Chapter 9 Surgical procedures of the anterior chamber and anterior uvea
Diseases of the anterior chamber and anterior uvea (iris and ciliary body) are frequent in small and large animals. Causes of these diseases include trauma, congenital anomalies, inflammations, neoplasms, immune diseases, and degenerations. Aqueous humor within the posterior chamber, pupil, and anterior chamber delivers nutrients and removes wastes from the lens and posterior cornea, and contains cells, proteins, and other substances that may be useful for diagnostic and therapeutic techniques. Trauma affecting the anterior chamber and anterior uvea is common in small animals and horses, often resulting in hemorrhage within the anterior chamber (hyphema). Ocular trauma also includes foreign bodies that may be embedded within the cornea, those partially extending into the anterior chamber, or those that penetrate the globe and lodge in the orbit or posterior orbit.
Inflammations involving the anterior chamber and anterior uvea are common in older small animals, and may be isolated to one eye or affect both eyes with systemic diseases. Immune-mediate inflammation, termed equine recurrent uveitis (ERU), is the most common cause of blindness in the horse. An example of immune-mediated panuveitis in dogs is uveodermatologic or Vogt–Koyanagi–Harada syndrome. Both of these chronic uveitides are apt to cause secondary cataracts with posterior synechiae, secondary glaucoma, retinal degeneration, and optic nerve atrophy.
Leukocytic infiltration of the anterior uvea also affects the anterior chamber, causing the aqueous humor to become cloudy (‘aqueous flare’ – Tyndall effect), with gravitation of cells in the ventral anterior chamber producing hypopyon and the adherence of inflammatory cells on the posterior cornea (keratic precipitates). Anterior uveal inflammations in cats are usually associated with systemic diseases, and frequently require additional diagnostic procedures to establish the diagnosis (Fig. 9.1). Anterior chamber paracentesis (keratocentesis) with cytologic and protein (globulin, albumin, antibodies, etc.) analysis may assist in establishing the diagnosis.
Fig. 9.1 Anterior uveitis and hypopyon in a cat. Note the presence of clotted blood distributed throughout the hypopyon. As this condition is bilateral, cytologic examination of these cells may help establish the cause of the iridocyclitis.
Congenital anomalies of the anterior uvea are less frequent in small animals, but occur in purebred dogs with some frequency. Microphthalmia results in microcornea and micro-anterior chamber. Anterior uveal anomalies, such as persistent pupillary membranes, are not usually candidates for surgery, but may be associated with secondary hemorrhage that requires therapy. Congenital abnormalities of the anterior segment have also been described in the horse; specifically, anterior segment dysgenesis, megaloglobus, and other ocular abnormalities seen in the Rocky Mountain and similar breed horses.
Anterior uveal neoplasms are frequent in small animals, but less frequent in horses. Clinical differences in the malignancy of these tumors in the dog and cat necessitate different strategies for their clinical management. Anterior uveal neoplasms in the dog are primarily malignant melanomas and, less frequently, ciliary body adenomas and adenocarcinomas (Fig. 9.2). These neoplasms, affecting the iris, ciliary body or both structures, usually enlarge slowly and metastasize late and infrequently (about 5%). The clinical history of these neoplasms usually includes local irritation, ocular inflammation, and enlargement of the globe. Space-occupying masses are commonly associated with secondary glaucoma and retinal detachments. Hyphema or hemorrhage in the anterior chamber may result from the rapidly growing tumor, secondary glaucoma and lens luxation, and retinal detachments. Types of neoplasm affecting the anterior uvea of large animals are similar, with melanoma the most common primary tumor.
Fig. 9.2 Primary anterior uveal masses in the dog. (a) Sometimes the animal is presented because the owner notices a mass or something within the eye or pupil. (b) Other frequent presenting clinical signs include hyphema, anterior uveitis, and secondary glaucoma.
Small isolated iridal neoplasms in dogs and horses may be treated by iridectomy or iridocyclectomy combined with excision of the adjacent sclera, and more recently by diode laser photocoagulation (Fig. 9.3a,b). Diffuse iridal melanomas in cats are managed differently, because the potential for local infiltration and metastasis is greater. Cats with diffuse iridal melanomas are usually presented with a progressive brown to black pigmentation of the iris (Fig. 9.3c). The iridal mass increases in thickness late in the disease. Pupillary changes, secondary glaucoma, hyphema, and retinal detachments indicate that the iridal neoplasm is advanced, and an enucleation should be performed. Controversy exists among veterinary ophthalmologists and veterinary pathologists as to early clinical management of these neoplasms when the only clinical sign is the iridal pigmentation, which is progressing slowly. Diffuse iridal melanomas in cats usually involve the majority of the iris and are not amenable to sectional iridectomy or iridocyclectomy.
Fig. 9.3 Iridal melanomas may present in different ways in the dog and cat. (a) The single ventrolateral pigmented mass in this dog was noted by the owner because of the contrast to the surrounding light brown iridal tissues. (b) In contrast, this owner noted multiple pigmented masses in their dog’s iris. Multiple masses complicate the therapy options. (c) In this cat, the entire iris has become densely pigmented and the pupil irregular.
