Limbus

Surgical entry into the anterior chamber usually occurs through the peripheral cornea, limbus, and at the limbal–scleral junction (Fig. 9.5). Most approaches for anterior uvea and glaucoma surgeries enter the anterior chamber at the limbus (‘blue zone’), but may be accompanied by limited hemorrhage. Peripheral corneal incisions are usually used for cataract and lens removals, are performed faster, and preferred by most veterinary ophthalmologists. The limbus–scleral incision is not usually performed because of the resultant hemorrhage but offers the possibility of the least postoperative corneal astigmatism.

Fig. 9.5 Relationships of the iris and ciliary body to the other structures of the eye.

The limbus (‘blue zone’) is the 0.5–1.0 mm junction of the clear cornea and opaque white anterior sclera (Fig. 9.6). The limbus may contain some pigmentation (especially laterally), and a few small blood vessels. On its anterior surface the non-keratinized squamous epithelium of the cornea begins to transform to keratinized squamous epithelium of the bulbar conjunctiva. The limbus prevents direct observation of the anterior chamber angle and the aqueous humor outflow passages into the ciliary or sclerociliary cleft. In this zone regular corneal stroma lamellae begin to form the irregular dense connective tissues of the sclera. Blood vessels, absent in the normal cornea, are present in the anterior scleral tissues next to the limbus.

Fig. 9.6 The surgical limbus is the area between the external transition of the corneal to conjunctival epithelium (A) and the internal termination of Descemet’s membrane, anterior insertions of the pectinate ligaments, and anterior border of the corneoscleral trabeculae (B). H & E, 25×.

The outer anterior boundary of the limbus is the transition of the corneal epithelia into conjunctival epithelia, and the inner posterior boundary is the termination of Descemet’s membrane, insertion of the primary pectinate ligaments, and the corneoscleral trabecular meshwork. Hence, limbal incision and entry of the anterior chamber is just caudal of the termination of Descemet’s membrane and anterior to the insertion of the pectinate ligaments and the anterior chamber angle.

Iris

The most striking gross difference between the cat and dog irides, and the horse and cow irides, is the shape of the pupil. The round pupil of the dog, when the anterior uveal tissues are severely inflamed, becomes very small and subject to temporary or permanent occlusion. In contrast, the vertical slit pupil of the domestic cat, and the horizontal oval pupil of the horse and cow with its considerable pupillary margin length, are less likely to occlude secondary to iridocyclitis. Nevertheless, iris bombé glaucoma from pupillary occlusion may occur in these three species, but are less common.

Iridal coloration also varies between species, and among the different breeds of these species. The central portion of the iris with a normal size pupil usually touches the axial portion of the anterior lens capsule. The central iris is often most pigmented and subject to marked changes in thickness. Iridal color changes occur in these species in response to chronic inflammations and intraocular neoplasms with progressive pigmentation.

The animal iris is comprised of highly vascular, friable, and spongy tissues and, in contrast to humans, will usually hemorrhage when incised by a sharp scalpel or scissors. Dorsal and ventral branches from the medial and lateral long posterior ciliary arteries and veins enter the basal iris or anterior ciliary body to form an incomplete vascular circle providing the majority of the blood supply to the anterior uvea (Fig. 9.7). In dogs, this incomplete vascular annular circle occupies the basal iris in about 50% of animals; in the remaining animals the vascular circle is positioned in the anterior portion of the ciliary body. As a result, hemorrhage occurs about one-half of the time when the basal iris is incised. Radial arteriolar branches from this circle terminate in capillary beds in the animal pupil. The minor arteriolar circle of the iris, observed in humans, is lacking or incomplete in many animal species. Incision of the basal iris of animals is expected to hemorrhage and may require cautery for hemostasis, which, if possible, should be performed with the peripheral iris protracted from the anterior chamber. The inflamed animal iris can markedly thicken; this increased thickness is a significant deterrent to surgical- and laser-produced iridotomies, and unfortunately these small holes will usually seal and close within a few days.

