Anesth Analg 2006;103:876-881
© 2006 International Anesthesia Research Society
doi: 10.1213/01.ane.0000237287.53957.18
ANESTHETIC PHARMACOLOGY
Section Editor:
James G. Bovill
Epidural Lidocaine Induces Dose-Dependent Neurologic Injury in Rats
Tomoko Muguruma, MD,
Shinichi Sakura, MD, and
Yoji Saito, MD
From the Department of Anesthesiology, Shimane University School of Medicine, Izumo City, Japan.
Address correspondence and reprint requests to Dr. Muguruma, Department of Anesthesiology, Shimane University School of Medicine, 89-1 Enya-cho, Izumo City, 693-8501, Japan. Address e-mail to hamutaro{at}med.shimane-u.ac.jp.
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Abstract
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Although epidural lidocaine administered as a bolus has been shown to cause little neurotoxicity, local anesthetics are often administered repetitively or continuously into the epidural space, and in high doses may induce neurologic injury. We investigated whether epidural lidocaine is neurotoxic when a large dose is continuously administered in rats, and whether the functional impairment and histologic damage is dose dependent. In Experiment 1, 13 rats received a 120-min epidural infusion (at 5 µL/min) of saline or 2% lidocaine. Four days after infusion, rats given 2% lidocaine developed significantly more prolonged tail-flick latencies and showed more apparent morphologic damage than those given saline. In Experiment 2, 41 rats were randomly divided into 5 groups to receive an epidural infusion of saline for 120 min or 5% lidocaine for 15, 30, 60, or 120 min at a rate of 5 µL/min. Rats given 5% lidocaine for 120 min developed a significant increase in tail-flick latency. Paw pressure thresholds did not change in any group. Nerve injury scores for rats given 5% lidocaine for 30, 60, and 120 min were significantly higher than those for rats given saline. Significant difference in damage in nerve roots was also observed among rats given the anesthetic for different durations of time; nerve injury scores with 120-min infusion were higher than with 15- and 30-min infusions, and injury with 60-min infusion was greater than with 15-min infusion. In conclusion, these results suggest that epidural lidocaine causes dose-dependent neurotoxicity after continuous infusion in rats.
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Introduction
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Increasing laboratory evidence (1–4) suggests that some local anesthetics are potentially neurotoxic, and that neurologic impairment after neuraxial blockade may result from a direct neurotoxic effect of local anesthetics. Interestingly, reported cases of injury are far less common after epidural anesthesia than after spinal anesthesia (5,6). However, the difference may merely reflect more frequent use of spinal anesthesia for blockade of lumbar and sacral nerves, which are the most frequent site of clinical injury.
An in vivo animal study (7) showed that, when intrathecal and epidural lidocaine were administered as a bolus to produce similar anesthetic effects, neurotoxic effects of epidural lidocaine were much less severe than those of spinal lidocaine and similar to those of epidural saline. However, in clinical practice, patients often receive epidural local anesthetic repetitively or continuously with the total dose of anesthetic being much larger than that by a single injection; epidural anesthesia is intended to be used for larger and longer surgical operations than spinal anesthesia. Local anesthetic neurotoxicity is dose-dependent in in vitro (8–13) as well as in vivo (2,14) experiments. Thus, large doses of local anesthetic might induce neurologic injury, even when administered into the epidural space. Accordingly, the current experiments sought to determine whether epidural lidocaine induced dose-dependent, functional impairment and morphologic damage when large doses were continuously administered in rats.
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METHODS
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The protocol was approved by the Animal Research and Use Committee of Shimane University. Two experiments were conducted on male Sprague-Dawley rats (245–320 g), which were maintained on a 12-h light–dark schedule and were housed individually with free access to food and water. To reduce the influence of handling on behavioral reactions, all rats were trained in the test situation at least twice before epidural catheterization.
Surgical Procedure
For epidural catheterization, anesthesia was induced with sodium pentobarbital (Dainippon Pharmaceutical, Osaka, Japan) (30 mg/kg intraperitoneally) and maintained with 1.5% halothane. Using an aseptic technique, a midline skin incision was made over the spinous processes of the L2–5 vertebrae with a foam block placed under the animal’s abdomen. The fascia was opened, and superficial muscles around the spinous processes were dissected and retracted laterally. With the use of fine forceps, the interspinous ligament was pierced, and a heat-connected catheter, composed of double-stretched polyethylene tubing PE-10 (1.2 cm), PE-10 (10 cm), and PE-20 (7 cm), was introduced into the epidural space. The double-stretched PE-10 was passed through the L4–5 intervertebral space and advanced 1.2 cm in the caudal direction. Before starting experiments, rats were allowed at least 2 days to recover from the operation. Rats having any problem with tail movement or motor dysfunction in the hindlimbs were not used in the experiments.
