• View in gallery
    Figure 1—

    Schematic representation of stimulating and recording electrode placements (ventral aspect of wing) for recording sensory evoked potentials and SNCVs from the radial and ulnar nerves in a mallard duck (Anas platyrhynchos).

  • View in gallery
    Figure 2—

    Representative recordings of the CNAP and CDP of the ulnar nerve following distal sensory nerve stimulation in an anesthetized mallard duck before (A) and after (B) surgical transection of the nerve branches of the brachial plexus. Wing temperature was 35.8°C during both assessments. Following nerve transection, the ulnar CNAP (A1 in both panels) is preserved but the CDP (A2 in both panels) is abolished. All horizontal divisions marked by dots (time) represent 2-millisecond intervals. For each tracing, stimulus intensity (mA), amplitude calibration (μV/division), and number of signals (N) averaged are stated.

  • 1.

    Niederhauser UB, Holiday TA. Electrodiagnostic studies in diseases of muscles and neuromuscular junctions. Semin Vet Med Surg (Small Anim) 1989;4:116125.

    • Search Google Scholar
    • Export Citation
  • 2.

    Holliday TA. Electrodiagnostic examination: somatosensory evoked potentials and electromyography. Vet Clin North Am Small Anim Pract 1992;22:833857.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 3.

    Holliday TA, Ealand BG, Weldon NE. Sensory nerve conduction velocity: technical requirements and normal values for branches of the radial and ulnar nerves of the dog. Am J Vet Res 1977;38:15431551.

    • Search Google Scholar
    • Export Citation
  • 4.

    Cuddon PA, Delauche AJ, Hutchison JM. Assessment of dorsal nerve root and spinal cord dorsal horn function in clinically normal dogs by determination of cord dorsum potentials. Am J Vet Res 1999;60:222226.

    • Search Google Scholar
    • Export Citation
  • 5.

    Shell L, Richards M, Saunders G. Brachial plexus injury in two Red-tailed hawks (Buteo jamaicensis). J Wildl Dis 1993;29:177179.

  • 6.

    Shell L, Jortner BS, Ehrich M. Assessment of organophosphorus-induced delayed neuropathy in chickens using needle electromyography. J Toxicol Environ Health 1988;25:2133.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 7.

    Clippinger TL, Platt SR, Bennett RA, et al. Electrodiagnostic evaluation of peripheral nerve function in rheas and barred owls. Am J Vet Res 2000;61:469472.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 8.

    Gregory NG, Wooten SB. Effect of electrical stunning on somatosensory evoked potentials in chickens. Br Vet J 1989;145:159164.

  • 9.

    Necker R, Meinecke CC. Conduction velocities and fiber diameters in a cutaneous nerve of the pigeon. J Comp Physiol [A] 1984;154:817824.

  • 10.

    Necker R. Projection of a cutaneous nerve to the spinal cord of the pigeon I. Evoked field potentials. Exp Brain Res 1985;59:338343.

  • 11.

    Woodbury CJ. Physiological studies of cutaneous inputs to dorsal horn laminae I–IV of adult chickens. J Neurophysiol 1992;67:241254.

  • 12.

    Mazliah J, Barron S, Bental E, et al. The effects of long-term lead intoxication on the nervous system of the chicken. Neurosci Lett 1989;101:253257.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 13.

    Purser DA, Berrill KR, Majeed SK. Effects of lead exposure on peripheral nerve in the cynomolgus monkey. Br J Ind Med 1983;40:402412.

  • 14.

    Tseng HP, Wang YH, Wu MM, et al. Association between chronic exposure to arsenic and slow nerve conduction velocity among adolescents in Taiwan. J Health Popul Nutr 2006;24:182189.

    • Search Google Scholar
    • Export Citation
  • 15.

    Yokoyama K, Feldman RG, Sax DS, et al. Relation of distribution of conduction velocities to nerve biopsy findings in n-hexane poisoning. Muscle Nerve 1990;13:314320.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 16.