Fig. 9.7 The base of the iris in both dogs and cats often contains large blood vessels that restrict the benefits of basal iridectomies, and can hemorrhage profusely when transected. H & E, 100×.

The animal iris is remarkably similar microscopically among the mammalian animal species, with often the main difference being the size and shape of the pupil. The iris separates the anterior and posterior chamber, with the former considerably larger. Aqueous humor, produced primarily by ciliary body processes, flows from the posterior chamber, through the pupil to enter the anterior chamber, and eventually exits the conventional and uveoscleral outflow passages. The different layers of the iris (from anterior to posterior) include: 1) the anterior border; 2) stroma and iridal sphincter musculature; and 3) the posterior epithelial layers including the iridal dilator muscles (Fig. 9.8). The anterior border of the iris, consisting of fibroblasts and melanocytes, has direct contact with the aqueous humor, and forms a more-or-less continuous cellular surface. The iris stroma consists of fine collagenous fibers, fibroblasts and melanocytes, and numerous blood vessels. The considerable extracellular spaces accommodate the physical changes in iris size secondary to the variations in pupil size.

Fig. 9.8 The iridal sphincter muscle is located at the pupillary margin and is the dominant muscle affecting pupil size in mammals. H & E, 100×.

The iridal sphincter, a non-striated muscle in the dog, cat, horse, and cow, and a striated muscle in birds, is located in the stroma near the pupil margin. In mammals, the iridal sphincter muscle is innervated by ciliary parasympathetic fibers from the ciliary ganglion within the orbit, but sympathetic innervation has also been demonstrated. The posterior layer of the iris is formed by the dilator muscles and the posterior iridal pigmented epithelium. The iridal dilator muscles, arranged like the spokes of a wheel, are highly developed pigmented myoepithelial cells, innervated primarily by the sympathetic fibers from the cranial cervical ganglion in mammals. The single layer posterior iridal epithelium is heavily pigmented and has direct contact with the posterior chamber and anterior surface of the lens.

In the avian species, pupil size and motility are under voluntary control, and light-induced testing of pupil reflexes of very limited value. If one undertakes entry into the anterior chamber and even cataract surgery in the avian species, the commercially available mydriatics have no effect. Often systemic ketamine is part of the avian general anesthesia protocol and can maintain a dilated pupil during anterior chamber, iris, and lens surgeries. Alternatively, intracameral use of non-depolarizing neuromuscular blocking agents has been described for mydriasis during cataract surgery in birds.

The corpora nigra (also known as granula iridica) are numerous black nodules or masses on the upper pupillary border and fewer small nodules on the ventral pupillary border of the iris in the horse and cow, and are especially well developed in South American camelids or cria (llamas, alpaca, etc.). Their function seems to assist the horizontal oval pupil in limiting light entry through the lens and vitreous, and to the retina. Fortunately, the corpora nigra are relatively avascular. They may become cystic, avulse secondary to trauma or undergo atrophy, or form synechiae to either the anterior lens capsule or posterior cornea secondary to chronic inflammation.

Ciliary body

The ciliary body is the second component of the anterior uvea, and continues posteriorly with the choroid. The ciliary body has contact with both anterior and posterior chambers, the sclera externally, the lens and vitreous internally, and the retina and choroid posteriorly (Fig. 9.9). The ciliary body is the principal source of aqueous humor, controls accommodation, and is important in the control of intraocular pressure (IOP). Aqueous humor is produced by active secretion by the non-pigmented ciliary body epithelium and by ultrafiltration from the capillaries in the ciliary body processes. Aqueous humor provides nutrients and removes waste products for the lens, anterior vitreous, iris, and posterior cornea. Aqueous humor, once leaving the ciliary body processes, enters the posterior chamber, traverses the pupil, and exits the anterior chamber through the iridocorneal angle and trabecular meshwork including the cilioscleral cleft or sinus within the anterior aspect of the ciliary body, or posteriorly through the uveoscleral route. Hence, the aqueous humor dynamics of the ciliary body are directly related to maintenance of IOP, essential for most of the intraocular tissues’ health and functions. The ciliary body, through its musculature, changes the tension on the zonulary attachments to the lens equator, and effects accommodation. Accommodation, or changes in the anterior–posterior length of the lens, appears minor in most mammals, and is seen mainly in young animals. Accommodation and the ciliary body musculature in non-human primates, some avian species, and humans are highly developed.

