Isolation of Retinal Arterioles for Ex Vivo Cell Physiology Studies

All experiments described here were performed in accordance with guidelines contained within the UK Animals (Scientific Procedures) Act of 1986 and were approved by the Queen's University of Belfast Animal Welfare and Ethical Review Body.

1. Isolation of Retinal Arterioles

Note: The following protocol is used to isolate retinal arterioles. This method is optimized for rat retinal arterioles but can be used with mouse retinas. The yield of vessels, however, is lower when using mice.

1. Make up a solution of low Ca²⁺ containing Hanks' (LCH) as shown in Table 1.
   NOTE: This solution can be stored for up to 3 days at 4 °C (when using stored LCH, ensure that it equilibrates to room temperature and the pH is correct before use).
2. Assemble the following equipment prior to collection of tissue to ensure rapid microdissection of the retinas: serrated forceps, curved scissors, dissection dish (silicone-coated Petri dish), single edged blade, 2 sets of forceps (blunt) 1 glass Pasteur pipette (fire polished to a smooth but not narrow tip), 1 disposable plastic pipette (with tip trimmed off to increase the aperture to ~ 5 – 7 mm), 1 glass round bottom test tube (5 mL) and holder. Decant approximately 50 mL of LCH into a glass beaker and prepare a second beaker to decant waste fluids.
   NOTE: See Table of Materials for specifications and manufacturer details.
3. Euthanize the rat using CO₂ followed by cardiac puncture to exsanguinate (avoid cervical dislocation or concussion as these may damage blood vessels in the eye). Retract the eyelids with serrated forceps, then use the curved scissors to cut around the orbital muscles and through the optic nerve. Gently extract the eyes from the orbit taking care to cut through any remaining attachments with scissors.
4. Place the eyes immersed in LCH solution on the dissection dish (eyes can be transported on ice in a solution of LCH if necessary). Use one set of blunt forceps to anchor the eye to the dish by holding the orbital muscle attachments or optic nerve; ensure that the anchoring point is as close as possible to the sclera to stabilize the eye.
5. Using the blade, cut through the cornea along the ora serrata and remove the lens by pressing gently on the sclera with forceps.
6. Cut the eye cup in half symmetrically through the optic disc. Using closed forceps gently 'brush' out the retinas from the two halves of the eyecup taking care to remove any remaining attachments at the optic disc. Repeat the process with the second eye and transfer the dissected retinas into the test-tube using the plastic pipette and a small drop of LCH medium.
7. Using the plastic pipette fill the test tube with LCH to ~ 5 mL and allow the retinas to settle to the bottom.
8. Wash the tissue approximately 3 times by removing ~ 4 mL of solution from the tube and adding fresh LCH using the plastic pipette. Where necessary, remove extraneous tissue such as suspensory ligaments, vitreous humor and hairs.
9. Wash the inside of the glass Pasteur pipette (polished tip) with LCH to prevent the tissue from sticking to the pipette. Using the same pipette, remove approximately 4 mL of solution from the test-tube. Then add approximately 2 mL of fresh LCH.
10. Gently dissociate (triturate) the retinas by drawing the tissue through the tip of the glass pipette slowly and expel the contents into the test-tube. Note, try not to introduce bubbles at this stage.
11. Repeat step 1.10 until the retinas are broken up to a size of approximately 2 – 3 mm².
12. Wash the inside of the pipette with approximately 2 mL of LCH and expel this into the test-tube. Allow the contents to settle to the bottom over 5 – 10 min.
13. Repeat the trituration process as outlined in 1.10 – 1.12 with a little more force until the tissue pieces are approximately 1 mm² in size. Again, allow the tissue to settle for 5 – 10 min.
14. Repeat steps 1.10 – 1.12 a third time with more force until the contents are fully homogenized and no pieces of retina remain (the solution should become milky in appearance).
    NOTE: This technique will yield up to 8 arteriolare segments (relatively free of surrounding neuropile and with some branches remaining intact) per isolation measuring 8 – 45 µm in diameter and 200 – 2500 µm in length.
15. Transfer a small quantity of the isolate into a physiological recording chamber or add fluorescent dyes for measurement of intracellular Ca²⁺ concentration ([Ca²⁺]_i; see section 3). Dispose of remnants of eye cups and solution removed during the isolation process in accordance with local rules on handling of animal waste.
    NOTE: While it is advisable to experiment on vessels immediately after isolation, surplus isolate can be stored at 4 °C for up to 8 h.

2. Arteriolar Pressure Myography

NOTE: Arteriolar pressure myography is carried out as follows using equipment detailed in Figure 1A, B and the Table of Materials.

