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A method for chronically recording brain-stem and cortical auditory evoked potentials from unanesthetized mice
78
Electroencephalograph), and clinical Neurophysiology, 185, 6 0 : 7 8 - 8 3 Elsevier Scientific Publishers Ireland, Ltd.
Short communication A METHOD FOR CHRONICALLY RECORDING BRAIN-STEM EVOKED POTENTIALS FROM UNANESTHETIZED MICE i
AND CORTICAL
AUDITORY
T.M. W E L C H , M.W. C H U R C H and D.W. S H U C A R D 2
Brain Sciences Laboratories, National Jewish Hospital and Research Center, and Department of Psychiatry, University of Colorado School of Medicine, Denver, CO (U.S.A.) (Accepted for publication: August 24, 1984)
The brain-stem auditory evoked potential (BAEP), a shortlatency far-field sensory evoked potential, has become an important clinical and experimental tool for assessing the integrity of the subcortical auditory pathway in h u m a n s and animals. The cortical auditory evoked potential (CAEP), a long-latency sensory evoked potential, is widely used to investigate sensory and cognitive processing of auditory information at the cortical level. Animal investigations offer the opportunity to study the BAEP and CAEP under well controlled conditions. Of particular significance when recording evoked potentials is the state of the animal being studied. Animal BAEP recordings are most often done under sedation or anesthesia. Although anesthesia affords control over artifacts due to movement or muscle activity, the use of anesthesia also leads to unwanted side effects such as hypothermia. In our experience, anesthetic-induced hypothermia readily occurs in small animals such as the laboratory mouse. Further, relatively small temperature decreases significantly affect the BAEP (Stockard et al. 1978a; Marshall and Donchin 1981; Williston and Jewett 1982). It also has been shown that anesthesia and sedation may affect the latencies a n d / o r amplitudes of both the BAEP (Squires et al. 1978; Stockard et al. 1978b; Henry 1979; Church and Williams 1982; Sutton et al. 1982; Church et al. 1984a, b) and the C A E P (Brazier 1972; Sutton et al. 1982) even in thermoregulated animals. Moreover, use of an anesthetic may interact with or mask the clinical and experimental conditions under investigation. Based on the above, it may be more appropriate to record BAEPs and CAEPs without the use of anesthesia. In the present report, we describe a technique for obtaining BAEPs and CAEPs in unanesthetized mice. The technique is useful for investigating the CNS effects of pharmacological agents, pathological conditions, aging and development without the possible confounding effects of anesthesia.
1 Supported in part by N I M H Grant No. M H 15442 and N1H Grant No. N S / A M 15855. 2 Address reprint requests to: D.W. Shucard, Ph.D., Brain Sciences Laboratories, National Jewish Hospital, 3800 East Colfax Ave., Denver, CO 80206, U.S.A.
Method Subjects The subjects were 9 BDF 1 female mice weighing 25 29 g before surgery with ages ranging from 4 to 6 months. These animals were bred in our animal facilities, housed individually in plexiglass cages (26 c m x 1 6 c m x l 3 cm), kept on a 12 h light/dark cycle, and fed lab chow and water ad lib.
Surgery All subjects were anesthetized with 80 m g / k g (i.p.) of sodium pentobarbital and secured on a small operating table. After a midline scalp incision was made, the skin and connective tissue were scraped away from the skull surface. Muscle and other soft tissue were cauterized to retard tissue regrowth. Once the surrounding tissue was cleared away, the skull surface was cleaned with 70% ethanol. Burr holes were then drilled into the skull to accommodate 3 stainless steel skull screws (0-80× 1 / 8 " stainless steel, flathead) that were placed in the following configuration: two screws, one on each side, were placed 2 m m lateral to the midline (sagittal suture) and one-half the distance between bregma and lambda. The third screw was placed 1.5 m m posterior to and 1.5 m m lateral to the left of lambda. When inserting the skull screws, care was taken not to cause brain compression or penetration of the dura. Once the skull screws were in place, male gold pins, previously connected to each of the screws via a teflon-coated silver wire (0.02 m m diameter), were inserted into a plastic strip connector cut to a length of 0.8 cm and a height of 0.5 cm. Each strip connector had 3 holes which accommodated the 3 male pin connectors, one connector attached to each of the wire leads from the 3 skull screws. Dental acrylic was then applied to the skull screws, skull surface and strip connector. Once a small base of dental acrylic was built, a female headpost was placed posterior to the strip connector and cemented into place with acrylic. The skull screws, strip connector and headpost were then all secured with dental acrylic. The headpost was machined from a no. 10-34 brass rod, had a length of 0.8 cm, and was drilled lengthwise and tapped to accommodate a no. 440 screw. The skull and surrounding tissue were then infused with an antibiotic ointment (bacitracin) and a single wound clip was used to partially
0013-4649/85/$03.30 © 1985 Elsevier Scientific Publishers Ireland, Ltd.
AEPs IN U N A N E S T H E T I Z E D MICE
A
79 close the incision. Animals were thermoregulated throughout the surgery and allowed 3-8 weeks to recover.
Adaptation
i
i i
After surgery but prior to evoked potential recording, the mice were gentled to reduce stress. This procedure involved handling the animals for 15 min every other day for a 2 week period. Adaptation to the restrainer and recording conditions was accomplished by placing each mouse in a mock recording setup for 1 h prior to the day of the actual experiment. The animal restrainer was also placed in each mouse cage for 1 night to further facilitate the animal's adaptation to the restrainer.