Fig. 9.9 The anterior uvea consists of the iris and ciliary body. Base of the iris (A), iridal pupillary margins (B), and ciliary body: pars plicata ciliaris (C) and pars plana ciliaris (D). SEM, 25×.

(Courtesy of Dr Don Samuelson, University of Florida.)

The ciliary body is divided grossly into: 1) anterior pars plicata (corona ciliaris or ciliary processes), and 2) posterior pars plana ciliaris. The anterior pars plicata consists of about 70 major and minor ciliary processes, the primary source for aqueous humor formation and the attachment of the zonules. The posterior flat pars plana ciliaris extends from the ciliary processes to the periphery of the retina, and serves as the termination for some of the lenticular zonules. Each quadrant of the pars plana ciliaris varies markedly in thickness by fractions of millimeters, and these differences are critical when entry into the vitreous is attempted as during retinal detachment surgery in the dog, cat, and horse.

The pars plana ciliaris varies in width, with the ventral and medial aspects the shortest in the dog and cat. In dogs, the posterior boundary of the pars plana ciliaris is 8 mm from the limbus dorsally and laterally, but only 4 mm behind the limbus ventrally and medially. Hence, entry into the anterior vitreous space through the pars plana ciliaris is usually through the larger dorsal and lateral quadrants.

In horses, the width of the ciliary body and, in particular, the pars plana ciliaris also varies by quadrant. These pars plana ciliaris widths range from 2.92 mm dorsally, 3 mm temporally or laterally, 0.33 mm nasally or medially, and 2.33 mm ventrally in fresh enucleated globes. Measurements by Frühauf et al for 12 o’clock vitrectomy suggest that the width of the dorsal equine pars plana ciliaris ranges from 7 to 12.06 mm posterior to the limbus, with an average of 5.06 ± 0.58 mm. Fortunately, both exposure of the eye and the dorsal quadrant of the pars plana ciliaris (about 3 mm wide) accommodate the single port vitrectomy in the horse. These measurements may also be increased when the equine eye is enlarged from glaucoma.

Microscopically the ciliary processes consist of a core of stroma rich in blood vessels, and two layers of epithelium (Fig. 9.10). The inner non-pigmented ciliary epithelium’s primary function is the formation of aqueous humor. It continues posteriorly as the inner neurosensory retina. The non-pigmented ciliary epithelia are linked to each other with tight gap junctions that represent ultrastructurally the blood–aqueous barrier. The deeper pigmented ciliary epithelium provides the majority of the pigmentation of the ciliary processes. The core of connective tissue and blood vessels within the ciliary process supply the energy needs of the two layers of epithelia, and the ultrafiltration portion of the aqueous humor. The more external aspect of the ciliary body consists of smooth muscles. With limited accommodation, the primary ciliary body musculature in the cat and dog is meridionally arranged and extends to the iridocorneal angle, forming the collagen beams for the trabecular meshwork. These ciliary muscles are richly innervated with parasympathetic nerve endings. Ciliary blood vessels are surrounded with numerous sympathetic nerve endings.

Fig. 9.10 The ciliary body’s main function in dogs and cats involves the formation of aqueous humor. The ciliary body process consists of a layer of non-pigmented and pigmented epithelia, and a central core rich in blood vessels. H & E, 100×.

Iridocorneal angle

The anatomy of the iridocorneal angle is presented in Chapter 10. The pathogenesis of most animal primary and secondary glaucomas appears to result from diseases of the outflow pathways, with the subsequent increase in resistance to aqueous humor outflow, and increase in IOP.

