Establishment of a Severe Dry Eye Model Using Complete Dacryoadenectomy in Rabbits

All vertebrate animal studies were completed in accordance and compliance with all relevant regulatory and institutional guidelines. All studies were approved by the Institutional Review Board of Stony Brook University and performed in accordance with the Association for Research in Vision and Ophthalmology (ARVO) statement for the use of animals in ophthalmic and vision research.

1. Animals and housing

1. Use New Zealand White (NZW) rabbits weighing 2−3 kg.
2. House rabbits individually in a strictly controlled environment: temperature (65 ± 5 °F), humidity (45 ± 5%), and lighting (12 h on/off cycle).
   NOTE: Due to aggressive behaviors often exhibited between rabbits that are group-housed, keep animals in individual cages to prevent inadvertent ocular injury.
3. Give rabbits unlimited access to standard rabbit chow and water.
4. Provide no other dietary enrichments so as to prevent inadvertent vitamin A supplementation which might affect dry eye.
5. Acclimate rabbits at least two weeks before recording DED parameters.

2. Removal of the nictitating membrane

NOTE: For simplicity, the technique for the right eye is described below. Complete this procedure on the left eye in an identical manner.

1. Remove the nictitating membrane bilaterally during the acclimation period (usually the first week).
2. Place rabbit in an appropriately sized restraining bag.
3. Administer a subcutaneous injection of acepromazine (1 mg/kg) over the shoulders using a 1 cc syringe and 26 G needle to sedate the rabbit. The endpoint for this mild sedation is when the animal maintains a relaxed head position without normal scanning movements and its ears are no longer fully upright.
4. Using a micropipette, apply 25 µL of preservative-free lidocaine (1%) to the eye. Insert a wire lid speculum between the eyelids.
5. Grasp the nictitating membrane at its apex with 0.3 forceps (or equivalent) and pull it over the corneal surface. Inject 1% lidocaine with 1:100,000 epinephrine into the subconjunctival space of the nictitating membrane using a 26 G needle. Inject approximately 0.3 mL to form a modest-sized bleb over the nictitating membrane. Injection volumes over 1 mL are well within a safe dose range for rabbits (2−4 mg/kg).
6. Remove the wire speculum. Wait approximately 5 min for the lidocaine and epinephrine to take effect. During this time, perform the same procedure in the fellow eye.
7. Replace the wire lid speculum. Grasp and extend the nictitating membrane over the corneal surface using 0.3 forceps. Cut the membrane at its base with tenotomy scissors or equivalent.
   NOTE: Bleeding is usually minimal, but keep a high-temperature battery cautery unit nearby and use as needed to minimize bleeding. Direct pressure over the cut base of the nictitating membrane can also be used to stop small bleeding if it occurs.
8. Remove the wire lid speculum. Place topical antibiotic ointment (neomycin, polymyxin, bacitracin, and hydrocortisone) over the corneal surface.
9. Perform an identical procedure to the fellow eye as indicated by the protocol.
10. Place animals back in individual cages and allow to heal for at least one week, or until the conjunctival surface has healed completely from a clinical standpoint, before performing any further assays or interventions.
    NOTE: Complete clinical healing is indicated by a lack of any swelling, injection or discharge from the conjunctival surfaces. Animals should keep their eyes open normally, without the presence of protective ptosis.

3. Measurement of dry eye parameters and collection of tear samples

1. Measure the following DED parameters, as appropriate to the experimental protocol: tear osmolarity, tear break-up time, Schirmer's tear test, and rose bengal staining. Perform them as previously described¹², with a team of at least two investigators.
   NOTE: A team of at least two investigators allows for the efficient measurement of larger groups of animals (6 or more) around the same clock time thereby preventing possible circadian variation from impacting results.

