Reduction of Intraocular Pressure in Mouse Eyes Treated with Latanoprost

Abstract

purpose. To determine whether topical treatment of mouse eyes with latanoprost alters intraocular pressure (IOP).

methods. In a masked study design, NIH Swiss mice received a 2-μL topical instillation of 0.00015%, 0.0006%, 0.0025%, or 0.01% latanoprost or vehicle (phosphate-buffered saline [PBS]). After 1, 2, or 3 hours, the animals were anesthetized, and a fluid-filled glass microneedle connected to a pressure transducer was inserted through the cornea into the anterior chamber to measure IOP. The reduction of IOP after latanoprost measurement was calculated by the comparison between treated and nontreated eyes in the same mouse. The effect of latanoprost after a single 0.01% dose was also measured at 6, 12, and 24 hours. As in the previous study, the identity of all eye drop solutions was masked.

results. In mouse eyes receiving topical PBS, the mean IOP was 14.8 ± 2.2 mm Hg (n = 173 males). There was no significant difference in IOP between male and female eyes and between right and left eyes. At 1 hour after topical treatment with 0.00015% or 0.0025% latanoprost, IOP increased by as much as 11% ± 7%. At 2 and at 3 hours after application, IOP decreased in a dose-dependent manner. These decreases were significant in eyes receiving 0.0025% or 0.01% latanoprost (P < 0.05, Student-Newman-Keuls test) and the largest decrease (14% ± 8%) was noted 2 hours after treatment with 0.01% latanoprost. At 6, 12, or 24 hours after treatment, there was no difference in latanoprost- and PBS-treated eyes.

conclusions. Latanoprost reduces mouse IOP in a dose-dependent manner. The mouse may be a useful model for studying the effect of drugs on IOP.

The availability of an animal to study the effects of molecular genetic manipulations on intraocular pressure (IOP) would provide an important new model for investigating glaucoma. The mouse has been extensively studied in many experimental fields because of the feasibility of using transgenic technology, as well as the mouse’s ease of handling and relatively low cost when compared with other mammals. ¹ Moreover, the mouse eye has many structural similarities to the human eye including a well-defined trabecular meshwork, Schlemn’s canal, ciliary body, ² and vascularized retina. ³ Recently, John et al. ⁴ reported an invasive method to measure mouse IOP and showed differences of IOP among four strains of inbred mice. 

The IOP response of the mouse eye to drug treatment has not been reported previously. The present investigation was undertaken to characterize normal IOP in the NIH Swiss mouse and to determine its response to latanoprost, a widely used treatment for lowering IOP in glaucoma. 

Methods

Animal Husbandry and Validation of IOP Measurement

All experiments were performed in compliance with the ARVO Statement for the Use of Animals in Ophthalmic and Vision Research. NIH Swiss mice were obtained from Harlan Sprague-Dawley (San Diego, CA). Mice were received 6 weeks after birth and were housed in clear cages covered loosely with air filters and containing white pine shavings for bedding. The environment was kept at 21°C with a 12-hour light and 12-hour dark cycle. All mice were fed ad libitum. Animal age ranged from 6 to 9 weeks. Body weight ranged from 25 to 36 g at the time of IOP measurement. In all experiments, IOP was measured only once in each mouse. 

The mice were anesthetized by intraperitoneal injection of a mixture of ketamine (100 mg/kg, Ketaset; Fort Dodge Animal Health, Fort Dodge, IA) and xylazine (9 mg/kg, TranquiVed, Vedco, Inc., St. Joseph, MO), prepared at room temperature. To minimize stress, the animals were gently immobilized in a plastic film restrainer (Braintree Scientific Inc., Braintree, MA), and anesthesia was administered with a 30-gauge needle. A timer was started immediately after the injection. Each mouse was monitored carefully to assess the state of anesthesia. After the mouse lost consciousness and failed to blink after topical instillation of phosphate-buffered saline (PBS), the measurement was performed. 