    Dorfman LJ, McGill KC, Cummins KL. Electrical properties of commercial concentric EMG electrodes. Muscle Nerve 1985;8:18.

  • 17.

    Nickel R. Nerves of the spinal cord. In: Nickel R, Schummer A, Seiferle E, eds. Anatomy of the domestic birds. New York: Springer-Verlag Inc, 1977;131143.

    • Search Google Scholar
    • Export Citation
  • 18.

    International Species Inventory System. Physiologic data references values for Anas platyrhynchos. Eagan, Minn: International Species Inventory System, 2002.

    • Search Google Scholar
    • Export Citation
  • 19.

    Redding RW, Ingram JT. Sensory nerve conduction velocity of cutaneous afferents of the radial, ulnar, peroneal, and tibial nerves of the cat: reference values. Am J Vet Res 1984;45:10421045.

    • Search Google Scholar
    • Export Citation
  • 20.

    Necker R. Thermal sensitivity of different skin areas in pigeons. J Comp Physiol 1977;116:239246.

  • 21.

    Bagley RS, Wheeler SJ, Gay JM. Effects of age on temperaturerelated variation in motor nerve conduction velocity in healthy chickens. Am J Vet Res 1995;56:819821.

    • Search Google Scholar
    • Export Citation
  • 22.

    Kornegay JN, Gorgacz EJ, Parker MA, et al. Motor nerve conduction velocity in normal chickens. Am J Vet Res 1983;44:15371540.

  • 23.

    Lee AF, Bowen JM. Effect of tissue temperature on ulnar nerve conduction velocity in the dog. Am J Vet Res 1975;36:13051307.

  • 24.

    Wheeler SJ. Influence of limb temperature on sensory nerve conduction velocity in horses. Am J Vet Res 1989;50:18171819.

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Somatosensory evoked potentials and sensory nerve conduction velocities in the thoracic limb of mallard ducks (Anas platyrhynchos)

Deena J. BrennerVeterinary Medical Teaching Hospital, University of California, Davis, CA 95616

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R. Scott LarsenDepartment of Medicine and Epidemiology, University of California, Davis, CA 95616

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Peter J. PascoeDepartment of Surgical and Radiological Sciences, University of California, Davis, CA 95616

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Raymund F. WackDepartment of Medicine and Epidemiology, University of California, Davis, CA 95616

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D. Colette WilliamsVeterinary Medical Teaching Hospital, University of California, Davis, CA 95616

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Peter J. DickinsonDepartment of Surgical and Radiological Sciences, University of California, Davis, CA 95616

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Abstract

Objective—To develop a clinically applicable technique for recording cord dorsum potentials (CDPs) following stimulation of the radial and ulnar nerves and establish reference values for radial and ulnar sensory nerve conduction velocities (SNCVs) in the wings of ducks.

Animals—8 clinically normal adult female mallard ducks (Anas platyrhynchos).

Procedures—Radial and ulnar compound nerve action potentials (CNAPs) and CDPs were recorded following distal sensory nerve stimulation. The CDPs were recorded from the interarcuate space between the last cervical vertebra and the first thoracic vertebra. Surgical dissection and transection of the brachial plexus in 1 anesthetized duck were performed to identify nerve root location and confirm functional loss of nerve conduction assessed by loss of the CDP.

Results—Radial and ulnar CNAPs and CDPs were consistently recorded in all birds. Median radial SNCV was 38.3 m/s (range, 36.0 to 49.0 m/s), and ulnar SNCV was 35.3 m/s (range, 28.0 to 40.0 m/s). Surgical transection of the brachial plexus resulted in complete loss of the CDP.

Conclusions and Clinical Relevance—Measurement of radial and ulnar SNCV or CDP is feasible in isoflurane-anesthetized mallard ducks. The CDP accurately reflects sensory nerve conduction through the brachial plexus. Assessment of brachial plexus function in mallard ducks via evaluations of SNCVs and CDPs may have application for diagnosis of traumatic injuries to the brachial plexus, evaluation of neuropathies associated with exposure to toxic chemicals, and assessment of the efficacy of interventions such as brachial plexus nerve blockade.