4. Surgical preparation and anesthesia

1. Lightly sedate animals placed in a restraining bag with subcutaneous acepromazine as above (1 mg/kg).
2. Remove all fur on the face and dorsal surface of the skull to visualize the surgical landmarks.
   1. Trim fur with cutting shears leaving residual fine fur about 1 mm in length (Figure 4A, left).
   2. Remove all residual fur using mild depilatory cream following the manufacturer's instructions (Figure 4A, right).
3. Mark surgical incision sites with a surgical pen.
   1. Identify the incision site over the posterior incisure by applying medial pressure to the globe causing a small bulge to develop in the skin over the posterior incisure from prolapse of the OSLG.
   2. Make a linear 2 cm mark in the anterior/posterior direction on the skin over the dorsal surface of the skull directly over this site with a surgical marking pen.
   3. When planning the incision for removal of the ILG, mark a long, curvilinear line around the eye (1 cm from the inferior and temporal lid margin) extending from the posterior (temporal) orbit to the anterior (medial) canthus. Make the marking extend along the posterior orbit to the level of the medial canthus or just superior to this (Figure 4B). In some dissections, the incisions to remove the OSLG and the ILG will be connected.
      NOTE: When performing bilateral surgery, mark both orbits at this time.
4. Trim a patch of fur 2 to 3 cm wide with shears over the lateral surface of each thigh to allow placement of a monopolar cautery plate.
   1. Apply ultrasound gel to ensure good electrical contact with the monopolar cautery plate.
5. Place a 25 G intravenous (IV) catheter in one of the marginal veins of the ear to administer medications or fluids if needed.
6. Give subcutaneous xylazine (1 mg/kg) and IV ketamine (15 mg/kg) for the initial induction of anesthesia (through the IV access).
   NOTE: If the rabbit is sedated beforehand with acepromazine sufficient to retain the endpoint described in step 2.3, use gas mask sedation with isoflurane as an alternative.
7. Place a laryngeal mask airway held in place using an elastic band or string to secure and maintain the airway.
   1. Connect the mask to the anesthesia machine with oxygen flow set at 1 L/min.
   2. Set the isoflurane at 5% initially and then reduce as tolerated based on the level of animal sedation.
   3. Maintain isoflurane at or above 2% until final wound closure.
      NOTE: Assess the level of sedation by monitoring for respiratory rate and movements in response to surgical or painful stimuli. Increase the depth of anesthesia if respiratory rate increases above 10 breaths per minute, if the rabbit begins to chew on the airway maintainer, or if any movements in response to painful stimuli are observed.
8. Monitor pulse oximetry, capnography, blood pressure, rectal body temperature and heart rate using a multi-parameter monitoring device or other appropriate devices.
   1. Monitor vitals continuously during the procedure and record every 10 to 15 min.
9. Position the rabbit on the operating room (OR) table over a heating pad to prevent hypothermia. Tilt the table in a reverse Trendelenburg position at approximately 30° to minimize bleeding.
10. Prepare the surgical area with a povidone-iodine solution diluted to half-strength with sterile water and drape so as to maintain a sterile field.

5. Complete surgical dacryoadenectomy

NOTE: The complete surgical dacryoadenectomy, as described herein, was done using 0.3 tissue forceps, tenotomy scissors, non-toothed tissue forceps, and scissors. These instruments can be interchanged with similar instruments that perform the same function based on surgeon preference.