1. Transfer an aliquot of retinal vessel homogenate (1 – 2 mL; isolated as per section 1) to a physiological recording chamber (partially filled with nominally Ca²⁺free Hanks' solution; 0Ca²⁺ Hanks'; see Table 1) mounted on the stage of an inverted microscope. Leave the vessels to settle on the bottom of the recording chamber for at least 5 min prior to experimentation.
2. Visually scan across the recording chamber to identify arterioles > 200 µm in length and possessing an open end through which the vessel can be cannulated (identification of vessel type is further explored in the results section). Position the vessel in the center of the field of view. If no suitable vessels are present, transfer another aliquot of vessel homogenate into the recording chamber as per step 2.1.
3. Anchor one end of the vessel using fine forceps and a small tungsten wire slip (75 µm diameter and ~ 2 – 4 mm in length) placed over the vessel (see Figure 1C(i)). To do this hold the wire in the forceps and locate the forceps above the recording chamber.
   1. Identify the corresponding shadow of the forceps under the microscope, lower the forceps into the bath until the wire becomes apparent. Position the wire over the vessel approximately 200 µm from the open end and drop the wire perpendicular to the long axis of the vessel.
      NOTE: The weight of the tungsten wire is sufficient to occlude the vessel distally stopping flow of luminal contents during cannulation and pressurization.
4. Move the arteriole into a suitable position within the recording chamber such that it lies horizontally across the bath with the open end in line with the pressurization cannula (see Figure 1C(ii-iv)). Do this by gently nudging the occluding tungsten wire slip with an additional section of wire held within the forceps; do not touch the arteriole as it will irreversibly adhere to the forceps.
5. Where necessary (for long arteriole segments), add additional tungsten wire slips over the vessel such that the distance between the occluded and open ends of the vessel is no more than 200 µm; this helps to prevent the arteriole moving out of the field of view upon pressurization. If the vessel has any side branches, these should also be occluded with additional tungsten wire slips. If a side branch cannot be occluded discard the vessel.
6. Perfuse the chamber with 0Ca²⁺ Hanks' at 37 °C to remove extraneous retinal tissue and to warm the bath to physiological temperatures.
   NOTE: A standard gravity-fed bath perfusion and in-line heater system can be used for this purpose (see Figure 1A). 0Ca²⁺ Hanks' is used to facilitate the cannulation procedure, since removal of external Ca²⁺ ensures that the arterioles remain dilated.
7. Fabricate pressurization cannulas from filamented borosilicate glass capillaries (tip diameter ~3 – 10 µm depending on the diameter of the vessel to be cannulated; smaller vessels require narrower cannulas; see the Table of Materials) using a microelectrode puller and back-fill with 0Ca²⁺ Hanks' solution. Ensure the shank of the cannula is tapered gently towards the tip to facilitate cannulation.
8. Mount the cannula in a pipette holder attached to a 10 mL syringe via a Y-connector and a 3-way tap; this is used to pressurize the system, pressure being recorded using a manometer attached to the other side-arm of the Y-connector (see Figure 1B).
   NOTE: A 'helper' pipette is required to assist with the cannulation process. Fabricate the pipette from a borosilicate glass capillary pulled on a microelectrode puller to a tip diameter of 0.5 – 2 µm. Heavily fire polish the pipette (often until closed) using a microforge.
9. Carefully position the helper pipette at right angles to the long-axis of the vessel close to the open end using a 3-axis mechanical manipulator (see Figure 1D). Position the helper pipette just above the vessel, but not touching it.
10. Position the cannulation pipette at the open end of the vessel (see Figure 1D and 2A(i)) using a fine micromanipulator; position the tip immediately adjacent to the opening, as assessed by adjusting the plane of focus on the microscope (at 20X). When both the end of the vessel and the cannula tip are in focus at the same time the cannula is correctly positioned for cannulation (see Figure 2A(ii)).
