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REGIONAL ANESTHESIA AND PAIN MANAGEMENT

Reproducibility of the Drug Effects over Time on Chronic Lumbar Epidural Catheterization in Rats

Tomoki Nishiyama, MD, PhD^*, and Kazuo Hanaoka, MD, PhD

*Department of Anesthesiology, The University of California, San Diego, California; and Department of Anesthesiology, The University of Tokyo, Faculty of Medicine, Tokyo, Japan

Address correspondence and reprint requests to Tomoki Nishiyama, MD, PhD, 3-2-6-603, Kawaguchi, Kawaguchi-shi, Saitama, 332-0015, Japan.

	Abstract

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Chronically implanted epidural catheters lead to a reaction that impedes drug action. The purpose of this study in a rat model with chronically implanted epidural catheters was to investigate the change in opiate activity and histology over time with this model. A skin incision of 1–2 cm was made at the T 13 level on the back of male Sprague-Dawley rats under halothane anesthesia. Muscles were dissected bluntly from the vertebrae, and the intervertebral ligament was cut to insert an epidural catheter (polyethylene tube, outer diameter of 0.14 mm) 2-cm caudally. The longer portion of the catheter was passed through a trocar subcutaneously to exit the dorsal neck area. One, two, and six days after catheterization, the effects of morphine on thermal stimulation using the hot-box test and histology were investigated. Analgesic effects of morphine 6 days after catheterization were significantly less than those on the first and second days. Histologically, evidence of inflammation around the catheter was noted as early as 4 h after catheterization. Pericatheter fibrosis was severe after 2 days. We conclude that this model of chronic epidural catheterization in the rat evoked a histologically defined, pharmacodynamically significant, local reaction 2 to 6 days after catheter implantation.

Implications: A rat model with chronically implanted epidural catheters should be used for testing the analgesic effects of drugs within two days after catheterization.

	Introduction

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There are many studies on the effects of drugs on nociception using rodent models with chronic epidural catheterization (1–4). Chronically catheterized rats require 4 to 7 days to recover from surgery (1–4). Grouls et al. (3) found no fibrosis around the epidural catheter 4 days after implantation, but did not discuss details. Durant and Yaksh (5) examined, in chronic epidurally catheterized rats, the effects of epidurally administered opioid and nonopioid drugs. They noted a progressive loss of drug effect over a 10-day period after catheter implantation. Histologically, fibrosis around the catheter began 1 day after catheter implantation, and after 10 days of catheterization a thick fibrosis prevented diffusion from the catheter tip (5). This result has implications for other studies that use rats 4 to 7 days after catheterization (1–4). We investigated the changes of the analgesic effects and histology of the spinal cord and catheter area in detail over shorter intervals of a new rat model with a chronic epidural catheter (6).

	Methods

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Experiments were performed according to a protocol approved by our animal care committee.

A new rat model with chronic epidural catheterization has been described in detail in a previous study (6). The epidural catheters were made with a 2-cm length of polyethylene tube (PE-5; Clay Adams, Parsippany, NJ) (outer diameter: 0.14 mm) and heat-melted to a 12-cm length of polyethylene tube (PE-10; Clay Adams, Parsippany, NJ) (outer diameter: 0.61 mm). One loose overhand knot was made at the connection between the PE-5 and PE-10 portions, and the second one was made 5 mm distal to the first knot. Each knot was covered with dental acrylic more than 3 days before insertion and served to secure the catheter. The dead space in the catheter was 8 ± 1 µL (mean ± SD, n = 8).