IOP measurement was performed as described previously by John et al., ⁴ with minor modifications. As diagrammed in Figure 1a sterilized, water-filled microneedle was used to cannulate the anterior chamber. To make the microneedle, borosilicate glass tubing (outer diameter, 1.0 mm; inner diameter 0.58 mm; Kwik-Fil; World Precision Instruments [WPI], Inc., Sarasota, FL) was pulled with a pipette puller (P-87; Shutter Instruments, Novato, CA), and the tip was beveled to 30° with a microgrinder (Micropipette Beveler; WPI). The outer diameter of the tip was 50 to 75 μm. The microneedle was connected to a pressure transducer (Blood Pressure Transducer; WPI), which relayed its signal to a bridge amplifier (Quad Bridge; ADInstruments [ADI], Castle Hill, New South Wales, Australia). The amplifier was connected to an analog-to-digital converter (Power Laboratory; ADI) and a computer (G4 Macintosh; Apple Computer Inc., Cupertino, CA). All data were collected and analyzed by computer (Chart 3.0 software; ADI). 

After the needle was filled and mounted to the transducer, the valve to the needle was closed, and the pressure in the system was raised to 40 mm Hg by a 10-mL syringe connected to the tubing circuit between the pressure manometer and the transducer. The system pressure was varied from 0 to 50 mm Hg, according to the manometer, and transducer readings were calibrated using the software. Next, the tip of the needle was placed in PBS drops and zero pressure readings were confirmed. 

As soon as the mice were anesthetized sufficiently, they were placed on a platform made with synthetic modeling clay (Polyform; Elk Groove Village, IA) shaped like a vaulting horse and gently immobilized with stainless steel wire head and tail holders that were pressed into the clay. When manipulating the animals, care was taken to avoid exerting pressure on the neck, which could alter IOP. All the following procedures were performed under a dissecting microscope. PBS (2 μL) was placed on the eye to prevent corneal dehydration and to allow a clear microscopic view of the anterior chamber. The microneedle tip was placed in the drop of PBS over the eye, and the pressure was confirmed to be 0 mm Hg. The tip of the microneedle was inserted into the anterior chamber through the cornea by use of a micromanipulator. After insertion, the microneedle tip usually rested 50 to 100 μm into the chamber. The microneedle was then repositioned to minimize corneal deformation and to ensure that the eye remained in its normal position. After measurement, the microneedle was withdrawn from the anterior chamber so that the tip was located again in the drop of PBS. Rapid return of the pressure reading to zero was required for the data to be included. If prolonged pushing was required for insertion, the data were discarded, because the IOP measurement may have been artificially low. 

Effect of Latanoprost on Mouse IOP

Latanoprost (Cayman Chemical Co., Ann Arbor, MI) was diluted to concentrations of 0.00015%, 0.0006%, 0.0025%, and 0.01% in PBS just before application. It was applied randomly to the right or left eye, and PBS was applied to the contralateral eye. The person administering the drops and then recording the IOP was masked to the identity of the various eye drop solutions. With a micropipette, 2 μL latanoprost solution was administrated topically to one eye, and 2 μL vehicle solution was administrated to the other. The time between drop administration and IOP measurement was 1, 2, 3, 6, 12, or 24 hours. All measurements were completed before the masking code was broken. The effect of the drug’s application was calculated as the difference of IOP between the treated eye and the contralateral control eye in each mouse. 

Statistical Analysis

Results from each latanoprost group (latanoprost in one eye and PBS in the contralateral eye) were compared with the vehicle-only control group (PBS in both eyes) using analysis of variance and the Student-Newman-Keuls (SNK) test. P < 0.05 was considered statistically significant. All data are presented as the mean ± SD. 

Results

Validation of IOP Measurement

Effect of Anesthesia.

To evaluate the effect of anesthesia, the IOP of one eye was monitored in 26 mice as soon as possible after the mouse lost consciousness. The right or left eye of one mouse was selected at random for the measurement. In 62% of the mice, IOP was stable from 3 to 7 minutes after anesthesia. In the remaining mice, IOP gradually increased at approximately 0.25 mm Hg/min for 2 minutes, followed by steady state pressure for approximately 2 minutes, and then gradual reduction of IOP at approximately 0.25 mm Hg for the next 3 minutes. Figure 2 shows the gradual increase and decrease of mean IOP that occurred in 26 mice between 3.7 and 7.3 minutes after injection of anesthesia. The peak of the IOP increase occurred 5.0 ± 0.8 minutes after anesthesia and was 15.8 ± 2.3 mm Hg. The time interval during which IOP ranged within the 3% of the peak IOP (approximately 0.5 mm Hg) was 72 seconds before and after the peak. Hence, for subsequent single measurements, the IOP was recorded at 4 minutes if the reading was not increasing and the variation was less than 0.2 mm Hg for at least 10 seconds. 