11. Advance the pressurization cannula into the vessel aperture using the x-axis controls of the fine micromanipulator (see Figure 2A(iii)). Assess the success of cannulation by moving the cannula along the y-axis. If the cannula remains within the vessel it is successfully cannulated (see Figure 2A(iv)). If the cannula moves outside of the vessel, the cannula is likely to be above the vessel; if so repeat step 2.10 – 2.11 with the cannula at a lower plane in the z-axis.
12. Upon successful cannulation gently lower the helper pipette onto the vessel to restrain the arteriole. Guide the arteriolar wall over the pressurization cannula with the helper pipette, at the same time as the cannulation pipette is advanced further into vessel lumen to a distance of approximately 30 – 50 µm (see Figure 2A(v,vi)).
    NOTE: This procedure requires simultaneous, controlled movement of both manipulators (helper pipette moving towards the cannula while the cannula moves into the vessel lumen) and will require extensive practice to achieve a high success rate.
13. Change the bath solution to normal Hanks' containing 2 mM Ca²⁺ (see Table 1) at 37 °C. Typically this causes a slight narrowing of the arteriole and the formation of a tight seal around the pressurization cannula. The helper pipette can then be moved away from the vessel.
14. Allow a tight seal to develop for 15 min. View the vessel using a USB camera (and image acquisition software) attached to the microscope and adjust the focus to maximize the contrast between the vessel boundaries and the external and intraluminal spaces (see Figure 2B). Image the vessel at 0.5 – 5 frames/s.
15. After 1 min of recording at 0 mmHg, increase the intraluminal pressure to the desired level by closing the 3-way tap attached to the pressurization cannula (see Figure 1B) and apply a small amount of positive pressure to the system via the syringe. Observe the pressure change on the manometer.
    NOTE: Pressure can be added in a step wise fashion up to 100 mmHg or to one value for example 40 mmHg (approximating retinal arteriolar pressure in vivo)²³. The vessel should rapidly dilate confirming successful cannulation. Please note, pressure-induced vasoconstriction (i.e, the myogenic response) will subsequently develop over a period of 15 min, but because of the small size of these vessels, this may not always be visually detectable.
16. Carefully monitor the vessel during initial pressurization for obvious leaks. Sources of leak may originate from cleaved side branches or inadequate sealing of the cannula to the inside of the arteriole.
    1. Watch for intraluminal contents leaving the vessel. Smaller, less obvious leaks may become apparent over time as a drop in pressure on the manometer. If possible, occlude the opening with additional tungsten wire slips taking care not to disturb the cannula. Exclude preparations with obvious leakage from further analysis.
17. Apply the drug of interest abluminally to the vessel prior to, or following, 15 min pressurization depending on the aims of the experiment (the former allows effects on the development of the myogenic tone to be determined, while the latter enables analysis of effects on steady-state myogenic tone). A typical drug delivery system is pictured in Figure 3A.
    1. Position the delivery outlet perpendicular to the portion of vessel of interest (as illustrated in Figure 1D) at a distance ~ 500 µm away from the vessel of interest taking care to ensure that the outlet is optimally angled to fully superfuse the area (see Figure 3B). Ensure that changes in perfusion do not physically disturb the vessel.
18. Following completion of an experiment, apply a solution of 0Ca²⁺ Hanks' containing 10 µM wortmannin (myosin light chain kinase inhibitor) to maximally dilate the vessel to determine the passive diameter.
    NOTE: The strength of the cannulation seal can be tested at the conclusion of the experiment by raising the intraluminal pressure to >100 mmHg. The majority of vessels will remain attached even at this high pressure indicating a tight seal.
19. Perform offline analysis using edge detection to track changes in the arteriolar diameter (see Table of Materials for suggested software). Normalize the arteriole diameter to the passive diameter for comparison between vessels.