Functional Examination
Neurologic function was measured by an investigator who was blinded to the group assignment. The tail flick (TF) test was performed on rats placed in an acrylic restraint to measure the response of the tail to noxious somatic stimulus using TF equipment (model DS20; Ugo Basile, Comerio-Varese, Italy). A 100-W projector lamp was focused on three segments of the tail approximately 3, 5, and 7 cm from the tip. The time at which rats withdrew the tail from the heat source was defined as the TF latency, and the mean value of the three segments was used for analysis. A cutoff time of 10 s was used to avoid damage to the tail.
The paw pressure (PP) test was performed to measure the response of the legs to noxious mechanical stimulus (15). Pressure was applied to the dorsal surface of both hindpaws using a device (type 7200; Ugo Basile, Comerio-Varese) capable of progressively increasing the pressure at a rate of 15 g/s. Rats were wrapped with a cloth for the test. The pressure at which rats withdrew the paw from the device was defined as the PP threshold, and the mean value of both paws was used for analysis. A cutoff pressure of 400 g was used to avoid damage to the paws.
Motor function (MF) of the posterior limbs was assessed by bilaterally grading the motor block as follows: 0, none; 1, partially blocked; and 2, completely blocked (16). Motor blockade was graded as none when the rat had no visible limb weakness and normal gait; as partially blocked when the limb was able to move but not able to support the normal posture; and as completely blocked when the limb was flaccid, with no detectable resistance to extension of the limbs. The normal baseline score was 0, and the score with bilateral complete block was 2 + 2 = 4. TF, PP, and MF tests were performed sequentially at the same time point with a 15-s interval.
Experimental Protocols
In Experiment 1, 13 rats were randomly divided into 2 groups to receive a continuous epidural infusion of saline (Group S, n = 6) or 2% lidocaine in saline (Group L, n = 7) for 120 min at a rate of 5 µL/min. In Experiment 2, 41 rats were randomly divided into 5 groups to receive a continuous epidural infusion of saline for 120 min (Group S120, n = 7) or 5% lidocaine in saline for 15 (L15, n = 9), 30 (L30, n = 9), 60 (L60, n = 8), or 120 min (L120, n = 8) at a rate of 5 µL/min. Lidocaine solutions were prepared by dissolving crystalline lidocaine hydrochloride (Sigma Chemical, Steinheim, Germany) in sterile normal saline (Otsuka Pharmaceutical, Tokyo, Japan). The osmolarity and pH of all the solutions were measured (Auto&Stat, OM-6030, Arkray, Kyoto, Japan; and pH meter, F-22, Horiba, Kyoto, Japan). The osmolarities of saline, 2% and 5% lidocaine solutions were 289, 413, and 587 mOsm/L, respectively, and the pH 6.06, 4.82, and 4.71, respectively. All infusions were administered by a mechanical infusion pump (model 975, Harvard Apparatus, South Natick, MA) with the rats placed in a horizontal acrylic restraint. A segment of calibrated PE tubing was inserted between the syringe and the epidural catheter, and the injection was monitored by observing the movement of a small air bubble within the tubing. The TF, PP, and MF tests were performed immediately before and after infusion, and animals receiving lidocaine solutions that failed to show the cutoff value of the TF latency after infusion were excluded from the study. Anesthetic level was also assessed immediately after infusion by clamping the skin of the hindlimbs or trunk. All the tests were repeated 4 days later.