Apparatus The recording environment consisted of an animal restrainer mounted in a box placed inside an acoustic chamber. The animal restrainer was designed to restrict body movement and immobilize the animal's head. Fig. 1A illustrates the design of the restrainer. The animal restrainer was fastened to a floor bar in the box with a small C-clamp to prevent it from moving. Recording leads, suspended from the top of the box, connected the skull electrodes via the strip connector with the evoked potential recording system. The sides of the box were padded with 2.54 cm thick foam rubber for acoustical dampening. The box, containing the animal in its restrainer, was placed in an electrically shielded 104 cm x 96.5 cm × 81 cm acoustic chamber. Fig. 1B illustrates the position of the mouse after placement in the device during recording.
Stimuli
Fig. 1. A: design of the animal restrainer. The mouse restrainer consists of a body tube which encases and supports the animal's body. Inside the body tube is a backstop that can be adjusted to the animal's body length. A 0.5 cm slot, cut lengthwise along the bottom of the body tube, allows the experimenter to pull the animal by the tail into the restrainer. An adjustable neck collar, incorporated into the front of the restrainer, prevents the animal from withdrawing its head into the restrainer. A male screw, mounted on a sliding bar at the front and top of the restrainer, is screwed into the female headpost cemented on the animal's head. B: position of the mouse during recording. With the mouse's head extended past the opening of the restrainer, the collar (a) is adjusted around the mouse's neck and tightened into place. The male screw attached to the restrainer (b) is threaded into a female headpost (d) which is permanently affixed to the strip connector. The recording leads (e) are
Rarefaction 'click' stimuli were used to produce BAEP recordings, Click stimuli were created by activating an earphone with a 100 #sec square wave pulse generated by an Iconix system. The earphone was mounted directly in front of each animal at ear level and at a distance of 13.8 cm from the animal's ears. Click intensity was measured in dB SPL peak equivalent (p.e.) at the mouse's ears. Spectral energy of the click was broadly distributed below 4 kHz with maximal energy between 3.0 and 3.5 kHz. Paired tones were used to produce CAEP recordings. Each tone was 100 msec in duration with an intensity of 80 dB SPL at the mouse's ears. A 6 kHz tone was selected to evoke CAEPs because initial investigations in our laboratory indicated that this frequency produced robust and well-delineated mouse CAEPs.
Recordings The BAEP was recorded ipsilaterally between the skull screw electrode placed to the left and posterior to lambda (positive) and the screw electrode placed to the left of the sagittal suture (negative). The remaining screw electrode served attached to the strip connector (f). The animal's head is held steadfast by the male screw that descends from the top of the restrainer and is threaded into the female headpost cemented to the mouse's head (d).
80 as a ground. BAEPs were recorded from each animal as a function of parametric variations in click repetition rate and stimulus intensity. With respect to repetition rate, BAEPs were collected in response to clicks presented at rates of 10, 50 and 100/see (intensity = 90 dB SPL p.e.). With respect to intensity, BAEPs were collected in response to clicks presented at 30, 35, 40, 50, 70, 90 and 110 dB SPL p.e. (rate = 10 clicks/sec). Each animal's BAEPs were amplified by an amplifier with sensitivity set at 1 ~ V / m m and a bandpass of 300-3000 Hz. The amplified signals were averaged by a signal averager, with positivity displayed upwards, and printed out on an X-Y plotter. BAEP activity was sampled at a rate of 40 vsec per address and 256 responses were averaged. The analysis epoch of 10.24 msec (256 addresses) consisted of a 0.50 msec prestimulus and a 9.74 msec poststimuhis period. The 0.50 msec prestimulus period consisted of a 0.40 msec acoustic transit time and a 0.10 msec equipment delay. This prestimulus period allowed for adequate temporal separation of stimulus artifact from the first BAEP component. The CAEP was recorded contralaterally between the screw electrode placed to the left and posterior to lambda (negative) and the screw electrode placed to the right of the sagittal suture (positive). The remaining screw served as a ground. CAEPs to the 80 dB SPL paired tones were evaluated at inter-stimulus intervals of 2.0, 1.0 and 0.6 sec with a time interval of 4.0 sec between successive tone pairs. Each animal's CAEPs were amplified with sensitivity set at 3 0 / t V / m m and a bandpass of 3 - 3 0 Hz. The amplified signals were averaged with positivity displayed upwards and printed out via the X-Y plotter. CAEP activity was sampled at a rate of 1 msec per address and 32 responses were averaged. The analysis epoch of 512 msec (512 addresses) consisted of a 100 msec prestimulus and a 412 msec poststimulus period. Several aspects of the procedure were c o m m o n to BAEP and CAEP recordings. BAEPs and CAEPs were recorded from each of the 9 mice, but during different recording sessions. Ambient noise level in the acoustic chamber was 15-20 dBA and the ambient temperature was 22-24°C. After placement in the restrainer inside the acoustic chamber, each animal was allowed a 10-15 min adaptation period prior to recording. During this adaptation period, animals typically reduced movement and their rectal temperatures stabilized. Core temperature was monitored throughout each recording session by a rectal probe. An artifact rejection system, sensitive to significant deviations from background EEG, suspended averaging during movement. Results and Discussion
The purpose of the present report is to describe a methodology. For this reason, only a brief description of the parametric changes in the mouse BAEP and CAEP will be presented here. A more detailed description of changes in the mouse auditory evoked potentials as a function of stimulus parameters will be presented elsewhere (manuscript in preparation). Fig. 2A presents a series of BAEPs from one mouse collected over a broad range of click intensities. As this figure
T.M. W E L C H ET AL.