3. Ca²⁺ Imaging

Note: Isolated retinal arterioles are prepared for conventional (microfluorimetry) and confocal Ca²⁺ imaging as follows (using equipment detailed in Table of Materials).

1. Prepare retinal homogenates in a 5 mL glass test tube as described in section 1. To 1 mL of retinal homogenate add Fura-2AM (microfluormetric Ca²⁺ recording) or Fluo-4AM (confocal Ca²⁺ imaging) to give a final concentration of 5 µM (protect from light); dye indicators preferentially load into smooth muscle cells.
2. Maintain at room temperature in the dark for 2 h flicking the side of the tube every 15 – 20 min to re-suspend the retinal tissue. Thereafter, dilute the homogenate 1:4 with LCH solution to reduce further dye loading.
   NOTE: Vessels remain viable for up to 6 h (when stored at 4 °C and in the dark); however it is recommended that experiments are commenced as soon as possible to reduce the impact of compartmentalization of the dye in internal organelles or extrusion of the dye over time.
3. Transfer 1 – 2 mL of retinal homogenate to a glass-bottomed recording chamber (partially filled with LCH) mounted on the stage of an inverted microscope connected to a Ca²⁺ microfluorimetry or confocal microscopy system.
   1. Anchor arterioles perpendicular to the direction of flow across the recording chamber, with tungsten wire slips (50 µm diameter, 2 – 4 mm in length) spaced ~ 100 – 200 µm apart along the vessel length (see Figure 3C) using a similar technique to that described in step 2.3.
4. Perfuse the bath with normal Ca²⁺ Hanks' solution at 37 °C. Position the drug delivery outlet as pictured in Figure 3C and apply drugs to vessels as described in step 2.17.
5. For conventional Ca²⁺ imaging, illuminate the vessel with 340/380 nm light via an oil immersion UV objective (e.g., 100X, NA 1.3) and collect emitted fluorescence at 510 nm via a photon counting photomultiplier tube (PMT).
6. At the end of each experiment, measure background fluorescence by quenching the vessel with a MnCl-containing solution (see Table 1). Use background-corrected fluorescence ratios (R-F340/F380) for analysis or convert to intracellular Ca²⁺ ([Ca²⁺]_i) using R_min and R_max measurements (see Table 1; applied prior to quenching²⁴) and the Grynkiewicz equation²⁵.
7. For confocal Ca²⁺ imaging, excite Fluo-4 loaded vessels at 488 nm and capture the resultant fluorescence emissions at 490 – 535 nm in either line scan mode (xt) or xyt scan modes (512 x 512 pixels; suggested pinhole 300 nm). Normalize measurements to the fluorescence recorded at time t=0s (F/F₀). Assess any changes in [Ca²⁺]_i during drug applications or pressurization offline in specific regions of interest (ROIs) using image analysis software.
8. If needed, perform Ca²⁺ imaging with pressure myography.
   NOTE: The descriptions above have been optimized for unpressurized vessels; however it is possible to combine Ca²⁺ imaging with pressure myography as follows.
   1. Isolate arterioles as per section 1. Load with Ca²⁺ indicator dyes as per step 3.1 – 3.2. Transfer the arterioles to the recording chamber and cannulate as per section 2. Take care to limit exposure to light during the mounting procedure.
   2. Measure Ca²⁺ as per step 3.5 or 3.7 above. Pressurize the vessel and repeat the measurement of Ca²⁺_. Simultaneous measurement of vessel diameter is dependent on the mode of Ca²⁺ measurement and equipment used.
      NOTE: For confocal Ca²⁺ measurement it may be necessary to image separate regions pre- and post- pressurization due to bleaching of the dye during long exposures to light.

4. Patch-Clamp Electrophysiology

Note: Whole-cell and single-channel recording is possible from individual arteriolar smooth muscle cells still embedded within their parent arterioles as follows (using equipment detailed in Table of Materials).