Abstract

Objective—To develop a clinically applicable technique for recording cord dorsum potentials (CDPs) following stimulation of the radial and ulnar nerves and establish reference values for radial and ulnar sensory nerve conduction velocities (SNCVs) in the wings of ducks.

Animals—8 clinically normal adult female mallard ducks (Anas platyrhynchos).

Procedures—Radial and ulnar compound nerve action potentials (CNAPs) and CDPs were recorded following distal sensory nerve stimulation. The CDPs were recorded from the interarcuate space between the last cervical vertebra and the first thoracic vertebra. Surgical dissection and transection of the brachial plexus in 1 anesthetized duck were performed to identify nerve root location and confirm functional loss of nerve conduction assessed by loss of the CDP.

Results—Radial and ulnar CNAPs and CDPs were consistently recorded in all birds. Median radial SNCV was 38.3 m/s (range, 36.0 to 49.0 m/s), and ulnar SNCV was 35.3 m/s (range, 28.0 to 40.0 m/s). Surgical transection of the brachial plexus resulted in complete loss of the CDP.

Conclusions and Clinical Relevance—Measurement of radial and ulnar SNCV or CDP is feasible in isoflurane-anesthetized mallard ducks. The CDP accurately reflects sensory nerve conduction through the brachial plexus. Assessment of brachial plexus function in mallard ducks via evaluations of SNCVs and CDPs may have application for diagnosis of traumatic injuries to the brachial plexus, evaluation of neuropathies associated with exposure to toxic chemicals, and assessment of the efficacy of interventions such as brachial plexus nerve blockade.

Electrodiagnostic examination is a fairly noninvasive method of assessing functional neuromuscular disorders in veterinary and human medicine. Routine electrodiagnostic examinations in domestic animals often include electromyography, motor and sensory nerve conduction evaluations, and assessment of SSEPs.1,2 Results of these diagnostic tests can help clinicians distinguish between myopathic and neuropathic (axonal or demyelinating) disease processes as well as differentiate between proximal and distal lesions within the peripheral nerves. Motor nerve conduction velocities can be determined by stimulating a minimum of 2 sites along a peripheral nerve and recording the compound muscle action potential (M wave) from a muscle innervated by that nerve.1 The SNCVs are determined by recording the conducted volley along a peripheral nerve following sensory nerve stimulation. The CNAP is a reflection of peripheral sensory nerve function.1,2 Cord dorsum potentials are stationary field potentials that are generated by pools of spinal cord interneurons in the lumbar and cervical intumescences. The presence of a CDP indicates that conducted sensory nerve action potentials from the brachial or lumbar plexus have reached the spinal cord.3,4 Despite the development of neuromuscular disorders in avian species, electrodiagnostic evaluations are not routinely performed, and to our knowledge, there is little information describing electrodiagnostic techniques in birds. Electromyography has been used to confirm the presence of brachial plexus avulsions in red-tailed hawks (Buteo jamaicensis)5 and assess organophosphate-induced delayed neuropathy in chickens.6 Reference values for ulnar MNCVs have been determined for rheas (Rhea americana) and barred owls (Strix varia),7 and cerebral SSEPs have been evaluated in studies8 to assess the efficacy of electrical stunning following stimulation of the radial nerve in chickens. Conduction velocities and evoked potentials have also been recorded following radial nerve stimulation and during open surgical laminectomy procedures in pigeons.9,10 Results of similar invasive experiments investigating evoked potentials and SNCVs following stimulation of the caudal cutaneous femoral nerve in chickens have been reported.11