Cysts of the iris and ciliary body are not infrequent in older dogs, cats, and horses (including those of the granula iridica). Iris cysts arise from the posterior iridal epithelium, and appear as densely pigmented, single or multiple spherical bodies. In dogs, they may be free-floating within the anterior chamber, in the pupil, or still attached to the posterior iridal surface (Fig. 9.4). In cats, most iridal cysts remain attached to the pupillary margin. In dogs presented with iridal cysts in the anterior chamber, the drug-induced mydriasis associated with the ophthalmic examination may liberate additional cysts into the anterior chamber that were trapped behind the pupil in the posterior chamber. The free-floating cysts gravitate to the most ventral portion of the anterior chamber. Manipulation of the animal’s head and ocular movements will often cause these bodies to move within the anterior chamber. Sometimes these iridal cysts become trapped in the anterior chamber angle, and may simulate a basal iridal melanoma. These small round black iridal cysts transilluminate, distinguishing them from anterior uveal melanomas. B-scan ultrasonography will also reveal hollow centers. Treatment includes temporization, laser-induced rupture or deflation, and paracentesis or lavage from the anterior chamber.
Fig. 9.4 Pigmented iris (usually black) or ciliary (usually red or clear) cysts in the dog may have different clinical significance. (a) This single iris pigmented cyst in a 12-year-old Golden Retriever can be deflated by laser therapy or aspiration. The cyst readily transilluminates and the prognosis is good. (b) In contrast, these ciliary cysts (clear cysts behind the pupil) and two pigmented iridal cysts (one dorsal and one in the ventral anterior chamber) in a 3-year-old Golden Retriever often progress to uveitis, pigment deposition on the anterior lens capsule, secondary cataract and glaucoma formation, and warrant a much more guarded prognosis.
Unfortunately, not all anterior uveal cysts appear innocuous. Anterior uveal cysts have been associated with glaucoma in the Golden Retriever and Great Dane breeds. In the Golden Retriever, nearly one-half of the dogs develop glaucoma and most of these animals lose their vision. The clinical significance of iridal and ciliary body cysts is not known; histologically, the ciliary body cysts contain periodic acid–Schiff (PAS)-positive material which may be deleterious to aqueous outflow pathways. The high number of these cysts within the posterior chamber may also displace the basal iris forward and compress the opening of the sclerociliary cleft. Laser deflation of these cysts seems not to delay the onset of the glaucoma, but more study is indicated.
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.
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.
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.
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.
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.
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.
Surgical entry of the anterior chamber elicits an acute inflammatory response by the anterior uvea (iris and ciliary body) which is mediated by the release of prostaglandins. Similarly, the surgical approach to the iris and ciliary body usually involves entry into the anterior chamber. The highly vascular sclera presents a significant barrier for entry to the iris, ciliary body, and posterior segment (including the vitreous space and retina). Cautery of the sclera is often necessary to control hemorrhage. Because penetration of the retina may result in retinal detachment, access for vitreous and retinal surgeries is limited to the pupillary approach, or through incisions in the pars plana ciliaris.
The pathophysiology of the anterior uvea during intraocular surgery includes several significant considerations. Both the iris and ciliary body are highly vascular. Tearing with forceps or incisions with scissors or scalpel result in variable hemorrhage. Control of the iridal and ciliary body hemorrhage requires strategies that can be effective in the presence of aqueous humor and blood. Judicious wet-field cautery, diode laser, intracameral adrenaline (epinephrine) injections, and viscotamponade are the usual methods to control anterior uveal hemorrhage. For large bleeders of the iris, wet-field cautery units are superior.
Direct surgical trauma or entry into the anterior chamber in the dog and cat results in variable miosis. In some breeds, such as the Miniature Schnauzer, the miosis may result in a pupil of only 1–2 mm (sometimes referred to as atropine- or mydriatic-resistant miosis). Pupillary constriction may limit the visualization of the lens or posterior segment, and impede, delay or preclude the surgery. This miosis, previously thought to be associated with the release of histamine or other substances, is now believed to be secondary to the release of uveal endogenous prostaglandins. The prostaglandins elicit an immediate contraction by the iridal sphincter muscles, breakdown of the blood–aqueous barrier at the level of the ciliary body epithelium with the resultant plasmoid or secondary aqueous humor, an acute elevation in IOP with subsequent decrease in IOP (ocular hypotony), and hyperemia of the anterior uveal and bulbar conjunctival blood vessels. Ocular hypotony from anterior uveal inflammation and release of prostaglandins is associated with an increase in the unconventional or uveoscleral aqueous humor outflow through the posterior ciliary body.
Administration of non-steroidal anti-inflammatory drugs (NSAIDs) prior to intraocular surgery, and prevention of the release of prostaglandins during anterior chamber entry, may directly affect the eventual success or failure of the surgery. Pre-, peri-, and postoperative administrations of topical antiprostaglandins and indometacin, as well as systemic carprofen (Rimadyl® Pfizer Animal Health, Exton, PA) and flunixin meglumine (Banamine® Schering-Plough, Kenilworth, NJ), are used to suppress the release of intraocular endogenous prostaglandins. As a result, intra- and postoperative control of the pupil in animals with topical mydriatics has become more successful.