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4 1
2
~
~
70dB
1 2
3
5
~
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~
~
3
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~
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ill5~ V
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Fig. 2. Brain-stem auditory evoked potentials (BAEPs) from one mouse as a function of click intensity and repetition rate. All BAEP components occurred within 10 msec relative to the arrival of the stimulus at the mouse's ears. The respective group mean latencies of components P1 through P5 were 0.9, 1.6, 2.5, 3.1 and 4.2 msec while their mean peak-to-trough amplitudes were 2.8, 2.0, 4.3, 9.2 and 1.1 ~V in response to clicks with an intensity of 90 dB SPL p.e. and a repetition rate of 10/sec. A: as click intensity increased, the amplitudes of the various BAEP components increased while their latencies shortened. B: as click rate increased, the amplitudes of the various BAEP components decreased while their latencies were prolonged. Arrows near the beginning of each BAEP trace indicate the arrival of the click stimuli at the mouse's ear.
AEPs IN U N A N E S T H E T I Z E D MICE illustrates, the typical mouse BAEP was comprised of 5 vertexpositive wave forms (labeled P1 through P5) with the P4 component being the most prominent, especially at the lower click intensities. As the intensity of the clicks increased, the latency of the peaks decreased and their amplitudes increased. Fig. 2B illustrates BAEPs from one mouse taken during the click repetition rate condition. As click rate increased, peak latencies increased and amplitudes decreased. BAEP latencies were highly reliable and consistent both within and across animals. BAEP morphology for an individual animal was quite consistent over extended periods of time but differed somewhat between animals. Fig. 3 presents the CAEPs from one mouse evoked by paired tones at 3 different ISls. The CAEP had 3 early components and a large slower late positivity. There was an increase in latency and a decrease in amplitude of the various CAEP components between tone 1 and tone 2 responses. This effect was generally greater for progressively shorter ISis. There was consistency in the CAEP wave form within animals and across recording sessions. There were, however, noteworthy differences in CAEP morphology between animals. The morphology of the wave form and the changes seen in the BAEPs as a function of click rate and intensity were consistent with those reported in other species of laboratory animals (Grinnell 1963a, b; Huang and Buchwald 1978; Church et al. 1984b) as well as in humans (Rowe 1978; Stockard et al. 1978b). The morphology of the wave form of the mouse CAEP was similar to the CAEP recorded from the rat (Tokimoto et al. 1977; Bhargava et al. 1978). In addition, the increases in latencies and decrements in amplitudes of the CAEP from tone 1 to tone 2 were consistent with the changes reported in the human literature (D.W. Shucard et al. 1977; J.L. Shucard et al. 1981 ; Thomas and Shucard 1983). When the mice were initially placed in the acoustic chamber, their rectal temperatures averaged 36.9+0.5°C. Over a 10-15 min pre-recording time period, their rectal temperatures gradually increased by an average of 0.7 + 0.1 °C before stabilizing at 37.6_+0.4°C. It is well known that core temperature changes of 0.5°C or more can significantly affect the BAEP latencies of small laboratory animals (Jones et al. 1980; Williston and Jewett 1982). It is an important aspect of the procedure, therefore, to allow adequate time for core temperature stabilization before recording BAEPs in unanesthetized animals. Animals also typically reduced movement and gnawing behavior during this pre-recording adaptation period and remained relatively movement free thereafter. Contamination of evoked potentials from movement artifact is a common concern when recording from unanesthetized animals. Movement artifact from our unanesthetized mice posed little problem because (a) the animals were typically passive as a result of the adaptation regiment, (b) the design of our animal restrainer rendered the animal's head virtually steadfast even when an animal attempted movement, and (c) when occasional movement artifact was detected in the recording leads an artifact rejection system suspended averaging of the evoked potential. In conclusion, our recording technique has provided con-
81 sistent and reproducible evoked potentials over time in the unanesthetized mouse. Following the procedures outlined above, we were able to obtain excellent recordings during a typical 1 h
N~ N90
~f~
~
2.0sec
ISI-
~,~-~
A ISl - 1.Osec
TONE 1
~~ V ~
ISI - 0.6sec
TONE
2
J 25 ~v 40 msec
Fig. 3. Cortical auditory evoked potentials (CAEPs) to paired tones from one mouse as a function of inter-stimulus intervals of 2.0, 1.0 and 0.6 sec. Arrows indicate stimulus onset. The early components, labeled according to polarity and approximate latency, were designated as P10-N35, N35-P70 and P70N90. The respective mean amplitudes of these CAEP components were 62.0, 53.0 and 29.0 ttV in response to tone 1 with an inter-stimulus interval of 2 sec and inter-pair interval of 4 sec.
82
T.M. W E L C H ET AL.
session. In addition, to date we have recorded from animals up to 4 months after surgery. Thus, this method could be used in a wide variety of investigations involving acute and chronic measures of electrophysiological activity in the mouse.
We would like to thank Tasleem Qaasim for the typing of this manuscript and Barry Silverstein for his diagram of the animal restraining device and his assistance with the photography.
Summary
References
Anesthetizing or sedating animals affords control over movement artifact during electrophysiological recording. However, the use of chemical restraint leads to unwanted side effects such as drug-induced hypothermia. Hypothermia is problematic because BAEP amplitudes and latencies are affected significantly by core temperature changes. Moreover, several recent studies have indicated that anesthetics and sedatives m a y significantly alter the BAEP and CAEP even in thermoregulated subjects. There is a need, therefore, to develop a practical technique for obtaining chronic BAEPs and CAEPs in restrained, unanesthetized animals. The present report describes a technique that permits the gathering of consistent, reliable evoked potential recordings over time in unanesthetized mice. The preparation is useful for studying the CNS effects of pharmacological agents, pathological conditions, aging and development in the mouse and could be adapted for use with other small animals as well.