The purpose of the study reported here was to develop a clinically applicable technique for recording CDPs following stimulation of the radial and ulnar nerves and establish reference values for radial and ulnar SNCVs in the wings of ducks. We hypothesized that CNAPs and CDPs could be consistently measured in birds following noninvasive stimulation of the radial and ulnar nerves and that the presence of the CDP would reflect the integrity of the brachial plexus. Mallard ducks (Anas platyrhynchos) were chosen as the model species for evaluating these techniques because of their size, tractability, and wing anatomy. It was anticipated that the SNCV and CDP values could potentially be used for assessment of brachial plexus integrity and function in mallard ducks and may be clinically applicable in the diagnosis of traumatic injuries to the brachial plexus; evaluation of toxic neuropathies associated with chemicals such as lead,12,13 arsenic,14 and n-hexane15; or assessment of the efficacy of therapeutic interventions such as a brachial plexus nerve blockade in this avian species.

Materials and Methods

Ducks—Eight adult female mallard ducks (Anas platyrhynchos) that weighed 0.9 to 1.1 kg were obtained from a commercial breeder. Physical examination, CBC, and plasma biochemical analyses were performed on each duck to evaluate its health status prior to study commencement. The study was approved by the Institutional Care and Use Committee of the University of California, Davis (protocol No. 06-12209).

Anesthesia and instrumentation—Each duck was manually restrained, and anesthesia was induced with 5% isofluranea administered via a face mask. Once relaxation occurred, each duck was intubated with an uncuffed 3.0- or 3.5-mm-diameter endotracheal tube and maintained via inhalation of 1.5% to 3.5% isoflurane for the duration of the procedure. A capnometerb was attached to the endotracheal tube, and end-tidal CO2 concentration was measured. Intermittent positivepressure ventilation was instituted to maintain end-tidal CO2 values at 25 to 45 mm Hg. Core body temperature was measured by use of an esophageal thermometer.c A circulating warm water blanket and external heat lamp were used to maintain body temperature at 37° to 39°C. Heart rate was monitored by use of a 9.7-Hz adult flat Doppler probed placed over the medial metatarsal artery. A cuff with a width approximately equal to 40% of the pelvic limb circumference was placed proximal to the Doppler probe and attached to a manual sphygmomanometere for measurement of systolic arterial blood pressure. Depth of anesthesia was monitored on the basis of a combination of clinical signs (eg, muscle tone and withdrawal reflexes) and autonomic responses (heart rate and arterial blood pressures), and alterations in the concentration of inhalant anesthetic were made accordingly.

Experimental design—In 5 of the 8 birds, both the left and right wings were evaluated (a washout period of at least 2 weeks was allowed to elapse between evaluations); in 3 birds, only 1 wing was evaluated. The single test wing or order of wing testing was determined by stratified random selection.

Each bird was placed in lateral recumbency with the test wing fully extended and suspended from an elevated clamp. This provided access to the entire ventral aspect of the wing. Pairs of polytetrafluoroethylenecoated stainless steel monopolar electrodesf were used for both stimulating and recording purposes; tip exposure was 5 mm for all electrodes. An additional platinum subdermal needle electrodeg was used as a ground electrode and inserted SC between the radius and ulna at a point halfway between the stimulating and recording electrodes. A surface temperature probef was positioned on the wing just cranial to the ground electrode. Wing temperature was maintained at 32° to 36°C by use of an infrared heat lamp.

Prior to each procedure, electrodes were sterilized in boiling deionized water for a minimum of 20 minutes. Electrodes were then electrolytically cleaned to reduce electrode impedance16 to ≤ 10 kΩ. All stimulating and recording electrodes were 20 mm in length except the cord dorsum recording electrode, which was 25 mm in length.