1. Isolate and anchor retinal arterioles in the recording chamber as described in steps 1, 2.3, and 3.3. For patch-clamp recording, the tungsten wire slips are spaced 200 – 800 µm apart along the vessel.
2. Remove basal lamina surrounding the arteriole and electrically uncouple adjacent smooth muscle cells and underlying endothelial cells prior to patch clamping. To do this, superfuse the vessel with a sequential series of enzyme solutions (Table 1) at 37 °C for the required duration.
   NOTE: The duration of enzyme exposure is optimized for rat retinal arterioles (protease, ~ 8 min; collagenase, ~ 12 min) and depends on the flow rate of the drug delivery system (~1 mL/min on our set-up) and the unit activities of the batch of enzymes being used.
3. Evaluate the level of digestion on visual separation of endothelial and smooth muscle layers during the collagenase step as shown in Figure 4. Once separation of the cell layers is observed (as shown in Figure 4B), apply DNAse I solution (~ 4 min) then terminate the digestion by replacement of the bath solution with normal Ca²⁺ Hanks' solution.
4. Remove any remaining strands of basal lamina and/or peripheral neuropile by carefully sweeping the closed tips of fine forceps along the surface of the vessel.
5. Pull and polish glass capillaries (Table of Materials), fill with pipette solution (Table 1) and place securely in the pipette holder of the patch-clamp headstage. Position the pipette at a 45° angle in relation to the recording chamber and advance it towards the vessel using the coarse setting on a motorized micromanipulator.
6. Select a smooth muscle cell protruding from the vessel wall and whose surface is free from visible debris (see Figure 4B). Position the tip of the patch pipette vertically over the cell of interest and lower gradually using the fine/slow movement of the micromanipulator to make contact with the smooth muscle membrane.
   1. Assess the contact between cell and pipette visually from cell movement and a change in the pipette resistance measured using the membrane (seal) test protocol in the acquisition software (5-10 mV negative step from 0 mV, frequency 25 KHz; see Figure 5A(i)).
7. When the seal resistance has increased 5-fold (e.g., rising from 2 to 10 MΩ; Figure 5A(ii)) apply negative pressure transiently to the back of the pipette holder via a y-connector, 3-way tap, syringe and tubing attached to the pipette holder (assembled in a similar manner to that used in pressurization of arterioles, see Figure 1B).
8. Repeat applications of negative pressure until a gigaseal (>1 GΩ resistance; as indicated by the membrane test in the acquisition software) is formed (Figure 5A(iii)). Note, this process may take 1 – 5 min. If a gigaseal fails to form, choose a different cell to patch clamp using a fresh patch pipette.
9. Apply a holding potential to the cell via the acquisition software as per the experiment being performed (typically 0, -40 or -80 mV).
10. Study (on-cell) single channel activity in the present configuration. Alternatively, generate inside-out patches by rapidly retracting the patch pipette from the cell surface after gigaseal formation.
    1. As the free ends of the membrane can re-anneal under these conditions remove the pipette from the bath via rapid vertical movement of the manipulator (coarse setting). Rapidly return through the meniscus to remove the reformed membrane leaving only the patch inside the pipette tip.
11. Set the sampling frequency on the acquisition software to 5 KHz and activate continuous recording mode to record channel activity. Apply any voltages changes if required via the software or the amplifier.
12. If required, apply drugs to vessel/membrane patch as described in step 2.17.
13. If membrane stretch is required, apply negative pressure to the posterior of the pipette using the syringe attached to a manometer via a 3-way tap and y-connector.
14. Analyze single channel recordings for open probability and unitary conductance in accordance with standard procedures²⁶.
15. For whole cell current recording pull and polish pipettes as per the Table of Materials, fill with pipette solution containing amphotericin B (add 3 mg of amphotericin B to 50 µL of DMSO; add 3-6 µL of this to 1 mL of whole-cell pipette solution; sonicate to mix) and form a seal as per steps 4.6 – 4.9.
16. Due to the inclusion of amphotericin B there is no need to rupture the membrane within the pipette tip to gain whole cell access. Monitor access, which will be gained gradually, using the membrane test protocol in the acquisition software (-20 mV step from -40 mV, sample frequency 25 KHz; see Figure 5B).
17. Automated fitting of capacitive transients in some software yields real-time measurements of access resistance and cell capacitance (alternatively the relative decline of the transients can be assessed by eye). Once the access resistance falls to < 15 MΩ, perform series resistance compensations (Figure 5C) using the whole-cell parameter dials (capacitance and series resistance) on the amplifier.
    1. To do this, use the automated fitting values to initially set the dials (see Figure 5C(ii)) then increase the series resistance compensation dial (both prediction and correction if available on the amplifier) re-adjusting the whole cell compensations while reducing the capacitive transient (see Figure 5C(iii)).
    2. Taking care not to over compensate (as shown in Figure 5C(iv)). Typically, it is possible to compensate the series resistance by 75% in perforated patch mode (requires lag compensation to be set at maximum).
18. If no automated fitting is available, start by setting the whole-cell parameters dials to 15 pF and 15 mΩ (typical values for these cells) and then adjust to produce the outputs as shown in Figure 5C(ii). Increase the series resistance compensation dial to ~75% and readjust the whole-cell parameters dials as per Figure 5C(iii).
19. Prior to commencing experimentation note the cell capacitance measurement either from the initial automated fitting or from the dial after manual compensation.
    NOTE: This value is used to normalize currents to cell size. Compensation for series resistance may need to be adjusted during experimentation as access may continue to improve due to further incorporation of amphotericin B into the patch.
20. Apply drugs to vessels as described in step 2.17. Apply voltage protocols (either steps or ramps) as per experimental requirements.
    NOTE: Typical voltage step protocols, 0.1 – 1 s in duration, are applied from the holding potential in 10 – 20 mV increments every 5 – 10 s to produce a family of voltage-activated currents. Alternatively use ramp protocols to measure currents at a series of voltages in one 'sweep' by ramping the voltage slowly (100 – 200 mV/s) from e.g., -80 to +80 mV (applied every 5-10 s) with offline sampling of the current at 10 mV intervals during the ramp.
21. It is possible to combine Ca²⁺ imaging with patch-clamp recording as follows. Isolate arterioles as per section 1. Load with Ca²⁺ indicator dyes as per steps 3.1 – 3.2. Mount in the recording chamber as per section 4. Take care to limit exposure to light during these procedures.
    NOTE: To date it has not been possible to combine patch-clamp with pressure myography studies due to the loss of the basement membrane and vessel integrity during the enzymatic dissociation process