Bhargava, V.K., Salamy, A. and McKean, C.M. Effects of cholinergic drugs on auditory evoked responses (CER) of the rat cortex. Neuropharmacology, 1978, 17: 1009-1013. Brazier, M.A.B. The Neurophysiological Background for Anesthesia. Thomas, Springfield, IL, 1972. Church, M.W. and Williams, H.L. Dose- and time-dependent effects of ethanol on brain stem auditory evoked responses in young adult males. Electroenceph. clin. Neurophysiol., 1982, 54: 161-174. Church, M.W., Welch, T.M. and Shucard, D.W. Pentobarbitalinduced changes in the mouse auditory brainstem responses as a function of stimulus repetition rate and time post-drug. Abstr. Soc. Neurosci., 1984a, 10: 112. Church, M.W., Williams, H.L. and Holloway, J.A. Brain-stem auditory evoked potentials in the rat: effects of gender, stimulus characteristics, and ethanol sedation. Electroenceph. clin. Neurophysiol., 1984b, 59: 328-339. Grinnell, A.D. The neurophysiology of audition in bats: intensity and frequency parameters. J. Physiol. (Lond.), 1963a, 167: 38-66. Grinnell, A.D. The neurophysiology of audition in bats: temporal parameters. J. Physiol. (Lond.), 1963b, 167: 67-96. Henry, K.R. Differential changes of auditory nerve and brain stem short latency evoked potentials in the laboratory mouse. Electroenceph. clin. Neurophysiol., 1979, 46: 452-459. Huang, C.M. and Buchwald, J.S. Factors that affect the amplitudes and latencies of the vertex short latency acoustic responses in the cat. Electroenceph. clin. Neurophysiol., 1978, 44: 179-186. Jones, T.A., Stockard, T.J. and Weidner, W.J. The effects of temperature and acute alcohol intoxication on brain stem auditory evoked protentials in the cat. Electroenceph. clin. Neurophysiol., 1980, 4 9 : 2 3 30. Marshall, N.K. and Donchin, E. Circadian variations in the latency of brainstem responses and its relation to body temperature. Science, 1981, 212: 356-358. Rowe, M.J. Normal variability of the brain stem auditory evoked response in young and old adult subjects. Electroenceph. clin. Neurophysiol., 1978, 44: 459-470. Shucard, D.W., Shucard, J.L. and Thomas, D.G. Auditory evoked potentials as probes of hemispheric differences in cognitive processing. Science, 1977; 197: 1295-1298. Shucard, J.L., Shucard, D.W., Cummins, K.R. and Campos, J.J. Auditory evoked potentials and sex-related differences in brain development. Brain and Language, 1981, 13: 91-102.
Rrsum6 Mbthode d'enregistrernent chronique des potentiels ~voquks auditifs du tronc et du cortex chez la souris non anesth~sibe
Mettre un animal sous anesth6siant ou sous srdatif permet de contr61er les artrfacts de mouvement au cours d'enregistrements 61ectrophysiologiques. Toutefois, l'utilisation de ces substances comme moyen d'immobilisation entra~me rapparition d'effets secondaires non voulus telle l'induction pharmacologique d'une hypothermie. L'hypothermie pose un probl~me car l'amplitude et la latence des potentiels 6voqurs auditifs du tronc crrrbral sont significativement affectres par les modifications de la temprrature centrale. De plus, plusieurs 6tudes rrcentes ont indiqu6 que les anesthrsiques et les srdatifs pouvaient altrrer significativement les potentiels 6voqu6s auditifs du tronc et du cortex, m~me lorsque la temprrature des sujets 6tait rrgulre. Il 6tait donc nrcessaire de drvelopper une m r t h o d e pratique pour obtenir l'enregistrement des potentiels 6voqurs auditifs du tronc et du cortex chez des animaux en contention et sans anesthrsie. Le prrsent travail drcrit une technique qui permet d'obtenir des enregistrements de potentiels 6voqurs stables et fiables au cours du temps chez la souris non anesthrsi6e. La prrparation est utile pour l'rtude des effets sur le systrme nerveux central, d'agents pharmacologiques, de conditions pathologiques, du vieillissement ou du drveloppement chez la souris; elle peut ~tre adaptre h l'rtude d'autres petits animaux.
AEPs IN UNANESTHETIZED MICE Squires, K.C., Chu, N.S. and Starr, A. Auditory brain stem potentials with alcohol. Electroenceph. clin. Neurophysiol., 1978, 45: 577-584. Stockard, J.J., Sharbrough, F.W. and Tinker, J.A. Effects of hypothermia on human brainstem auditory response. Ann. Neurol., 1978a, 3: 368-370. Stockard, J.J., Stockard, J.E. and Sharbrough, F.W. Non-pathological factors influencing brainstem auditory evoked potentials. Amer. J. EEG Technol., 1978b, 18: 177-209. Sutton, L.N., Frewen, T., Marsh, R., Jaggi, J. and Bruce, D.A. The effects of deep barbiturate coma on multimodality evoked potentials. J. Neurosurg., 1982, 57: 178-185.
83 Thomas, D.G. and Shucard, D.W. The use of a control or baseline condition in electrophysiological studies of hemispheric specialization of function. Electroenceph. clin. Neurophysiol., 1983, 55: 575-579. Tokimoto, T., Osako, S. and Matsuura, S. Development of auditory evoked cortical and brain stem responses during the early postnatal period in the rat. Osaka City med. J., 1977, 23: 141-153. Williston, J.S. and Jewett, D.L. The Q10 of auditory brainstem responses in rats under hypothermia. Audiology, 1982, 21: 457-465.