Two stimulating electrodes for the radial nerve were placed percutaneously at the cranial edge of the wing near the center of the carpometacarpus; 1 electrode was placed dorsally and 1 electrode was placed ventrally beneath the secondary feathers (Figure 1). Two stimulating electrodes for the ulnar nerve were placed percutaneously at the caudal edge of the ventral aspect of wing near the center of the metacarpus. The cathode and anode of each pair were separated by a distance of 7 mm, and the electrode tips were positioned proximally with the electrodes parallel to one another. Each pair was connected to separate stimulator probes.h The stimulator probes were isolated by use of transformer isolation. The recording electrodes for the ulnar nerve were placed percutaneously, adjacent to the cutaneous ulnar vein near the medial aspect of the point of the elbow. The recording electrodes for the radial nerve were placed percutaneously, cranial to the point of the elbow approximately halfway between the caudal aspect of the elbow joint and the cranial edge of the wing. The reference electrodes were placed 7 mm cranial to the recording electrodes. Both sets of recording electrodes were oriented with the tips pointing proximally. Placement of the cord dorsum electrode was performed by palpating the depression between the last cervical vertebra and the first thoracic vertebra and gently inserting the needle into the interarcuate space. A reference electrode was positioned SC approximately 3 cm laterally adjacent to the cord dorsum electrode.

Figure 1—
Figure 1—

Schematic representation of stimulating and recording electrode placements (ventral aspect of wing) for recording sensory evoked potentials and SNCVs from the radial and ulnar nerves in a mallard duck (Anas platyrhynchos).

Citation: American Journal of Veterinary Research 69, 11; 10.2460/ajvr.69.11.1476

Stimulus intensity of 5 mA was used for both SNCVs and CDPs; a supramaximal stimulus was achieved at 7 Hz for 0.2 milliseconds by use of a rectangular stimulus pulse. At least 1,000 impulses were averaged. Distance between stimulation and recording sites on the surface of the skin was determined with a tape measure along the presumed course of the nerve. Latency was defined as the time between the stimulus onset and the first positive peak of the CNAP. Amplitudes for the radial and ulnar CNAPs and CDPs were determined by calculating the voltage difference between the first positive peak and the highest negative peak. Stimulus intensity, CNAP latency, CNAP amplitude, and CDP amplitude following sensory nerve stimulation were determined by use of a neurodiagnostic system.i The SNCV was calculated as the stimulus to recording site distance divided by the CNAP latency. Ducks were monitored for adverse effects for 24 hours after the procedures.

To assess the effect of brachial plexus transection on SSEPs, surgical exploration and open unilateral transection of all brachial plexus nerve roots were performed on 1 anesthetized duck. The duck was positioned in lateral recumbency with the test wing fully extended and suspended from an elevated clamp; after placement of electrodes as described, initial electrophysiology measurements were obtained. Without altering the position of the wing or electrodes, a 1.5-cm skin incision was made in the ventral axillary region. Via careful blunt dissection of the subcutaneous tissues, the brachial plexus and all 4 branches of the brachial plexus (ventral branches of the 2 most caudal cervical spinal nerves and 2 most cranial thoracic spinal nerves)17 were exposed and transected. Electrodiagnostic procedures were then repeated. The anesthetized duck was euthanatized via IV injection of pentobarbital sodium and phenytoin sodiumj following this procedure.

Statistical analysis—For ducks in which both wings were evaluated, electrophysiologic values derived from the 2 wings were averaged and the mean value was used for overall analysis; single values were used for analysis in birds in which only 1 wing was evaluated. Because of the small sample size, data sets were analyzed via nonparametric methods.k Association between each electrophysiologic measurement and wing temperature was evaluated by use of Spearman rank correlation coefficients. Ulnar SNCV was compared with radial SNCV by use of a Mann-Whitney U test. Results were considered significant at a value of P ≤ 0.05. Results are presented as median and range.

Results

Each duck was considered clinically normal on the basis of physical examination findings; results for a CBC and plasma biochemical analyses were within reference ranges for mallard ducks.18 Radial and ulnar CNAPs and CDPs were consistently recorded for all ducks (Table 1). There was no significant difference between the median radial SNCV (38.3 m/s; range, 36.0 to 49.0 m/s) and median ulnar SNCV (35.3 m/s; range, 28.0 to 40.0 m/s). No significant correlation was determined between SNCV and wing temperature in the temperature range tested (32.0° to 36.0°C).