IOP in Untreated Eyes.

The mean IOP in anesthetized male mice was 14.8 ± 2.2 mm Hg (mean ± SD, n = 173; Table 1 ). These measurements were collected between 8 AM and 10 PM. In preliminary studies, we observed a difference of IOP measured in the midday and at night. To preclude the possibility that such a difference, if it existed, would influence our results, we limited measurements to a specific time (10 AM to 2 PM). The mean IOP between 10 AM and 2 PM was found to be 14.2 ± 2.0 mm Hg (n = 80). The IOP in 20 female mice was measured between 10 AM and 2 PM and found to be 13.3 ± 1.2 mm Hg. This result was insignificantly different from the male mouse IOP measurements collected during the same period (t-test P = 0.11). 

Comparison of Bilateral IOP Measurements.

In 30 male mice, the IOP of the right eye (Td) and the left eye (Ts) was measured sequentially. To avoid the variation in IOP after anesthesia, the IOP of the first eye was recorded at 4 minutes after anesthesia, and the IOP of the second eye (T2) was measured within 1 minute after the measurement of the IOP of the first eye (T1). Thus, both the first and the second measurements were recorded within the plateau phase (Fig. 3) . As before, these measurements were collected between 10 AM and 2 PM to avoid the possible influence of normal diurnal IOP variation. Moreover, the order of measurements of Td and Ts was selected at random to avoid the possible systematic influence of order. The averages of Td and Ts were 14.8 ± 1.9 and 14.6 ± 1.6 mm Hg, respectively, and were not significantly different (t-test, P = 0.661, n = 30; Table 1 ). The mean difference between the IOP measurements of both eyes in each mouse (Td − Ts) was 5.3% ± 4.2% of the average IOP measurement (0.78 ± 0.62 mm Hg). The influence of measurement order, as determined by T1 − T2, was nonsignificant (t-test, P = 0.665, mean of T1 − T2 = 0.20 ± 0.98 mm Hg). 

Effect of Latanoprost on Mouse IOP

Dose response and time course of IOP change induced by latanoprost were determined by measuring IOP at 1, 2, and 3 hours after treatment with a single 0.00015%, 0.0006%, 0.0025%, and 0.01% dose administered at 9:00 AM (Fig. 4) . There were eight mice treated for each measurement point. The change of IOP induced by latanoprost was expressed as the difference between the IOP of the control eye and the IOP of the treated eye. At 1 hour after treatment, IOP increased by 8% ± 8% or 11% ± 8% in the eyes that had received 0.00015% or 0.0025% latanoprost, respectively. At 2 and at 3 hours after treatment, however, IOP decreased in a dose-dependent manner within the latanoprost-treated eyes. This reduction was significant at both 2 and 3 hours in eyes that had received either 0.0025% or 0.01% doses. (SNK test, P < 0.05). 

In view of these results, the long-term effect of 0.01% latanoprost was investigated at 6, 12, and 24 hours after administration. These results are plotted together in Figure 5 with the short-term results from Figure 4D . Although the mean IOP at 6 hours was less than in control vehicle-treated eyes, the reduction was nonsignificant. The differences in IOP from vehicle-treated eyes were also nonsignificant at 12 and 24 hours after treatment. In all these experiments, no change in the external appearance of the eyes was observed by stereo microscope. 

Discussion

Topical treatment with 0.0025% or 0.01% of latanoprost significantly decreased mouse eye IOP in a dose-dependent manner at 2 and 3 hours after application. Bilateral comparison of IOP using the microneedle method was rapid, and the measurements were unaffected by the order in which they were obtained. Measurements were discarded when there were difficulties with needle insertion through the cornea (<5% of attempts). Moreover, there was no significant difference in the IOP measurements of the left and right eyes of mice that received vehicle in both eyes. 