Electroencephalograph), and clinical Neurophysiology, 185, 6 0 : 7 8 - 8 3 Elsevier Scientific Publishers Ireland, Ltd.
Short communication A METHOD FOR CHRONICALLY RECORDING BRAIN-STEM EVOKED POTENTIALS FROM UNANESTHETIZED MICE i
AND CORTICAL
AUDITORY
T.M. W E L C H , M.W. C H U R C H and D.W. S H U C A R D 2
Brain Sciences Laboratories, National Jewish Hospital and Research Center, and Department of Psychiatry, University of Colorado School of Medicine, Denver, CO (U.S.A.) (Accepted for publication: August 24, 1984)
The brain-stem auditory evoked potential (BAEP), a shortlatency far-field sensory evoked potential, has become an important clinical and experimental tool for assessing the integrity of the subcortical auditory pathway in h u m a n s and animals. The cortical auditory evoked potential (CAEP), a long-latency sensory evoked potential, is widely used to investigate sensory and cognitive processing of auditory information at the cortical level. Animal investigations offer the opportunity to study the BAEP and CAEP under well controlled conditions. Of particular significance when recording evoked potentials is the state of the animal being studied. Animal BAEP recordings are most often done under sedation or anesthesia. Although anesthesia affords control over artifacts due to movement or muscle activity, the use of anesthesia also leads to unwanted side effects such as hypothermia. In our experience, anesthetic-induced hypothermia readily occurs in small animals such as the laboratory mouse. Further, relatively small temperature decreases significantly affect the BAEP (Stockard et al. 1978a; Marshall and Donchin 1981; Williston and Jewett 1982). It also has been shown that anesthesia and sedation may affect the latencies a n d / o r amplitudes of both the BAEP (Squires et al. 1978; Stockard et al. 1978b; Henry 1979; Church and Williams 1982; Sutton et al. 1982; Church et al. 1984a, b) and the C A E P (Brazier 1972; Sutton et al. 1982) even in thermoregulated animals. Moreover, use of an anesthetic may interact with or mask the clinical and experimental conditions under investigation. Based on the above, it may be more appropriate to record BAEPs and CAEPs without the use of anesthesia. In the present report, we describe a technique for obtaining BAEPs and CAEPs in unanesthetized mice. The technique is useful for investigating the CNS effects of pharmacological agents, pathological conditions, aging and development without the possible confounding effects of anesthesia.
1 Supported in part by N I M H Grant No. M H 15442 and N1H Grant No. N S / A M 15855. 2 Address reprint requests to: D.W. Shucard, Ph.D., Brain Sciences Laboratories, National Jewish Hospital, 3800 East Colfax Ave., Denver, CO 80206, U.S.A.
Method Subjects The subjects were 9 BDF 1 female mice weighing 25 29 g before surgery with ages ranging from 4 to 6 months. These animals were bred in our animal facilities, housed individually in plexiglass cages (26 c m x 1 6 c m x l 3 cm), kept on a 12 h light/dark cycle, and fed lab chow and water ad lib.
Surgery All subjects were anesthetized with 80 m g / k g (i.p.) of sodium pentobarbital and secured on a small operating table. After a midline scalp incision was made, the skin and connective tissue were scraped away from the skull surface. Muscle and other soft tissue were cauterized to retard tissue regrowth. Once the surrounding tissue was cleared away, the skull surface was cleaned with 70% ethanol. Burr holes were then drilled into the skull to accommodate 3 stainless steel skull screws (0-80× 1 / 8 " stainless steel, flathead) that were placed in the following configuration: two screws, one on each side, were placed 2 m m lateral to the midline (sagittal suture) and one-half the distance between bregma and lambda. The third screw was placed 1.5 m m posterior to and 1.5 m m lateral to the left of lambda. When inserting the skull screws, care was taken not to cause brain compression or penetration of the dura. Once the skull screws were in place, male gold pins, previously connected to each of the screws via a teflon-coated silver wire (0.02 m m diameter), were inserted into a plastic strip connector cut to a length of 0.8 cm and a height of 0.5 cm. Each strip connector had 3 holes which accommodated the 3 male pin connectors, one connector attached to each of the wire leads from the 3 skull screws. Dental acrylic was then applied to the skull screws, skull surface and strip connector. Once a small base of dental acrylic was built, a female headpost was placed posterior to the strip connector and cemented into place with acrylic. The skull screws, strip connector and headpost were then all secured with dental acrylic. The headpost was machined from a no. 10-34 brass rod, had a length of 0.8 cm, and was drilled lengthwise and tapped to accommodate a no. 440 screw. The skull and surrounding tissue were then infused with an antibiotic ointment (bacitracin) and a single wound clip was used to partially
0013-4649/85/$03.30 © 1985 Elsevier Scientific Publishers Ireland, Ltd.
AEPs IN U N A N E S T H E T I Z E D MICE
A
79 close the incision. Animals were thermoregulated throughout the surgery and allowed 3-8 weeks to recover.
Adaptation
i
i i
After surgery but prior to evoked potential recording, the mice were gentled to reduce stress. This procedure involved handling the animals for 15 min every other day for a 2 week period. Adaptation to the restrainer and recording conditions was accomplished by placing each mouse in a mock recording setup for 1 h prior to the day of the actual experiment. The animal restrainer was also placed in each mouse cage for 1 night to further facilitate the animal's adaptation to the restrainer.