Table 1—

Sensory nerve stimulation data (median [range]) for the radial and ulnar nerves in 8 mallard ducks (Anas platyrhynchos) and wing temperature conditions (median [range]) during assessments.

NerveSNCV (m/s)CNAP (μV)CDP amplitude (μV)Wing temperature (°C)
Radial38.3 (36.0 – 49.0)1.7 (0.9 – 6.3)6.9 (3.9 – 18.8)35.6 (34.0 – 36.6)
Ulnar35.3 (28.0 – 40.0)4.1 (2.3 – 12.9)10.6 (1.9 – 18.0)35.5 (34.6 – 36.7)

Dissection of the brachial plexus in 1 anesthetized duck clearly revealed the branches of the brachial plexus. Prior to surgery, initial electrophysiologic measurements including SNCV and CNAP values were obtained (radial SNCV, 49.0 m/s; ulnar SNCV, 40.0 m/s; radial CNAP amplitude, 1.5 μV; and ulnar CNAP amplitude, 12.9 μV). The initial CDP amplitudes were obtained from the radial nerve (3.9 μV) and from the ulnar nerve (18.0 μV) following stimulation. Following surgical transection of the branches of the brachial plexus, the CDP was no longer present following either radial nerve or ulnar nerve stimulation (0.0 μV); however, there was no substantial change in SNCV or CNAP amplitude (radial SNCV, 43.0 m/s; ulnar SNCV, 36.0 m/s; radial CNAP amplitude, 2.3 μV; and ulnar CNAP amplitude, 10.7 μV; Figure 2).

Figure 2—
Figure 2—

Representative recordings of the CNAP and CDP of the ulnar nerve following distal sensory nerve stimulation in an anesthetized mallard duck before (A) and after (B) surgical transection of the nerve branches of the brachial plexus. Wing temperature was 35.8°C during both assessments. Following nerve transection, the ulnar CNAP (A1 in both panels) is preserved but the CDP (A2 in both panels) is abolished. All horizontal divisions marked by dots (time) represent 2-millisecond intervals. For each tracing, stimulus intensity (mA), amplitude calibration (μV/division), and number of signals (N) averaged are stated.

Citation: American Journal of Veterinary Research 69, 11; 10.2460/ajvr.69.11.1476

One duck died during recovery from anesthesia, likely as a result of endotracheal tube obstruction with thick tracheal secretions. Gross necropsy of this bird and histologic examination of tissues did not reveal any additional abnormalities. No adverse effects were observed in any other ducks during the 24-hour period following these procedures.

Discussion

Radial and ulnar SNCVs were consistently measured in mallard ducks in the present study. Values for SNCV in the mallard ducks were much lower than those previously reported for dogs (mean ± SD radial SNCV, 61.7 ± 0.76 m/s; ulnar SNCV, 68.4 ± 2.71 m/s)3 and cats (radial SNCV, 83.6 ± 8.3 m/s; ulnar SNCV, 89.2 ± 10.3 m/s).19 Sensory nerve conduction velocities in other avian species are also reported to be slower than those recorded for mammals; cutaneous radial SNCV in pigeons was 50 m/s,9 and cut femoral SNCV in chickens was 43 m/s.11 Conduction velocity is affected by nervefiber diameter and the thickness of the myelin sheath. In pigeons, the cutaneous antebrachii lateralis nerve (cutaneous branch of the radial nerve) has a smaller nerve diameter and a thinner myelin sheath (as determined via electron microscopy) than the corresponding nerves in mammalian species of similar or larger body sizes; similar nerve characteristics in mallard ducks and other avian species may explain the comparatively slower SNCVs recorded in the present and previous studies.9,11 Further research is warranted to evaluate the relationship between these electrophysiologic findings and the anatomic or morphometric characteristics of sensory nerves in mallard ducks, compared with other avian species and mammals.