The present results support previous observations indicating that normal IOP in mice appears to be strain specific. In the present study, mean IOP in the NIH Swiss mouse was 14.8 ± 2.2 mm Hg (mean ± SD). In a previous report, IOP of C3H/HeJ, C57BL/6J, A/J, and BALB/cJ mice was 13.7 ± 0.8, 12.3 ± 0.5, 9.4 ± 0.5, and 7.7 ± 0.5 mm Hg (means ± SE), respectively. ⁴ These differences in strains may reflect genetic differences within tissues of the inflow and outflow pathways. In addition to illustrating the importance of characterizing the normal IOP of a particular mouse strain before experimental studies are performed, the current results demonstrate the importance of the cannulation method and the anesthesia protocol. 

A challenge in designing experiments to investigate the change in IOP is that IOP can vary widely among different mice. However, it is well established that in humans, as well as in many larger laboratory animals, IOP in one eye is similar to IOP in the other eye. ^{5 6} Hence, in experimental investigations of IOP regulation, the treated eyes are often compared with the contralateral control eye. During the use of general anesthesia, however, there is concern that IOP might change during anesthesia induction and that IOP might not be similar if the measurements of the two eyes are not acquired simultaneously. Indeed, in the present study we found that between 3 and 7 minutes after administration of anesthesia, IOP either was stable or gradually increased and then later gradually decreased. However, in the latter cases there was a 2-minute period between the increasing and decreasing phases in which IOP varied by less than 3%. By collecting the IOP measurements from both eyes during this plateau phase, possible variation due to anesthesia was minimized, and it was observed that IOP was essentially the same in both eyes. These data indicate that it is appropriate to conduct mouse experiments in which one eye receives an experimental treatment and the other eye receives a control treatment, if both IOP measurements are collected during the plateau phase. 

The microneedle technique has several advantages and limitations when compared with other IOP measurement techniques. A major limitation of noninvasive methods has been lack of calibration of the instrument to the mouse eye. In contrast, one of the advantages of the present microneedle technique is direct calibration, which allows for confirmation of good accuracy. A second advantage of our technique is that the chart recording output of the data recorder allows assessment of the quality of the measurement (the IOP trace should be stable during the measurement and should return to zero when the microneedle tip is moved from the anterior chamber to the overlying PBS droplet). Moreover, a permanent record of the IOP measurement is obtained. A limitation of this technique is that is cannot be repeated for several days, to allow healing of the corneal perforation. If repeated measurements were obtained in the same animal, inflammation and healing incited by the needle insertion also might affect the IOP after the first measurement. Another limitation is that general anesthesia is required, and many common anesthetics, including the ketamine-xylazine mixture used in the present study, have been shown to alter IOP in other rodents. ⁷ As shown in the present study, however, the anesthesia effects are reproducible, and useful readings can be obtained during the plateau phase of the IOP response. 

Topical application of 50 or 200 ng (2 μL of 0.0025% or 0.01% solution) latanoprost lowers mouse IOP measured at 2 and 3 hours after treatment. This dose appears to be similar to the optimal dose for human eyes when allowance is made for the difference in size of the anterior chamber volume. The optimally effective dose of latanoprost for IOP reduction in humans is 1.5 μg ⁸ and the average volume of the anterior chamber in older humans is 160μ L. ⁹ The aqueous volume of mouse anterior chamber can be estimated by measurement with a microneedle and is approximately 8 μL (data not shown). Assuming that the permeability of latanoprost into anterior chamber in mouse and human eyes is similar, combining these measurements to estimate the effective dose for the mouse yields a dose of 75 ng per eye (1.5 μg × 8 μL/160 μL). This calculated dose corresponds well with the present observations that significant IOP reduction occurred after 50-ng (2 μL of 0.0025% solution) and 200-ng (2 μL of 0.01% solution) treatments. Moreover, this similarity in dose requirement suggests that there may be similarities in the tissue responses in human and mouse eyes. 