Histological Examination
After the last functional examination, the rats were killed by intraperitoneal pentobarbital and perfused intracardially with a phosphate-buffered 2.0% paraformaldehyde–2.5% glutaraldehyde fixative. Methyl green solution was injected to confirm the location of the catheter after the perfusion. The spinal cord and nerve roots were dissected out and immersed in the same fixative for 5 h. Two specimens (10 mm rostal and caudal to the conus medullaris) from each rat were postfixed with cacodylate-buffered 1% osmium tetroxide, dehydrated in a series of graded alcohol solutions, and embedded in epoxy resin. From the embedded tissue, 1-µm transverse sections were obtained using the microtome (MT6000; RMC, Tucson, AZ) and stained with toluidine blue dyes. Histopathologic evaluation was performed using light microscopy by a pathologist blinded to the group assignment and to the results of behavioral measurements. Sections obtained from 10 mm rostral to the conus (caudal spinal cord) were used for qualitative evaluation. Quantitative analysis of nerve injury was performed using the sections obtained from 10 mm caudal to the conus (cauda equina). Each fascicle present in the cross section was assigned an injury score of 0–3 (where 0 = normal, 1 = mild, 2 = moderate, and 3 = severe) as described previously (3,17–20). The injury score for each rat was then calculated as the average score of all fascicles in the cross section. In addition, ultrathin sections were obtained using the same microtome described above and double-stained with uranyl acetate and lead citrate for electron microscopy (EM-002B, Topcon, Tokyo, Japan).
Statistical Analysis
Sample size was determined by a power analysis based on the variability (sd 0.3) observed in our previous study (7) and an expected difference of 0.7 in nerve injury score with ß set at 0.2 and 
at 0.05. A minimal sample of five in each group met these criteria. To assess whether the groups studied were equivalent before administration of the test solutions, raw baseline latencies were compared using Student’s t-test and one-way analysis of variance for experiments 1 and 2, respectively. To assess sensory function 4 days after infusion, TF latencies and PP thresholds were compared with baseline values using the paired t-test. In addition, both values obtained 4 days after infusion were converted to the percent maximal possible effect, calculated as (postdrug value – baseline value)/(cutoff value – baseline value) x 100 and analyzed using Student’s t-test or one-way analysis of variance followed by the Scheffé test for experiments 1 or 2, respectively. MF scores were compared using the Kruskal–Wallis test followed by the Mann–Whitney U-test. Injury scores were compared using the Mann–Whitney U-test or the Kruskal–Wallis test followed by the Mann–Whitney U-test for Experiments 1 or 2, respectively. P < 0.05 was considered significant.
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RESULTS
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In Experiment 1, one rat given 2% lidocaine was excluded from the study because postmortem examination revealed that the catheter was located in the subarachnoid space. The remaining 12 rats (six in each group) were included in the data analysis. Baseline TF latencies and PP thresholds did not differ between groups. TF latency did not change in rats given saline throughout the experiment but significantly increased in those given 2% lidocaine when assessed 4 days after infusion (Fig. 1). Neither group of rats developed a persistent increase in PP threshold or MF scores. Sections obtained from the cauda equina of rats given saline showed little damage in the fascicles, whereas those in group L contained mild to moderate damage, including edema and axonal degeneration with swelling in the nerve roots. Nerve injury scores were significantly different between groups (Fig. 2). Sections obtained from the spinal cord of rats in both groups showed no apparent abnormal findings but minimal axonal degeneration in the posterior columns in some animals given lidocaine.
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Figure 1. Sensory function 4 days after epidural infusion of 2% lidocaine (n = 6) and normal saline (n = 6). Tail-flick latencies were converted to the percent maximal possible effect (%MPE), calculated as (postdrug value – baseline value)/(cutoff value – baseline value) x 100. Data are presented as mean ± sem *P < 0.05 compared with saline.
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Figure 2. Nerve injury score for sections obtained 10 mm caudal to the conus 4 days after epidural infusion of 2% lidocaine (n = 6) and normal saline (n = 6). Each fascicle present in the cross section was assigned an injury score of 0–3 (0 = normal, 1 = mild, 2 = moderate, and 3 = severe). The injury score for each rat was then calculated as the average score of all fascicles in the cross section. The box represents the 25th–75th percentiles, and the median is represented by the solid line. Error bars above and below the box mark the 10th and 90th percentiles. *P < 0.05 compared with saline.
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In Experiment 2, four rats (one in each group of rats receiving lidocaine) were excluded from the study because postmortem examination revealed that the catheter was located in the subarachnoid space, and one rat in group L30 that failed to develop anesthesia during infusion was not studied. All the other animals had the catheter in the epidural space after perfusion and were included in the data analysis. Baseline TF latencies and PP thresholds did not differ among groups. TF latency percent maximal possible effect did not change in rats given saline throughout the experiment. In contrast, when assessed 4 days after infusion, TF latencies of rats in group L120 increased significantly from baseline, and the increase in group L120 differed significantly from that in groups S120 and L30 (Fig. 3). No group of rats developed a persistent increase in PP threshold or MF scores.