In the ducks of the present study, the CDP was consistently recorded. When the brachial plexus was surgically transected in 1 bird, there was complete loss of the CDP, yet the radial and ulnar SNCV and CNAP values changed minimally. These results indicate that the CDP accurately reflects sensory nerve conduction through the brachial plexus in mallard ducks.

Efforts were made to maintain core body and wing temperatures at constant values throughout the electrodiagnostic evaluations. Wing temperatures in the ducks of the present study were consistent with wing temperatures among healthy pigeons.9,20 No significant correlation between wing temperature and radial or ulnar SNCV in mallard ducks was evident in our study. In other investigations, linear relationships between MNCV and limb temperature in juvenile chickens21 and between tibial MNCV and cloacal temperature in healthy adult chickens22 were identified. Linear associations between ulnar MNCV and tissue temperature in dogs23 and between lateral palmar SNCV and limb temperature in horses24 have also been determined. However, the 4 aforementioned studies investigated variation in conduction velocity over a wider range of temperatures. In the present study, wing temperature was maintained within a narrow temperature range (32° to 36°C) to maximize the consistency in electrophysiologic measurements among birds and over time. It is likely that any correlation between temperature and SNCV would be difficult to detect because of the narrow temperature range and the relatively low number of ducks. Additional studies involving electrophysiologic measurements over a wider range of wing temperatures would be necessary to determine whether similar temperature-dependent variation in SNCV is present in avian sensory nerves.

Female ducks were obtained for use in our study in an attempt to minimize aggression in a group housing setting. Differences between males and females of this species are expected to be minimal. To the authors' knowledge, data from males and females were pooled in previous avian electrophysiologic studies,6–10,20–22 and there are no reports of significant differences in electrophysiologic measurements obtained from male and female birds. Although variability in conduction velocities and action potential amplitudes was evident among individual birds, radial and ulnar SNCV, CNAP, and CDP values were consistently obtained in our study and may serve as baseline values for adult mallard ducks. The procedures were tolerated well with no apparent adverse effects. The technique used in our study has potential for use in the assessment of brachial plexus function and evaluation of peripheral neuropathies associated with toxic chemicals in birds, and it may have clinical applicability in assessments of the efficacy of brachial plexus nerve blockade in avian species.

ABBREVIATIONS

CDP

Cord dorsum potential

CNAP

Compound nerve action potential

MNCV

Motor nerve conduction velocity

SNCV

Sensory nerve conduction velocity

SSEP

Somatosensory evoked (spinal cord) potential

a.

Isoflo, Abbott Animal Health, Abbott Park, Ill.

b.

Tidal wave 715, Respironics, Carlsbad, Calif.

c.

Yellow Springs 700 Thermistor, Measurement Specialties, Dayton, Ohio.

d.

Parks Medical Electronics, Aloha, Ore.

e.

Speidel & Keller, Jungingen, Germany.

f.

Nicolet Biomedical, Madison, Wis.

g.

Grass-Telefactor, West Warwick, RI.

h.

Nicolet S403 stimulator probes, Nicolet Biomedical, Madison, Wis.

i.

Nicolet Viking IV EMG/Evoked Potential System, Nicolet Biomedical, Madison, Wis.

j.

Beuthanasia, 1 mL/kg, IV, Schering-Plough Animal Health, Kenilworth, NJ.

k.

Graphpad Instat, version 3.06, 32 bit for Windows, GraphPad Software Inc, San Diego, Calif.

References

  • 1.

    Niederhauser UB, Holiday TA. Electrodiagnostic studies in diseases of muscles and neuromuscular junctions. Semin Vet Med Surg (Small Anim) 1989;4:116125.

    • Search Google Scholar
    • Export Citation
  • 2.

    Holliday TA. Electrodiagnostic examination: somatosensory evoked potentials and electromyography. Vet Clin North Am Small Anim Pract 1992;22:833857.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 3.