Apparatus The recording environment consisted of an animal restrainer mounted in a box placed inside an acoustic chamber. The animal restrainer was designed to restrict body movement and immobilize the animal's head. Fig. 1A illustrates the design of the restrainer. The animal restrainer was fastened to a floor bar in the box with a small C-clamp to prevent it from moving. Recording leads, suspended from the top of the box, connected the skull electrodes via the strip connector with the evoked potential recording system. The sides of the box were padded with 2.54 cm thick foam rubber for acoustical dampening. The box, containing the animal in its restrainer, was placed in an electrically shielded 104 cm x 96.5 cm × 81 cm acoustic chamber. Fig. 1B illustrates the position of the mouse after placement in the device during recording.
Stimuli
Fig. 1. A: design of the animal restrainer. The mouse restrainer consists of a body tube which encases and supports the animal's body. Inside the body tube is a backstop that can be adjusted to the animal's body length. A 0.5 cm slot, cut lengthwise along the bottom of the body tube, allows the experimenter to pull the animal by the tail into the restrainer. An adjustable neck collar, incorporated into the front of the restrainer, prevents the animal from withdrawing its head into the restrainer. A male screw, mounted on a sliding bar at the front and top of the restrainer, is screwed into the female headpost cemented on the animal's head. B: position of the mouse during recording. With the mouse's head extended past the opening of the restrainer, the collar (a) is adjusted around the mouse's neck and tightened into place. The male screw attached to the restrainer (b) is threaded into a female headpost (d) which is permanently affixed to the strip connector. The recording leads (e) are
Rarefaction 'click' stimuli were used to produce BAEP recordings, Click stimuli were created by activating an earphone with a 100 #sec square wave pulse generated by an Iconix system. The earphone was mounted directly in front of each animal at ear level and at a distance of 13.8 cm from the animal's ears. Click intensity was measured in dB SPL peak equivalent (p.e.) at the mouse's ears. Spectral energy of the click was broadly distributed below 4 kHz with maximal energy between 3.0 and 3.5 kHz. Paired tones were used to produce CAEP recordings. Each tone was 100 msec in duration with an intensity of 80 dB SPL at the mouse's ears. A 6 kHz tone was selected to evoke CAEPs because initial investigations in our laboratory indicated that this frequency produced robust and well-delineated mouse CAEPs.
Recordings The BAEP was recorded ipsilaterally between the skull screw electrode placed to the left and posterior to lambda (positive) and the screw electrode placed to the left of the sagittal suture (negative). The remaining screw electrode served attached to the strip connector (f). The animal's head is held steadfast by the male screw that descends from the top of the restrainer and is threaded into the female headpost cemented to the mouse's head (d).
80 as a ground. BAEPs were recorded from each animal as a function of parametric variations in click repetition rate and stimulus intensity. With respect to repetition rate, BAEPs were collected in response to clicks presented at rates of 10, 50 and 100/see (intensity = 90 dB SPL p.e.). With respect to intensity, BAEPs were collected in response to clicks presented at 30, 35, 40, 50, 70, 90 and 110 dB SPL p.e. (rate = 10 clicks/sec). Each animal's BAEPs were amplified by an amplifier with sensitivity set at 1 ~ V / m m and a bandpass of 300-3000 Hz. The amplified signals were averaged by a signal averager, with positivity displayed upwards, and printed out on an X-Y plotter. BAEP activity was sampled at a rate of 40 vsec per address and 256 responses were averaged. The analysis epoch of 10.24 msec (256 addresses) consisted of a 0.50 msec prestimulus and a 9.74 msec poststimuhis period. The 0.50 msec prestimulus period consisted of a 0.40 msec acoustic transit time and a 0.10 msec equipment delay. This prestimulus period allowed for adequate temporal separation of stimulus artifact from the first BAEP component. The CAEP was recorded contralaterally between the screw electrode placed to the left and posterior to lambda (negative) and the screw electrode placed to the right of the sagittal suture (positive). The remaining screw served as a ground. CAEPs to the 80 dB SPL paired tones were evaluated at inter-stimulus intervals of 2.0, 1.0 and 0.6 sec with a time interval of 4.0 sec between successive tone pairs. Each animal's CAEPs were amplified with sensitivity set at 3 0 / t V / m m and a bandpass of 3 - 3 0 Hz. The amplified signals were averaged with positivity displayed upwards and printed out via the X-Y plotter. CAEP activity was sampled at a rate of 1 msec per address and 32 responses were averaged. The analysis epoch of 512 msec (512 addresses) consisted of a 100 msec prestimulus and a 412 msec poststimulus period. Several aspects of the procedure were c o m m o n to BAEP and CAEP recordings. BAEPs and CAEPs were recorded from each of the 9 mice, but during different recording sessions. Ambient noise level in the acoustic chamber was 15-20 dBA and the ambient temperature was 22-24°C. After placement in the restrainer inside the acoustic chamber, each animal was allowed a 10-15 min adaptation period prior to recording. During this adaptation period, animals typically reduced movement and their rectal temperatures stabilized. Core temperature was monitored throughout each recording session by a rectal probe. An artifact rejection system, sensitive to significant deviations from background EEG, suspended averaging during movement. Results and Discussion
The purpose of the present report is to describe a methodology. For this reason, only a brief description of the parametric changes in the mouse BAEP and CAEP will be presented here. A more detailed description of changes in the mouse auditory evoked potentials as a function of stimulus parameters will be presented elsewhere (manuscript in preparation). Fig. 2A presents a series of BAEPs from one mouse collected over a broad range of click intensities. As this figure
T.M. W E L C H ET AL.