Male Sprague-Dawley rats (300–350 g; Harlan Industries, Indianapolis, IN) were anesthetized with halothane (4%) in an anesthesia induction box. Upon loss of responsiveness and spontaneous movement, the rat was removed from the induction box and anesthetized continuously with halothane (2%) in an air-oxygen mixture during spontaneous respiration. The dorsal thoracolumbar spinal region was clipped and surgically prepared with alcohol and Betadine. A 1–2-cm midline skin incision was made at the most prominent thoracic spinous process, which is T 13. Muscles were bluntly dissected from the vertebrae and retracted to expose the intervertebral space. The intervertebral ligament was carefully cut, and an epidural catheter inserted into the epidural space 2 cm caudally. The catheter tip was located at the L3-4 level, which was checked anatomically in the preliminary study. The first knot was put into the space between the two vertebrae. The second knot was covered by muscle, and the longer portion was passed through a trocar subcutaneously to exit the dorsal neck area. Failure to obtain fluid from the catheter indicated that the catheter was not placed intrathecally. The catheter was flushed with sterile saline, and no extra vertebral leakage was noted at volumes up to 20 µL. The catheter tip was plugged with a 28-gauge steel wire. The muscles and fascia were sutured with 3–0 Vicryl^TM and the skin was closed using 3–0 silk. Inhalation of halothane was stopped and animals were observed during recovery in a warm box. Only rats with no motor disturbances before the study (98% of the total number of catheter implanted rats) were used. Each rat was used for only one treatment. All animals were killed with an overdose of pentobarbital (100 mg/kg) after the study, and the spinal column was removed with the catheter to verify the position of the catheter tip. The data from rats with the catheter not in the epidural space (5%) were excluded from this study.

Morphine (morphine sulfate, opioid agonist; Merck, Sharpe, and Dohme, West Point, PA) was dissolved in normal saline of 20 µL that contained the desired quantity of the drug. After epidural drug injection, the catheter was flushed by the subsequent injection of 10 µL of normal saline. Microinjector syringes were used for all injections. For each dose, seven randomly selected rats received one of these doses of morphine (0 µg; control: only saline, 30, 100, or 300 µg) on the first, second, and sixth day after surgery.

All animals were tested for their acute nociceptive response using a hindpaw thermal withdrawal test (7). The rats were placed in a clear plastic cage on an elevated floor of clear glass. A radiant heat source (halogen projector lamp CXL/CXP 50 W 8 V; Ushio, Tokyo, Japan) was contained in a movable holder placed beneath the glass floor. The radiant heat source’s aperture was 4 mm in diameter, and bulb intensity was controlled by a constant voltage source. The interior of the box under the animal was prepared with a heat source such that the under-plate temperature was regulated to 30°C. The under-floor heat source was then positioned such that it focused at the plantar surface of one hindpaw where it was in contact with the glass. The calibration of the thermal test system was such that the average response latency in normal untreated rats measured before the initiation of an experiment was approximately 6 s. In the beginning of the test, the rat was placed in the box and allowed to adapt for approximately 20 min. The light beam was then activated. The time interval between the application of the light beam and the brisk hindpaw withdrawal response was measured. Cut-off time in the absence of a response to avoid tissue injury was set to 20 s (8).

The general behavior (including agitation and allodynia), motor function, pinna reflex, and corneal reflex were examined. The former two were scored as follows: 0, normal; 1, slight deficit; 2, moderate deficit; and 3, severe deficit. The reflexes were judged as present or absent. The presence of allodynia was examined by looking for agitation (escape or vocalization) evoked by lightly stroking the flank of the rat with a pointed probe. Motor function was evaluated by observing the placing/stepping reflex and the righting reflex. The former was evoked by drawing the dorsum of either hindpaw across the edge of the table. The latter was assessed by placing the rat horizontally with its back on the table, which normally gives rise to an immediate, coordinated twisting of the body to an upright position.

To determine the differences in analgesic actions of epidurally administered drugs according to the intervals between catheter implantation and testing, rats were assigned to groups to be examined on the thermal escape and behavioral and motor function tests. These measures were performed before and at intervals of 15, 30, 45, 60, 90, 120, 180, 240, and 300 min after epidural morphine injection with different doses of 0, 30, 100, or 300 µg on 1, 2, or 6 days after catheter implantation.