    Holliday TA, Ealand BG, Weldon NE. Sensory nerve conduction velocity: technical requirements and normal values for branches of the radial and ulnar nerves of the dog. Am J Vet Res 1977;38:15431551.

    • Search Google Scholar
    • Export Citation
  • 4.

    Cuddon PA, Delauche AJ, Hutchison JM. Assessment of dorsal nerve root and spinal cord dorsal horn function in clinically normal dogs by determination of cord dorsum potentials. Am J Vet Res 1999;60:222226.

    • Search Google Scholar
    • Export Citation
  • 5.

    Shell L, Richards M, Saunders G. Brachial plexus injury in two Red-tailed hawks (Buteo jamaicensis). J Wildl Dis 1993;29:177179.

  • 6.

    Shell L, Jortner BS, Ehrich M. Assessment of organophosphorus-induced delayed neuropathy in chickens using needle electromyography. J Toxicol Environ Health 1988;25:2133.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 7.

    Clippinger TL, Platt SR, Bennett RA, et al. Electrodiagnostic evaluation of peripheral nerve function in rheas and barred owls. Am J Vet Res 2000;61:469472.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 8.

    Gregory NG, Wooten SB. Effect of electrical stunning on somatosensory evoked potentials in chickens. Br Vet J 1989;145:159164.

  • 9.

    Necker R, Meinecke CC. Conduction velocities and fiber diameters in a cutaneous nerve of the pigeon. J Comp Physiol [A] 1984;154:817824.

  • 10.

    Necker R. Projection of a cutaneous nerve to the spinal cord of the pigeon I. Evoked field potentials. Exp Brain Res 1985;59:338343.

  • 11.

    Woodbury CJ. Physiological studies of cutaneous inputs to dorsal horn laminae I–IV of adult chickens. J Neurophysiol 1992;67:241254.

  • 12.

    Mazliah J, Barron S, Bental E, et al. The effects of long-term lead intoxication on the nervous system of the chicken. Neurosci Lett 1989;101:253257.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 13.

    Purser DA, Berrill KR, Majeed SK. Effects of lead exposure on peripheral nerve in the cynomolgus monkey. Br J Ind Med 1983;40:402412.

  • 14.

    Tseng HP, Wang YH, Wu MM, et al. Association between chronic exposure to arsenic and slow nerve conduction velocity among adolescents in Taiwan. J Health Popul Nutr 2006;24:182189.

    • Search Google Scholar
    • Export Citation
  • 15.

    Yokoyama K, Feldman RG, Sax DS, et al. Relation of distribution of conduction velocities to nerve biopsy findings in n-hexane poisoning. Muscle Nerve 1990;13:314320.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 16.

    Dorfman LJ, McGill KC, Cummins KL. Electrical properties of commercial concentric EMG electrodes. Muscle Nerve 1985;8:18.

  • 17.

    Nickel R. Nerves of the spinal cord. In: Nickel R, Schummer A, Seiferle E, eds. Anatomy of the domestic birds. New York: Springer-Verlag Inc, 1977;131143.

    • Search Google Scholar
    • Export Citation
  • 18.

    International Species Inventory System. Physiologic data references values for Anas platyrhynchos. Eagan, Minn: International Species Inventory System, 2002.

    • Search Google Scholar
    • Export Citation
  • 19.

    Redding RW, Ingram JT. Sensory nerve conduction velocity of cutaneous afferents of the radial, ulnar, peroneal, and tibial nerves of the cat: reference values. Am J Vet Res 1984;45:10421045.

    • Search Google Scholar
    • Export Citation
  • 20.

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Contributor Notes

Dr. Brenner's present address is the Department of Veterinary Services, San Diego Zoo's Wild Animal Park, Escondido, CA 92027.

Supported by the Center for Companion Animal Health, School of Veterinary Medicine, University of California, Davis.

Presented in part in abstract form at the American Association of Zoo Veterinarians Conference, Knoxville, Tenn, October 2007.

The authors thank Mr. John Doval for the technical artwork.

Address correspondence to Dr. Brenner.