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~
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Fig. 2. Brain-stem auditory evoked potentials (BAEPs) from one mouse as a function of click intensity and repetition rate. All BAEP components occurred within 10 msec relative to the arrival of the stimulus at the mouse's ears. The respective group mean latencies of components P1 through P5 were 0.9, 1.6, 2.5, 3.1 and 4.2 msec while their mean peak-to-trough amplitudes were 2.8, 2.0, 4.3, 9.2 and 1.1 ~V in response to clicks with an intensity of 90 dB SPL p.e. and a repetition rate of 10/sec. A: as click intensity increased, the amplitudes of the various BAEP components increased while their latencies shortened. B: as click rate increased, the amplitudes of the various BAEP components decreased while their latencies were prolonged. Arrows near the beginning of each BAEP trace indicate the arrival of the click stimuli at the mouse's ear.
AEPs IN U N A N E S T H E T I Z E D MICE illustrates, the typical mouse BAEP was comprised of 5 vertexpositive wave forms (labeled P1 through P5) with the P4 component being the most prominent, especially at the lower click intensities. As the intensity of the clicks increased, the latency of the peaks decreased and their amplitudes increased. Fig. 2B illustrates BAEPs from one mouse taken during the click repetition rate condition. As click rate increased, peak latencies increased and amplitudes decreased. BAEP latencies were highly reliable and consistent both within and across animals. BAEP morphology for an individual animal was quite consistent over extended periods of time but differed somewhat between animals. Fig. 3 presents the CAEPs from one mouse evoked by paired tones at 3 different ISls. The CAEP had 3 early components and a large slower late positivity. There was an increase in latency and a decrease in amplitude of the various CAEP components between tone 1 and tone 2 responses. This effect was generally greater for progressively shorter ISis. There was consistency in the CAEP wave form within animals and across recording sessions. There were, however, noteworthy differences in CAEP morphology between animals. The morphology of the wave form and the changes seen in the BAEPs as a function of click rate and intensity were consistent with those reported in other species of laboratory animals (Grinnell 1963a, b; Huang and Buchwald 1978; Church et al. 1984b) as well as in humans (Rowe 1978; Stockard et al. 1978b). The morphology of the wave form of the mouse CAEP was similar to the CAEP recorded from the rat (Tokimoto et al. 1977; Bhargava et al. 1978). In addition, the increases in latencies and decrements in amplitudes of the CAEP from tone 1 to tone 2 were consistent with the changes reported in the human literature (D.W. Shucard et al. 1977; J.L. Shucard et al. 1981 ; Thomas and Shucard 1983). When the mice were initially placed in the acoustic chamber, their rectal temperatures averaged 36.9+0.5°C. Over a 10-15 min pre-recording time period, their rectal temperatures gradually increased by an average of 0.7 + 0.1 °C before stabilizing at 37.6_+0.4°C. It is well known that core temperature changes of 0.5°C or more can significantly affect the BAEP latencies of small laboratory animals (Jones et al. 1980; Williston and Jewett 1982). It is an important aspect of the procedure, therefore, to allow adequate time for core temperature stabilization before recording BAEPs in unanesthetized animals. Animals also typically reduced movement and gnawing behavior during this pre-recording adaptation period and remained relatively movement free thereafter. Contamination of evoked potentials from movement artifact is a common concern when recording from unanesthetized animals. Movement artifact from our unanesthetized mice posed little problem because (a) the animals were typically passive as a result of the adaptation regiment, (b) the design of our animal restrainer rendered the animal's head virtually steadfast even when an animal attempted movement, and (c) when occasional movement artifact was detected in the recording leads an artifact rejection system suspended averaging of the evoked potential. In conclusion, our recording technique has provided con-
81 sistent and reproducible evoked potentials over time in the unanesthetized mouse. Following the procedures outlined above, we were able to obtain excellent recordings during a typical 1 h
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Fig. 3. Cortical auditory evoked potentials (CAEPs) to paired tones from one mouse as a function of inter-stimulus intervals of 2.0, 1.0 and 0.6 sec. Arrows indicate stimulus onset. The early components, labeled according to polarity and approximate latency, were designated as P10-N35, N35-P70 and P70N90. The respective mean amplitudes of these CAEP components were 62.0, 53.0 and 29.0 ttV in response to tone 1 with an inter-stimulus interval of 2 sec and inter-pair interval of 4 sec.
82
T.M. W E L C H ET AL.
session. In addition, to date we have recorded from animals up to 4 months after surgery. Thus, this method could be used in a wide variety of investigations involving acute and chronic measures of electrophysiological activity in the mouse.
We would like to thank Tasleem Qaasim for the typing of this manuscript and Barry Silverstein for his diagram of the animal restraining device and his assistance with the photography.
Summary
References
Anesthetizing or sedating animals affords control over movement artifact during electrophysiological recording. However, the use of chemical restraint leads to unwanted side effects such as drug-induced hypothermia. Hypothermia is problematic because BAEP amplitudes and latencies are affected significantly by core temperature changes. Moreover, several recent studies have indicated that anesthetics and sedatives m a y significantly alter the BAEP and CAEP even in thermoregulated subjects. There is a need, therefore, to develop a practical technique for obtaining chronic BAEPs and CAEPs in restrained, unanesthetized animals. The present report describes a technique that permits the gathering of consistent, reliable evoked potential recordings over time in unanesthetized mice. The preparation is useful for studying the CNS effects of pharmacological agents, pathological conditions, aging and development in the mouse and could be adapted for use with other small animals as well.