One hour after catheter implantation, or 1, 2, 4, or 6 days after the catheterization, groups of rats were killed with pentobarbital 100 mg/kg. Afterward, 20 µL of blue dye (followed by a flush with 10 µL of saline) was injected into the catheter and the vertebral bone was removed carefully to see the spread of the dye.

Two rats each were killed with pentobarbital 100 mg/kg to check histology at 1 h, or 1, 2, 4, or 6 days after catheter implantation or after sham operation (the same surgical procedure except for catheter insertion). After perfusion fixation using 10% formalin, the spinal cord with the vertebral bone and catheter was removed, decalcified, and fixed in 10% formalin. Tissues around the catheter tip were processed in paraffin and stained with hematoxylin-eosin for light microscopic examination. Spinal cord injury and inflammation were scored from 0 (no injury/inflammation) to 4 (severe injury/inflammation) by an animal pathologist who was blinded to the treatment.

Response latency data from hot-box measurements were converted to percent maximum possible effect (%MPE) according to the formula: %MPE = [(postdrug latency – baseline latency)/(cutoff time – baseline latency)] x 100. The area under the time effect curve (AUC) was calculated as time (min) x %MPE. ED₅₀ values (effective dose resulting in a 50% prolongation of the control hot-box latency) were calculated by a computer program that is made at the Department of Anesthesiology, University of California, San Diego, CA as a dose which produces a value of 50% MPE.

Dose-response data are graphically presented as mean ± SEM. One way analysis of variance was used to see the differences in the effects among the three doses, and a Fisher’s Protected Least Significant Difference test was used as a post hoc test. A P value of <0.05 was considered statistically significant.

	Results

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Dose-dependent increases in the peak effects and the AUC after epidural morphine were seen in all day groups. The time courses of the effect of epidural morphine were similar on all days, but the effects were less on the sixth day than on the first and second days (Fig. 1). The peak %MPE and the AUC showed no differences between the first day and the second day in all doses. However, on the sixth day, both peak %MPE and AUC were lower than those on the first and the second days (Fig. 2).

View larger version (21K): [in this window] [in a new window]   Figure 1. Time course of the effect of epidural morphine 300 µg. Closed circles indicate the data 1 day after catheter implantation, open circles 2 days, and closed squares 6 days after the catheterization (each group: n = 7). Bars indicate SEM. *P < 0.05 versus Day 1, **P < 0.01 versus Day 1, P < 0.05 versus Day 2, P < 0.01 versus Day 2.

View larger version (15K): [in this window] [in a new window]   Figure 2. Peak percent maximum possible effect (%MPE) (upper) and area under the time effect curve (AUC) (lower) of the analgesic effect of epidural morphine administration on the different days. Response latency data from hot-box measurements were converted to %MPE according to the formula: %MPE = [(postdrug latency – baseline latency)/(cutoff time – baseline latency)] x 100. The AUC was calculated as time (min) x %MPE. Closed circles indicate the data 1 day after catheter implantation, open circles 2 days, and closed squares 6 days after catheterization (each group: n = 7). Bars indicate SEM. *P < 0.05 versus Day 1, **P < 0.01 versus Day 1, P < 0.05 versus Day 2, P < 0.01 versus Day 2.

After 1 day of epidural catheterization, each rat in the 100 µg and 300 µg morphine-treated group showed a slight agitation and allodynia and three rats given morphine 300 µg showed motor disturbance (one: moderate in the righting reflex and the placing/stepping reflex; two: slight in the righting reflex). After 6 days of the catheterization, one rat given morphine 100 µg and two rats given 300 µg showed a slight agitation and allodynia (Table 1). These side effects were reversible during the study periods.

Signs of inflammation were observed around the catheter at 4 h after catheterization and continued to develop through 6 days after catheterization (Table 2). The catheter tract displayed significant fibrosis at 6 days after catheterization.