Simple entry into the anterior chamber to aspirate aqueous humor (anterior chamber paracentesis or keratocentesis), with the resultant decrease in IOP, immediately disrupts the blood–aqueous barrier. The secondary aqueous humor quickly restores the normal volume of the posterior and anterior chambers but contains high levels of albumin and globulins. These proteins result in increased turbidity of the aqueous humor and formation of frank fibrin clots within the anterior chamber. The fibrin clots may serve as scaffolds for attachments of the iris to the lens, posterior lens capsule, posterior cornea, and peripheral cornea. The fibrin clots can also be gradually replaced with fibroblasts, collagen, and pigment cells. The resultant opaque fibrous bands can permanently link the iris to the lens or its capsules, the posterior cornea or the edges of the pupillary margins, resulting in small and irregular pupils that impair vision.
Pre- and postoperative treatment with topical and systemic corticosteroids and NSAIDs, and intracameral heparin during surgery help to reduce the formation of intraocular fibrin. Introduction of tissue plasminogen activator (tPA), injected directly into the anterior chamber, can dissolve any existing aqueous humor fibrin and clotted blood within 1–2 weeks post-formation. Intracameral tPA cannot, however, prevent the formation of future aqueous humor fibrin. With any postoperative iridocyclitis, the iris becomes inflamed and thicker than normal. From its inflammatory products the iridal surface becomes sticky and readily adheres to any intraocular tissues it contacts. As a result, temporary to permanent attachments may develop involving the iris and lens (posterior synechiae), the iridocorneal angle (peripheral anterior synechiae), the posterior cornea (anterior synechiae), or its pupillary margins (pupillary synechiae). Permanent iridal contact with the anterior lens capsule results in anterior capsular and anterior cortical cataract formation. Permanent iridal contact with the posterior cornea results in edema and dense corneal scars. Formation of peripheral anterior synechiae within the iridocorneal angle predisposes the eye to angle-closure glaucoma by filling and closing the opening of the sclerociliary cleft. Adherence of the pupillary margins creates an irregular pupil and may completely occlude the pupil, resulting in an immediate iris bombé and loss of vision. Treatment strategies to prevent the formation of synechiae are summarized in Box 9.1.
• Restore the blood–aqueous barrier to decrease as much as possible the cellular and protein (fibrin) content of the secondary or plasmoid aqueous humor, and reduce the possibility of formation of fibropupillary membranes.
As iris and ciliary body inflammation always occurs following anterior chamber penetration and the resulting abrupt decrease in IOP, medical treatments should be implemented pre-, peri-, and postoperatively. If treatment is limited to the postoperative period, less than optimal results should be anticipated. The intensity of postoperative iridocyclitis is variable, and should be assessed daily. With these changes in the intensity of postoperative iridocyclitis, the types and frequencies of the topical and systemic medications should be adjusted accordingly.
The peak in the anterior uveal inflammation after intraocular surgery usually occurs 3–5 days postoperatively. An indirect method of assessing the intensity of the iridocyclitis is to measure IOP daily by applanation tonometry. After the initial elevation in IOP associated with iridocyclitis (which is often missed clinically because it lasts for only a few hours), daily tonometry, performed about the same time each day, can monitor the extent and duration of the ocular hypotony. As the iridocyclitis gradually resolves, IOP will return to normal levels. As a clinical guide, medical treatments should be administered until normal IOP returns. Normal IOP with intense iridocyclitis often signals the onset of secondary glaucoma and the obstruction of the aqueous filtration angle with inflammatory debris and the formation of peripheral anterior synechiae.
Preoperative treatment before entry into the anterior chamber or surgery of the iris and ciliary body involves: 1) dilatation of the pupil with a mydriatic (usually 1% atropine or 1% tropicamide) or a combination of mydriatics (1% atropine or 1% tropicamide plus 10% phenylephrine); 2) topical (usually 1% prednisolone or 0.1% dexamethasone) and systemic corticosteroids (e.g., prednisolone 0.5–1.0 mg/kg PO once or twice daily) to suppress preoperative or anticipated postoperative iridocyclitis; 3) topical and systemic NSAIDs and antiprostaglandins to reduce anterior uveal inflammation and assist in maintenance of the dilated pupil during surgery; and 4) topical and systemic antibiotics to prevent bacterial infection if contamination of the anterior chamber occurs during surgery.
Entry into the anterior chamber is usually through the periphery of the cornea or surgical limbus, or by a limbal–scleral incision. As the last approach usually causes hemorrhage from the posterior limbal and anterior scleral blood vessels, the clear corneal and limbal approaches are usually performed. Entry into the anterior chamber may be gained by a hypodermic needle, or scalpel blade (Beaver No. 6500 microsurgical blade or 55M (keratome)).
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).
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.
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.
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.