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Figure 3. Sensory function 4 days after an epidural infusion of 5% lidocaine for 15 min (L15, n = 8), 30 min (L30, n = 7), 60 min (L60, n = 7), and 120 min (L120, n = 7), or normal saline for 120 min (S120, n = 7). Tail-flick latencies were converted to the percent maximal possible effect (% MPE), calculated as (postdrug value – baseline value)/(cutoff value – baseline value) x 100. Data are presented as mean ± sem. *P < 0.05 compared with saline.  P < 0.05 compared with baseline.
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Sections obtained from the cauda equina of rats given saline showed little damage in the fascicles, whereas those in groups L30, L60, and L120 contained mild to moderate injury in the nerve roots. Histologic changes in nerve roots were characterized by edema and axonal degeneration, including appearance of swelling and atrophy (Fig. 4). Nerve injury scores for groups L30, L60, and L120 differed significantly from those for group S120 (Fig. 5). A significant difference in damage in nerve roots was also observed among rats given the anesthetic for different durations of time; nerve injury scores for group L120 were higher than those for groups L15 and L30, and injury in group L60 was greater than in group L15. Qualitative light microscopic examination revealed no apparent spinal cord injury in any group, but minimal axonal degeneration in the posterior columns in some animals given lidocaine for 60 and 120 min.
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Figure 4. Transverse sections obtained from 10 mm caudal to the conus 4 days after a 2-h epidural infusion of saline (A) or 5% lidocaine. (B) Arrows indicate damaged fascicles in cauda equina. Histologic changes in cauda equina were characterized by edema and axonal degeneration, including appearance of myelin ovoid, and swelling, atrophy, and loss of axons with macrophage infiltration.
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Figure 5. Nerve injury score for sections obtained 10 mm caudal to the conus 4 days after a epidural infusion of 5% lidocaine for 15 min (L15, n = 8), 30 min (L30, n = 7), 60 min (L60, n = 7), and 120 min (L120, n = 7), or normal saline for 120 min (S120, n = 7). Each fascicle present in the cross section was assigned an injury score of 0–3 (where 0 = normal, 1 = mild, 2 = moderate, and 3 = severe). The injury score for each rat was then calculated as the average score of all fascicles in the cross section. The box represents the 25th–75th percentiles, and the median is represented by the solid line. Error bars above and below the box mark the 10th and 90th percentiles. *P < 0.05 compared with saline. P < 0.05 compared with L15. #P < 0.05 compared with L30.
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Typical electron microscopic findings are shown in Figure 6. In the specimen from a rat in group S120, axons and myelin lamellae of myelinated fibers were almost intact. Unmyelinated fibers were also almost intact with clear neurofilaments, neurotubules, and mitochondria. In contrast, the specimen from a rat in group L120 included axonal degeneration, disintegrated myelin lamellae, and myelin ovoid. In addition, unmyelinated swollen fibers, unclear neurofilaments, and neurotubules, as well as degenerated mitochondria and degenerated Schwann sheaths were present.
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Figure 6. Typical electron microscopic findings of myelinated and unmyelinated fibers obtained 4 days after a 2-h epidural infusion of saline (A) or 5% lidocaine; (B) original magnification x 12,000. The arrows indicate unmyelinated fibers. Normal myelinated and unmyelinated fibers were observed in A. There were degenerated axons and disintegrated myelin lamellae in B. Unmyelinated fibers in B were swollen with unclear neurofilaments and neurotubules, and degenerated mitochondria. M = myelinated fiber.
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DISCUSSION
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To the best of our knowledge, this is the first study to observe the neurotoxicity of local anesthetics administered in the epidural space. In our previous study (7), a bolus injection of 100 µL of 10% lidocaine (10 mg) did not induce detectable neurologic injury. In the present study, the first experiment was conducted to determine whether a larger dose of epidural lidocaine had neurotoxic effects. The dosage was increased by using an infusion technique rather than a bolus injection of a larger concentration and/or volume of local anesthetic, since a concentration larger than 10% lidocaine would have further decreased clinical relevance of the study, and a bolus injection of a volume larger than 100 µL might not have been tolerable to our animal model. In our previous experiments (7), a single epidural injection of 100 µL caused severe excitation in some animals. The continuous infusion also has the advantage of producing consistent and restricted anesthetic distribution, which has been considered one of the causative factors to produce clinical injury (21). As a result, although the total dose of lidocaine was only 1.2 times as large as that used in our previous study (7), functional impairment and morphologic damage were observed with 2% lidocaine administered for 120 min at a rate of 5 µL/min (12 mg) into the epidural space.