	Discussion

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Van den Hoogen and Colpaert (9) first described the rat model with chronic epidural catheterization. They inserted a PE-10 catheter (outer diameter of 0.61 mm). The placement was accomplished by making a hole/groove, and removing the process and arch of vertebral bone. It was reported that at least one week was necessary for recovery. Because the degree of inflammation might vary with catheter diameter, we used a smaller tube (outer diameter of 0.14 mm) for the insertion part. In addition, in our model, we avoided any intervention in the bone. Such surgery might induce an inflammatory reaction. We avoided the use of glues to affix the catheter. A knot was prepared to fit between the two vertebral bones. This method served to prevent catheter removal and to prevent leakage from the epidural space. The procedure was minimally traumatic, as evidenced by the fact that body weight did not decrease after catheter implantation and all rats showed normal behavior one day after surgery.

Despite this attention to potential variables associated with local reaction, a significant and pharmacologically relevant reaction was noted in the rat. Thus, the ED₅₀ on day six after catheterization (1131 µg) was significantly higher than those on days one (62 µg) and two (94 µg) after catheterization. The values one and two days after catheterization are about 20 to 30 times higher than with intrathecal morphine (3.1 mg: 1.9–5.4 mg; mean 95% confidence interval) tested by the same thermal escape test (10). The potency ratios between epidural and intrathecal morphine observed in the rat are almost equivalent to those used clinically (11–15). The adequate clinical dose of intrathecally administered morphine should be 0.2 to 0.4 mg (11,12). On the other hand, three to five milligrams of morphine is used epidurally for analgesia (13–15). Therefore, the potency between epidural and intrathecal morphine might be about 1:10 to 25. Other experimental studies showed the ED₅₀ values of epidural morphine as 1.5 µg (2) to 2.2 µg (5) in the tail flick test, which were similar with that obtained by intrathecal morphine administration (1.2 µg) using the same test (4). From these observations, our model of epidural catheterization tested for the effects by thermal escape appears to resemble the clinical situation as discussed in our previous study (6).

Regarding the effects of the volume of epidurally administered solutions, we already investigated that 20 µL can give the maximal effects in this model (6). More than 20 µL was not used in the previous study because this volume was the maximum before leakage at the catheter insertion point. Twenty microliters induced sensory block only on the hindpaw but not on the forepaw (6). Based on these preliminary data, drugs for epidural injections were mixed such that all doses were administered in a 20-µL solution.

It should be noted that the epidural catheter reaction is not limited to the rat. Lebeaux (16) inserted an epidural catheter (outer diameter of 1.9 mm) in dogs by laminectomy. The effects of epidural lidocaine did not decrease until four days after catheter implantation. After three weeks, the histology showed inflammatory responses at the orifice of the catheter in the epidural space. In some dogs, the catheter tip was occluded by granulation tissues. Nagaro (17) reported details of the histology and the effects of lidocaine using chronic epidural catheterized dogs (outer diameter of the catheter: 0.8 mm). One week later, the catheterization inflammatory reaction was observed and four weeks later it became fibrous and encapsulated the catheter. Cephalad spread of anesthesia by lidocaine decreased after one week in his model. Thus, fibrous tissue formation and occlusion of the catheter occur in both rats and dogs.

We used polyethylene catheters to compare the results with previous studies (7,9,17). However, other materials, such as hydrogel (18) or carbohydrate polymer (19), are reported to prevent fibrosis. Therefore, catheters made by those materials should be tested in our new rat model of epidural catheterization in the future.

In conclusion, we describe the reaction of the rat tissues in the epidural space to chronically implanted epidural catheters. This local reaction is pharmacologically relevant, as the analgesic effects of morphine decreased within six days after catheterization, and fibrosis capsulated the catheter within four days after catheterization.

	Acknowledgments

 
We thank Professor Tony L. Yaksh, Department of Anesthesiology, University of California, San Diego, for his support.

	References

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