Keratocentesis/anterior chamber paracentesis

In keratocentesis a small gauge (25–30) hypodermic needle is inserted into the peripheral clear cornea or limbus to enter the anterior chamber and aspirate a small amount (0.1–0.2 mL) of aqueous humor. Alternatively, this technique may also be used for intracameral injection of materials such as tPA, adrenaline (epinephrine) or antibiotics. The indications for keratocentesis are summarized in Box 9.2. The aqueous humor sample has a limited volume, usually 0.1–0.3 mL. The value of each diagnostic procedure (cytology, culture, protein analyses, antibody titers) may require prioritization and only the most important tests performed (Fig. 9.11).

Fig. 9.11 Anterior uveitis and hypopyon in a shorthair cat with infectious peritonitis. Note the hypopyon also contains limited hemorrhage. The anterior uveitis affects both eyes.

For keratocentesis, short-acting general anesthesia or deep sedation is used to provide restraint. In the horse, general anesthesia or use of a retrobulbar nerve block in conjunction with sedation is required. Topical anesthetic is instilled on the cornea and the eyelids retracted by speculum. Exudates, if present, are removed by sterile cotton swabs and the corneoconjunctival surfaces are flushed liberally with 0.5% povidone–iodine solution.

For keratocentesis, a 25–30 g hypodermic needle and 1 mL syringe are used (Fig. 9.12a). A thumb forceps is used to grasp the bulbar conjunctiva and stabilize the eye. The hypodermic needle is directed through the peripheral cornea into the anterior chamber at an angle that avoids contact with the posterior cornea and the anterior iris (Fig. 9.12b). A small volume (0.1–0.3 mL) of aqueous humor is withdrawn. The needle bevel may be positioned down (or toward the iris) to avoiding snagging the iris and to provide a self-sealing needle puncture.

Fig. 9.12 Keratocentesis through the peripheral cornea. (a) The von Graefe thumb forceps is used to stabilize the eye and the hypodermic needle is inserted through the peripheral cornea. (b) The angle of the needle pathway must avoid the anterior iris and posterior cornea.

The hypodermic needle is carefully retracted. The needle track should be self-sealing; if a small amount of aqueous humor escapes, the needle hole is covered briefly with a sterile cotton swab. Depending on the amount of aqueous humor aspirated, IOP will be decreased proportionally.

As a variation of the clear corneal method, the hypodermic needle is inserted in the bulbar conjunctiva 2–4 mm posterior to the limbus (Fig. 9.13a). It is then carefully moved forward subconjunctivally to the limbus, and then inserted through the limbus at an angle between the posterior cornea and anterior iris (Fig. 9.13b). After aqueous humor sampling, the hypodermic needle is carefully and slowly retracted. If some aqueous humor leaks from the limbal penetration, it remains trapped in the bulbar subconjunctival space.

Fig. 9.13 Keratocentesis through the limbus and under the bulbar conjunctiva. (a) With the von Graefe thumb forceps holding the bulbar conjunctiva and globe, the hypodermic needle is inserted in the bulbar conjunctiva about 2–4 mm posterior of the limbus. The needle is advanced subconjunctivally to traverse the limbus. (b) The hypodermic needle angle is between the anterior iris and posterior cornea.

An alternative keratocentesis technique uses two 1 mL syringes connected by a two-way stopcock, and a single 25–30 g hypodermic needle. One syringe is used to aspirate 0.1–0.3 mL of aqueous humor; the other is filled with 0.5 mL sterile lactated Ringer’s or balanced salt solution (BSS), and is used to refill the anterior chamber with a volume equal to the aqueous humor removed (Fig. 9.14). This technique immediately replaces the lost aqueous humor, reduces the influx of the plasmoid or secondary aqueous humor, avoids any ocular hypotony, and requires only a single needle penetration into the anterior chamber.

Fig. 9.14 Use of two syringes, stopcock, and a single hypodermic needle permits keratocentesis and restoration of the lost aqueous humor as a single procedure.

When injecting materials intracamerally, a tuberculin syringe and 30 g needle are used. The volume to be administered is generally about 0.1 mL in small animals and up to 0.3 mL in large animals, and usually requires removal of an equal volume of aqueous humor before injection of the agent. The total volume to be administered should be the total volume in the syringe. The anterior chamber is entered in the same manner as for keratocentesis, the material injected, and the syringe removed as for keratocentesis. Leakage of a small volume of aqueous humor is acceptable as this will help maintain a more normal IOP since the anterior chamber has been volume-expanded.