The results of Experiment 2 confirmed that the neurotoxicity of epidural lidocaine was dose dependent. In this experiment, 5% lidocaine was used rather than 2%, since the neurotoxic effects observed in the first experiment were mostly mild. The observation of severe nerve injury was necessary for a demonstrating dose–effect relationship and required larger doses of lidocaine, which might also have been administered by increasing either the rate or duration of infusion. However, an increase in the rate could have increased the rostal spread of the anesthetic and induced more hemodynamic and respiratory changes. A longer infusion was not feasible as well, since placing an animal in a restraint for longer than 2 h could cause undue stress.
In the current study, we advanced epidural catheters in the caudal direction from the L4–5 intervertebral space to position the tip very close to the caudal end of the epidural space. There is substantial clinical and experimental evidence to suggest that maldistribution of local anesthetic is an important etiologic factor in neurologic injury, and our technique was specifically developed to produce a restricted distribution. Previous studies showing the neurotoxic effects of intrathecal local anesthetics have used rat models with the tip of an intrathecal catheter placed among the nerve roots of the cauda equina. Although the location of our epidural catheter was checked only after perfusion, it is likely that our epidural rat model had the tip of the catheter at a similar level to those intrathecal models during infusion.
Histological findings in the present study indicated that epidurally administered lidocaine induced damage mainly to the fascicles and less to the spinal cord. These are also common features observed in the neurologic injury with intrathecal local anesthetic (3,7,22). Thus, it appears that epidural local anesthetic may induce neurologic injury with a part of the anesthetic absorbed into the cerebrospinal fluid. However, the present findings do not exclude a possibility that there are also other mechanisms involved in epidural local anesthetic neurotoxicity, especially when a drug is administered at a more cephalad level.
It is possible that ischemia was responsible for the damage observed. However, since anesthetic level was restricted to the perineum in most animals and few rats developed level of blockade higher than that in the hindlimbs, it is unlikely that those rats suffered severe hypotension leading to hypoperfusion in the spinal nerves. In fact, preliminary data we have obtained from tail cuff blood pressure monitoring suggest that hemodynamics remain stable under the current experimental conditions.
There are some limitations in the study. First, the doses of lidocaine associated with injury in both experiments exceed, on a per kg basis, commonly administered clinical doses. In addition, the concentration used in the second experiment was 2.5 times larger than the largest concentration used for clinical epidural anesthesia. However, available data suggest that rats and humans differ in the dose–response relationship for epidural lidocaine. Fassoulaki et al. (23) found, using an intrathecal model, that rats required far larger doses (1.2 mg/kg) of lidocaine to develop sacral sensory anesthesia than do humans (0.3–0.6 mg/kg). Epidural administration requires larger dosage of local anesthetic than intrathecal administration, not only in humans but also in rats. Results of our previous study (7) using rats showed that the potency ratio was 4.72:1 for intrathecal:epidural lidocaine. Second, it is possible, although unlikely, that residual lidocaine could contribute to the functional impairment present 4 days after administration. However, the histological damage observed in the specimens from rats clearly indicates that epidural lidocaine can cause permanent nerve injury.
Clinical reports have shown far less frequent neurologic complications after epidural anesthesia than spinal (5,6), and previously available laboratory evidence (7) suggests that there is a significant difference in neurotoxicity of local anesthetic administered between epidural and intrathecal spaces. However, although we must be careful to extrapolate from laboratory data to clinical practice, the present findings suggest that patients may not be immune to neurologic injury when epidural lidocaine is administered in large doses.
In conclusion, when a large dose of lidocaine was administered continuously into the epidural space, persistent functional impairment occurred with histologic damage in the nerve roots in a dose-dependent fashion.
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ACKNOWLEDGMENTS
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The authors thank Toshiko Tsumori, PhD, (Assistant Professor, Department of Morphological Neuroscience, Shimane University, Izumo City, Japan) for neuropathologic assessment and Tsunao Yoneyama and Yuko Okui (Technicians, Center for Integrated Research in Science, Shimane University, Izumo City, Japan) for technical assistance.
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Footnotes
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Accepted for publication June 12, 2006.
Presented in part at the Annual Meeting of the American Society of Anesthesiologists, Atlanta, Georgia, October 22–26, 2005.
Supported by Japan Society for the Promotion of Science Grant 15591630.
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Abstract
Introduction