1. Remove the OSLG first.
   1. Infiltrate the incision sites (surgical marking pen lines and upper posterior lid) with a 50:50 mixture of 2% lidocaine with 1:100,000 epinephrine and 0.5% bupivacaine using a 5 cc syringe with a 30 G needle (Figure 5A).
      NOTE: Syringe and needle size are not critical.
   2. Use a Colorado needle connected to an electrosurgical unit to make the skin incisions along the surgical markings. Settings can vary based on clinical response and typically are between 10 to 15 units for both cut and coagulation (Figure 5B).
   3. Apply opposing tension across the skin incision to separate the tissues and expose the underlying frontoscutularis muscle fibers.
   4. Apply medial pressure on the globe to aid visualization of the OSLG, seen as bulging tissue located just medial or deep to the frontoscutularis muscle fibers. If necessary, move these muscle fibers to the side in order to expose the underlying incisure.
   5. With toothed forceps (0.3) and capsulotomy scissors, gently retract and cut the fibrous capsule overlying the OSLG. The OSLG typically has a pale tan color (Figure 5C).
   6. Using toothed or non-toothed forceps, grasp the OSLG gland tissue and gently pull it out through the superior incisure using a "hand over hand" technique. Cut small, fibrous bands using capsulotomy scissors to free the gland from its position in the orbit (Figure 5D).
      NOTE: As the OSLG gland tissue is removed, it will begin to coalesce into a large tube-like structure (main excretory duct).
   7. When the gland has been removed as completely as possible, use generous cautery with the Colorado needle to create tissue char, truncating the gland within the incisure as deeply as possible. This will later serve as a confirmatory landmark during removal of the PSLG.
2. Remove the PSLG.
   1. Evert the upper eyelid using a cotton-tipped applicator. The bulbous end of the PSLG is usually easily visible.
      NOTE: In some anatomic dissections, it may be possible to visualize the main excretory duct as a pale linear structure about 1 or 2 mm wide.
   2. Engage the PSLG with toothed forceps (0.3) and retract it from the eyelid surface while using capsulotomy scissors to cut around its base separating it from the underlying tarsus (Figure 6A).
   3. Control moderate bleeding with the monopolar cautery.
   4. Apply continuous traction on the separated tissue to maintain a tissue plane for dissection. This will allow the main excretory duct of the SLG to be removed as well (Figure 6B).
      NOTE: As the dissection is carried out, it will typically advance to the superior orbital rim where it is possible to see the cautery marks left behind from removal of the more superior and medially located OSLG.
3. Resect the ILG.
   1. Allow at least 5 min for the local anesthetic to take effect.
   2. Incise the skin, the depressor muscle of the inferior palpebra, the zygomaticolabial part of the zygomatic muscle, and orbicularis muscle with the Colorado microdissection needle and separate as for the OSLG in section 5.1.
   3. Maintain hemostasis with the monopolar cautery.
   4. As the incision is carried deeper through the skin marking, look for the sheen of a fascial plane over the zygomatic bone or superficial part of the masseter muscle. At this point, maintain the tissue plane and carry it superiorly toward the orbital rim using the Colorado needle for cutting (Figure 7A).
      NOTE: For the purpose of identifying the ILG, it is easiest to perform this part of the dissection over the head of the ILG which is typically inferior to the anterior limbus of the eye.
   5. After identifying and incising the capsule surrounding the ILG, identify the tan tissue of the ILG. Only the anterior portion of the ILG head will be visible (Figure 7B). However, the head can be followed medially as it passes beneath the zygomatic arch and transitions into the tail (Figure 7C).
   6. Use tenotomy scissors to cut the orbital septum along the inferior rim exposing the more posterior portion of the ILG tail. Once the tissue plane is identified, extend the dissection posteriorly along the entire incision line (Figure 7D).
      NOTE: The duct of the ILG passes through the lower fibrous connective tissues to enter the inferior conjunctival space in the temporal aspect of the lid. At the posterior rim, the tail of the ILG can have varying anatomic configurations. Sometimes it terminates inferior to the posterior (lateral) canthus, while in other dissections it extends more superiorly around the temporal orbit.
   7. Use extreme care to prevent inadvertent damage to the blood supply, which the ILG receives from branches of the carotid artery. The blood supply can be seen during this part of the dissection (Figure 7E).
   8. In cases where the tail terminates under the posterior (lateral) canthus, it may be necessary to bisect the temporal portion of the frontoscutular muscle to expose the tail of the ILG, which lies along the zygomatic bone.
   9. After the entire ILG has been isolated and exposed, remove it. Due to its large size, it is often preferable to cut the gland in half with scissors and remove the head separately from the tail.
   10. Proceed very cautiously when removing the head of the ILG as it lies immediately adjacent to a large venous sinus in the orbit. Although bleeding from this structure during surgical resections has not occurred, have ample hemostatic aids present to mitigate this risk.
   11. After removal of all gland tissue, close the deep connective tissue plane with multiple interrupted 5-0 ethylene terephthalate sutures. Close the superficial muscles and skin with a running 6-0 polyglactin 910 suture (Figure 7F) using 0.3 tissue forceps and a needle driver.

6. Postprocedural care

1. Undrape the animals and cleanse the surgical sites with sterile water.
2. Apply topical ophthalmic antibiotic and steroid ointment (neomycin, polymyxin, bacitracin, and hydrocortisone) to the incisions. Continue this application twice a day for 2 days.
3. Give a subcutaneous injection of 20 mL normal saline over the shoulder blades using a 26 G needle.
4. Give subcutaneous buprenorphine 0.01 mg/kg or ketoprofen 3 mg/kg for pain control using a 1 cc syringe and 30 G needle.
   NOTE: Animals should return to their normal dietary intake and activities within 1−2 days. Rabbits should be evaluated at least weekly for clinical signs of infection as evidenced by progressive swelling, pain, erythema, calor or purulent discharge over the incision sites. Animals must also be observed to make sure they do not begin to scratch the incision sites/suture lines. Trimming all claws prior to dacryoadenectomy may be helpful in this regard. If scratching the incision lines is observed, standard protective collars may be used to prevent self-injury.
5. Reverse the anesthesia.
   1. Remove the airway maintainer after the animal responds to stimuli and begins to show spontaneous chewing but before the airway maintainer can be damaged.
   2. Monitor animals for approximately 1−2 h or until they have completely recovered from anesthesia as evidenced by spontaneous movement in their cages.
   3. Assess animals for pain and treat appropriately.
6. Allow animals to recover for at least 1 week following surgery before making any clinical measures of DED.