The magnitude and timing of IOP reduction observed in latanoprost-treated mouse eyes also shows similarities with previous observations. A single application of 0.01% latanoprost reduced mouse IOP by 14% (1.97 mm Hg/14.2 mm Hg) in 2 hours compared with the nontreated eye. This ocular hypotensive effect in mouse was similar to the maximal IOP reduction seen in latanoprost-treated monkey eyes (approximately 15%) but lower than in human eyes (approximately 30%). ^{8 10} The magnitude of the IOP reduction in the mouse peaked at 2 hours and returned to baseline by 3 hours after treatment. This is similar to the monkey, in which the IOP trough spanned from 1 to 2 hours after treatment. ¹⁰ However, there was no effect of single application in mouse eyes at 6 hours, whereas in monkey and human eyes the IOP reductions may persist for 6 to 10 hours. ^{8 11} The basis for these difference in duration are not known, but may reflect, in part, faster clearance of the drug from the smaller mouse eye. Unlike the case in human, monkey, and mouse, latanoprost has no IOP-reducing effect in cat or rabbit. ^{12 13} These differences may reflect different distributions of EP and FP receptors or structural differences in the uveoscleral outflow pathway tissues. 

In conclusion, the intraocular pressure of mouse eye is reduced after treatment with latanoprost. This model can be used to investigate the molecular mechanism of latanoprost-mediated IOP reduction and also may be used to study the IOP-lowering effects of other drugs. 

 Supported in part by Grant EY-05990 from the National Eye Institute (RNW), the Foundation for Eye Research (MA), and the Joseph Drown Foundation (JDL). MA was supported by the Department of Ophthalmology, Faculty of Medicine, University of Tokyo, Tokyo, Japan.

 Submitted for publication June 21, 2001; revised August 22, 2001; accepted September 10, 2001.

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Figure 1.

 

Diagram of the equipment used for mouse IOP measurement.

Figure 2.

 

Effect of general anesthesia on mouse IOP. Data indicate the time course of mean IOP change after the intraperitoneal injection of anesthesia. Error bars indicate SD (n = 26 mice).

Table 1.

 

Normal IOP of NIH Swiss Albino Mice

Purpose	IOP*	Gender	n	Time Range
Average IOP throughout the day	14.8 ± 2.2	M	173	8 AM to 10 PM
Average IOP near the noon hour	14.2 ± 2.0	M	80	10 AM to 2 PM
Influence of gender	13.3 ± 1.2	F	20	10 AM to 2 PM
Bilateral measurements
Right eye	14.8 ± 1.9	M
Left eye	14.6 ± 1.6	M	30	10 AM to 2 PM
Difference between paired eyes	0.78 ± 0.62	M
Measurement order (T1− T2), †	0.20 ± 0.98	M

*  Data expressed as mean mm Hg ± SD.

^†  T1 and T2 are the IOP of the first and second measured eyes in each mouse.

Figure 3.

 

A sample recording of mouse IOP without treatment. In this case, the left eye IOP was measured first. Each accepted IOP (Td and Ts) was a mean of the area indicated by black bars, which show stable IOP with variation of less than 0.2 mm Hg for 10 seconds.

Figure 4.

 

Short-term effect of latanoprost on mouse IOP. Data indicate the mean difference in IOP (IOP of treated eye − IOP of control eye), and error bars indicate SD. Latanoprost (2 μL) or vehicle (2 μL) was applied at 9:00 AM and measurements were made 1, 2, or 3 hours later. The tested concentrations of latanoprost were 0.00015% (A), 0.0006% (B), 0.0025% (C), and 0.01% (D). *P < 0.05 by the SNK t-test. The number next to each data point indicates the number of mouse IOP measurements associated with that point. Numbers less than eight reflect measurements disqualified due to technical complications, such as microneedle clogging.

Figure 5.

 

Long-term effect on mouse IOP of treatment with 2 μL 0.01% latanoprost. For each measurement, treatment was performed at 9:00 AM. The data indicate the IOP difference due to treatment (IOP of treated eye − IOP of control eye) and are expressed as mean ± SD.* P < 0.05 by the SNK t-test. The number next to each data point indicates the number of mouse IOP measurements associated with that point.

The authors thank Simon W. John, PhD, Jackson Laboratory, Bar Harbor, Maine, for assistance and advice in the method for measuring mouse IOP.