Bhargava, V.K., Salamy, A. and McKean, C.M. Effects of cholinergic drugs on auditory evoked responses (CER) of the rat cortex. Neuropharmacology, 1978, 17: 1009-1013. Brazier, M.A.B. The Neurophysiological Background for Anesthesia. Thomas, Springfield, IL, 1972. Church, M.W. and Williams, H.L. Dose- and time-dependent effects of ethanol on brain stem auditory evoked responses in young adult males. Electroenceph. clin. Neurophysiol., 1982, 54: 161-174. Church, M.W., Welch, T.M. and Shucard, D.W. Pentobarbitalinduced changes in the mouse auditory brainstem responses as a function of stimulus repetition rate and time post-drug. Abstr. Soc. Neurosci., 1984a, 10: 112. Church, M.W., Williams, H.L. and Holloway, J.A. Brain-stem auditory evoked potentials in the rat: effects of gender, stimulus characteristics, and ethanol sedation. Electroenceph. clin. Neurophysiol., 1984b, 59: 328-339. Grinnell, A.D. The neurophysiology of audition in bats: intensity and frequency parameters. J. Physiol. (Lond.), 1963a, 167: 38-66. Grinnell, A.D. The neurophysiology of audition in bats: temporal parameters. J. Physiol. (Lond.), 1963b, 167: 67-96. Henry, K.R. Differential changes of auditory nerve and brain stem short latency evoked potentials in the laboratory mouse. Electroenceph. clin. Neurophysiol., 1979, 46: 452-459. Huang, C.M. and Buchwald, J.S. Factors that affect the amplitudes and latencies of the vertex short latency acoustic responses in the cat. Electroenceph. clin. Neurophysiol., 1978, 44: 179-186. Jones, T.A., Stockard, T.J. and Weidner, W.J. The effects of temperature and acute alcohol intoxication on brain stem auditory evoked protentials in the cat. Electroenceph. clin. Neurophysiol., 1980, 4 9 : 2 3 30. Marshall, N.K. and Donchin, E. Circadian variations in the latency of brainstem responses and its relation to body temperature. Science, 1981, 212: 356-358. Rowe, M.J. Normal variability of the brain stem auditory evoked response in young and old adult subjects. Electroenceph. clin. Neurophysiol., 1978, 44: 459-470. Shucard, D.W., Shucard, J.L. and Thomas, D.G. Auditory evoked potentials as probes of hemispheric differences in cognitive processing. Science, 1977; 197: 1295-1298. Shucard, J.L., Shucard, D.W., Cummins, K.R. and Campos, J.J. Auditory evoked potentials and sex-related differences in brain development. Brain and Language, 1981, 13: 91-102.
Rrsum6 Mbthode d'enregistrernent chronique des potentiels ~voquks auditifs du tronc et du cortex chez la souris non anesth~sibe
Mettre un animal sous anesth6siant ou sous srdatif permet de contr61er les artrfacts de mouvement au cours d'enregistrements 61ectrophysiologiques. Toutefois, l'utilisation de ces substances comme moyen d'immobilisation entra~me rapparition d'effets secondaires non voulus telle l'induction pharmacologique d'une hypothermie. L'hypothermie pose un probl~me car l'amplitude et la latence des potentiels 6voqurs auditifs du tronc crrrbral sont significativement affectres par les modifications de la temprrature centrale. De plus, plusieurs 6tudes rrcentes ont indiqu6 que les anesthrsiques et les srdatifs pouvaient altrrer significativement les potentiels 6voqu6s auditifs du tronc et du cortex, m~me lorsque la temprrature des sujets 6tait rrgulre. Il 6tait donc nrcessaire de drvelopper une m r t h o d e pratique pour obtenir l'enregistrement des potentiels 6voqurs auditifs du tronc et du cortex chez des animaux en contention et sans anesthrsie. Le prrsent travail drcrit une technique qui permet d'obtenir des enregistrements de potentiels 6voqurs stables et fiables au cours du temps chez la souris non anesthrsi6e. La prrparation est utile pour l'rtude des effets sur le systrme nerveux central, d'agents pharmacologiques, de conditions pathologiques, du vieillissement ou du drveloppement chez la souris; elle peut ~tre adaptre h l'rtude d'autres petits animaux.
AEPs IN UNANESTHETIZED MICE Squires, K.C., Chu, N.S. and Starr, A. Auditory brain stem potentials with alcohol. Electroenceph. clin. Neurophysiol., 1978, 45: 577-584. Stockard, J.J., Sharbrough, F.W. and Tinker, J.A. Effects of hypothermia on human brainstem auditory response. Ann. Neurol., 1978a, 3: 368-370. Stockard, J.J., Stockard, J.E. and Sharbrough, F.W. Non-pathological factors influencing brainstem auditory evoked potentials. Amer. J. EEG Technol., 1978b, 18: 177-209. Sutton, L.N., Frewen, T., Marsh, R., Jaggi, J. and Bruce, D.A. The effects of deep barbiturate coma on multimodality evoked potentials. J. Neurosurg., 1982, 57: 178-185.
83 Thomas, D.G. and Shucard, D.W. The use of a control or baseline condition in electrophysiological studies of hemispheric specialization of function. Electroenceph. clin. Neurophysiol., 1983, 55: 575-579. Tokimoto, T., Osako, S. and Matsuura, S. Development of auditory evoked cortical and brain stem responses during the early postnatal period in the rat. Osaka City med. J., 1977, 23: 141-153. Williston, J.S. and Jewett, D.L. The Q10 of auditory brainstem responses in rats under hypothermia. Audiology, 1982, 21: 457-465.