Post-keratocentesis treatment includes topical mydriatics, antibiotics, and corticosteroids to control the resultant mild iridocyclitis. As any pre-existing iridocyclitis is usually intensified by keratocentesis, the benefits and risks of keratocentesis should be ascertained.

Complications after keratocentesis are infrequent. Keratocentesis is used experimentally to break down the blood–aqueous barrier, and markedly elevate serum proteins in the resultant (secondary or plasmoid) aqueous humor. These effects seem directly related to the release of prostaglandins from the anterior uvea.

After keratocentesis, the anterior chamber is quickly reformed with new (secondary) aqueous humor rich in proteins (plasmoid aqueous). This secondary aqueous humor frequently contains fibrin clots. In glaucomatous eyes, keratocentesis acutely decreases IOP, but may worsen iridocorneal angle closure. A misdirected hypodermic needle can penetrate the posterior cornea, creating a temporary corneal opacity secondary to penetration of Descemet’s membrane. If the hypodermic needle touches the iris, limited hemorrhage may result. If iridal hemorrhage occurs, the secondary aqueous humor contains additional levels of serum proteins.

The use of anterior chamber paracentesis in glaucomatous eyes has a potential benefit of lowering medically non-responsive elevated IOP to reduce damage to the optic nerve and retina versus the risks of causing further anterior chamber angle closure, intraocular hemorrhage, forward shifting of the lens and vitreous, and retinal and/or choroidal detachments. While keratocentesis temporarily lowers IOP in glaucomatous eyes, the resultant release of anterior uveal prostaglandins may elevate the IOP further within a few hours. Hence, keratocentesis is recommended to lower IOP only after intensive antiglaucoma medical therapy, including intravenous mannitol, has failed to lower IOP to safe levels, or immediately before an appropriate glaucoma surgery is performed. When used for acute glaucoma, a 30 g needle should be used without a syringe, allowing the needle hub to passively fill and the IOP to be gradually reduced. This will help to minimize complications such as an expulsive choroidal hemorrhage.

Corneal/limbal incisions

Surgical entry into the anterior chamber is the fundamental step in most intraocular procedures. For surgery involving the iris, ciliary body, certain glaucoma procedures, phacoemulsification, extracapsular and intracapsular lens extraction, proficiency in performing clear corneal or limbal incisions into the anterior chamber is essential. In the 1950s and 1960s, entry into the dog and cat anterior chamber was through the limbus under a limbal- or fornix-based bulbar conjunctival flap. Since the 1980s, the clear corneal incision has become popular in all animal species. Clear corneal incisions can be performed rapidly, avoid limbal blood vessels and any hemorrhage, and are easier to appose with simple interrupted or continuous sutures. However, without protective cover by the bulbar conjunctiva, exact apposition of the corneal incision is essential to maintain the integrity of the wound and IOP postoperatively.

The peripheral cornea and limbus may be incised perpendicular to the corneal surface, as a beveled incision to the corneal surface, or an incision that combines both perpendicular and beveled characteristics (Fig. 9.15). Those entries that include a beveled or angled entry into the anterior chamber are self-sealing. Entry into the anterior chamber that involves incisions that are perpendicular to the corneal surface are not self-sealing, but result in the least corneal scar formation. Limbal incisions are usually performed under a limbal- or fornix- based bulbar conjunctival flap. If a limbal-based conjunctival flap is used, the flap is usually only 3–5 mm wide to minimize its adverse effect on observation of the anterior chamber during iris and lens surgeries. Sutures for closure of the limbal wound may be buried under the limbal- or fornix- based conjunctival flap, or the knots may be exposed at the external limbus. Suturing of the limbal wound under a limbal-based conjunctival flap is more tedious as braided sutures often snag on the flap.

Fig. 9.15 The different types of corneal and limbal incisions. (a) Perpendicular to the corneal surface. (b) Beveled incision. (c) Combination perpendicular–beveled, most popular.

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