Like all methods to estimate trabecular outflow facility, the fluorophotometric method rests on a number of assumptions. It is assumed that uveoscleral outflow varies little with changes in the IOP, and uveoscleral outflow facility is very small relative to trabecular outflow facility. This has been measured directly in animals and appears to be true under normal circumstances (Bill, 1966, 1967; Toris and Pederson, 1985). The fluorophotometry method would not be valid in some conditions that greatly aVect uveoscleral outflow facility such as the presence of a cyclodialysis cleft (Suguro et al., 1985). Additionally, there is a debate over the eVect, if any, of topical prostaglandins on uveoscleral outflow facility (Camras, 2003). The fluorophotometric method to assess outflow facility has been used in rabbits, cats, dogs, monkeys (Tables III–VI), and humans (See Chapter 8).

3. Perfusion Methods

The two level constant pressure perfusion technique (Ba´ra´ny, 1964) is used in research animals to achieve a more accurate estimate of outflow facility than the indirect methods of tonography and fluorophotometry. This technique requires insertion of needles into the eye, and if great care is taken to avoid infection and minimize injury, multiple measurements can be made in the same animal. The needle is attached to a reservoir of mock aqueous humor and the IOP is set by the level of the reservoir above the eye. The rate of fluid flow into the anterior chamber (F1) that is needed to maintain a stable intraocular pressure (IOP1) is measured. The reservoir is raised to a new level, and the new intraocular pressure (IOP2) and fluid inflow (F2) are measured. Equation (7) is used to calculate outflow facility. A similar method, the two level constant flow technique, measures the pressures needed to maintain a constant fluid flow into the eye at each of two diVerent flow rates. Like tonography, this method includes ocular rigidity, pseudofacility, and uveoscleral outflow facility in the measurement.

Arguably the most accurate method to assess trabecular outflow facility is the ‘‘flow to blood’’ method. A radioactive isotope is infused into the anterior chamber and the radioactivity detected in the blood during a specified time interval is considered to have drained solely through the trabecular meshwork. Changes in outflow facility can be made by sampling blood at diVerent times and infusion pressures and using Eq. (7) to calculate trabecular outflow facility.

D. Uveoscleral Outflow

The aqueous humor that enters the ciliary muscle exits the eye by multiple routes (Fig. 3). It percolates through the supraciliary space and across the anterior sclera or into the suprachoroidal space and across the posterior sclera

3

Choroid

6

6

4

Choroidal blood vessel

FIGURE 3 Uveoscleral outflow pathways. Most of the aqueous humor that flows anteriorly through the pupil (1) will drain into the anterior chamber angle (2). If it does not traverse the trabecular meshwork, it will enter the uveoscleral outflow pathway. This pathway starts with the ciliary muscle. From there, fluid can flow in many directions, including: across the sclera (3), within the supraciliary and suprachoroidal spaces (4), through emissarial canals (5), into choroidal vessels (6) and vortex veins (not drawn), and into ciliary processes (7) where it is secreted again.

(uveoscleral outflow) (Bill, 1971). It flows through the emissarial canals around the vortex veins (McMaster and Macri, 1968; Green et al., 1977; Krohn and Bertelsen, 1997) or traverses uveal vessels and into vortex veins (uveovortex outflow) (Pederson et al., 1977). Additional evidence suggests that the fluid enters the ciliary processes from the ciliary muscle and is secreted back into the posterior chamber. Figure 4 is an unpublished micrograph of our study tracking intracameral fluorescein isothiocyanate dextran (1 10 4 M, 70,000 MW) into the ciliary body of ketamine/xylazine anesthetized New Zealand white rabbits. The tracer was infused into the anterior chamber for 30 min at a pressure of 15 mmHg. The eye was quickly frozen in isopentane

FIGURE 4 Tracer filled ciliary processes of a rabbit. Fluorescence micrograph of two ciliary processes (arrows) of a rabbit infused into the anterior chamber with fluorescein isothiocyanate dextran (1 10–4 M, 70,000 MW) for 30 min at a pressure of 15 mmHg. The core of the left process is completely filled with tracer (light color) and the core of the right process is partially filled, leaving the tip void of tracer. This suggests that the direction of fluid movement is from the ciliary muscle into the process.

cooled in liquid nitrogen and slowly freeze dried over a period of 3 weeks to minimize postmortem tracer movement (Grayson and Laties, 1971). Cryosections were observed under a fluorescence microscope. Among other uveal tissues, the tracer was found in the core of ciliary processes. Fluid in the core is the source of material for aqueous humor. Therefore, some fluid originating from the anterior chamber is very likely secreted back into the posterior chamber.

A decrease in uveoscleral outflow has been found as a function of age in humans (Toris et al., 1999, 2002) and rhesus monkeys (Gabelt et al., 2003). Normal healthy monkeys 3–10 years of age drain 45–70% of their aqueous humor through the uveoscleral outflow pathway, whereas older animals 25–29 years of age drain aqueous humor at about half this rate (Gabelt et al., 2003). Uveoscleral outflow amounts to more than 80% of total outflow in mice, 1–20% in rabbits, 7–25% in cats, and 15% in dogs (Tables I and III–V).

1. Mathematical Calculation

The only noninvasive method to measure uveoscleral outflow requires assessment of the IOP, aqueous flow, outflow facility, and episcleral venous pressure in the same eye on the same day and calculating uveoscleral

outflow from Eq. (4). Large standard deviations are generated using this method because each variable has its own inherent variability. This approach is more useful to detect within and between group diVerences than it is to determine absolute values. Despite this limitation, the calculation method has been used to provide valuable information about drug eYcacy and disease eVects.

2. Intracameral Tracer Methods

Intracameral tracer methods are often used in research animals to determine uveoscleral outflow. The direct tracer method requires insertion of two needles into the eye: one connected by tubing to a pressure transducer to monitor the IOP and the other to a reservoir filled with a fluorescent or radioactive tracer. The IOP is maintained at a stable level by adjusting the height of the reservoir above the eye. At a precise time interval, usually 30–60 min, the eye is enucleated and dissected into tissues of the outflow pathway including iris, sclera, ciliary body, and choroid. The tracer is extracted from each tissue and quantitated. Uveoscleral outflow is calculated as the total volume of tracer that had accumulated in the tissue during the specified time interval. The sacrifice of the animal makes this method nonrepeatable.

The direct tracer method also has been used to measure the facility of uveoscleral outflow. Uveoscleral outflow is determined twice (F1, F2), once at each of two diVerent infusion pressures (IOP1, IOP2), necessarily in diVerent eyes, and uveoscleral outflow facility is calculated from Eq. (7).

The indirect isotope method involves infusing a radioactive tracer into the anterior chamber and monitoring the appearance rate of the tracer in the blood (an indication of trabecular outflow) and the disappearance rate of tracer from the anterior chamber (an indication of aqueous flow). Uveoscleral outflow is calculated as the diVerence between aqueous flow and trabecular outflow. This method does not involve sacrifice of the animal, and changes in uveoscleral outflow can be assessed over time.

E. Episcleral Venous Pressure

The aqueous humor that reaches Schlemm’s canal leaves the eye through collector channels and episcleral veins. The pressure in these veins averages about 7–11 mmHg in humans (Zeimer, 1989) and 10 mmHg in monkeys (Table VI). Normal values in mice, rabbits, cats, and dogs (8–14 mmHg, Tables I and III–V) are remarkably similar to primates despite the anatomical diVerences of the venous pathways among the various species.