Injection anaesthesia with fentanyl–midazolam–medetomidine in adult female mice: importance of antagonization and perioperative care

Abstract

Injection anaesthesia is commonly used in laboratory mice; however, a disadvantage is that post-anaesthesia recovery phases are long. Here, we investigated the potential for shortening the recovery phase after injection anaesthesia with fentanyl–midazolam–medetomidine by antagonization with naloxone–flumazenil–atipamezole. In order to monitor side-effects, the depth of anaesthesia, heart rate (HR), core body temperature (BT) and concentration of blood gases, as well as reflex responses, were assessed during a 50 min anaesthesia. Mice were allowed to recover from the anaesthesia in their home cages either with or without antagonization, while HR, core BT and spontaneous home cage behaviours were recorded for 24 h. Mice lost righting reflex at 330 ± 47 s after intraperitoneal injection of fentanyl–midazolam–medetomidine. During anaesthesia, HR averaged 225 ± 23 beats/min, respiratory rate and core BT reached steady state at 131 ± 15 breaths/min and 34.3 ± 0.25℃, respectively. Positive pedal withdrawal reflex, movement triggered by tail pinch and by toe pinch, still occurred in 25%, 31.2% and 100% of animals, respectively. Arterial blood gas analysis revealed acidosis, hypoxia, hypercapnia and a marked increase in glucose concentration. After anaesthesia reversal by injection with naloxone–flumazenil–atipamezole, animals regained consciousness after 110 ± 18 s and swiftly returned to physiological baseline values, yet they displayed diminished levels of locomotion and disrupted circadian rhythm. Without antagonization, mice showed marked hypothermia (22 ± 1.9℃) and bradycardia (119 ± 69 beats/min) for several hours. Fentanyl–midazolam–medetomidine provided reliable anaesthesia in mice with reasonable intra-anaesthetic side-effects. Post-anaesthetic period and related adverse effects were both reduced substantially by antagonization with naloxone–flumazenil–atipamezole.

Keywords

anaesthesia mouse antagonization fentanyl midazolam medetomidine

Anaesthesia in laboratory mice is often induced by intraperitoneal injection of a mixture of two or three drugs, which usually belong to different substance classes and thus exhibit different mechanisms of action with possible synergistic or additive effects, see examples.^1–5 Details of several substance combinations have been published over the years, with criteria for selecting and combining anaesthetic agents depending on the purpose for which anaesthesia is needed (e.g. surgery, imaging or terminal perfusion). In addition, the potential for drugs used in anaesthesia to interfere with experimental procedures and results can influence the choice of appropriate anaesthetic regimen.^4,5 Also, problems of availability, particularly in many countries where drugs are regulated under narcotic laws, can determine whether specific substances can be applied or not.^1–3

A well-known drawback of injection anaesthesia protocols for mice is the relatively long recovery phase, in which the animal is not in a deep stage of anaesthesia, but remains more or less immobilized while responding to stimuli. In this phase, hypothermia is the most prominent problem, but metabolic, respiratory and cardiovascular functions are usually also impaired.

To circumvent these problems, a mixture containing drugs that could be antagonized was suggested.⁶ A fully reversible three-component anaesthesia⁷ that has been shown to achieve safe and reliable anaesthesia in chinchillas,⁶ ferrets,⁸ and rats⁹ was proposed. This anaesthesia protocol has since been used in mice: Thal and Plesnila⁴ administered the anaesthetic mixture in the suggested species-specific dosages (fentanyl 0.05 mg/kg, midazolam 5 mg/kg, medetomidine 0.5 mg/kg) in female C57BL/6 mice. As Thal and Plesnila aimed to obtain long-term measurements (approximately 3 h) of blood pressure and partial pressure of carbon dioxide (PaCO₂) under standardized conditions, animals were ventilated mechanically and injection of anaesthetics had to be repeated. Remarkably, at 1.5–2 h after reversal of anaesthesia, a marked drop in body temperature (BT) to 32℃ was found by punctual measurements using a rectal probe.⁴

The protocol used by Thal and Plesnila⁴ and Henke and colleagues⁷ consisted of full reversal, i.e. all three components for inducing anaesthesia were antagonized at a predetermined time point. Consequently the analgesic effect of fentanyl was equally completely eliminated with naloxone, which could be a disadvantage if post-operative pain remains persistent after anaesthesia. With regards to post-procedural pain, it could be beneficial to omit naloxone and instead inject buprenorphine, which might antagonize some effects of fentanyl while providing pain alleviation for some hours. However, to date, the protocol of Henke and colleagues has often been applied in experiments which do not induce long-term substantial pain, but require anaesthesia for immobilizing the animal for a certain time, e.g. for marking, probe sampling, intratracheal instillation, measurements of physiological values,⁹ and the many types of imaging procedures (e.g. functional magnetic resonance imaging [fMRI] and positron emission tomography [PET]). For such painless investigations, mice are often subjected to repeated anaesthesia for which a fully reversible anaesthesia protocol might be useful, as (side-) effects of fentanyl are antagonized and prompt recovery from anaesthesia could be provided at a predefined time point.

We aimed to use the fully reversible mixture as a protocol for routine laboratory settings (e.g. 30–50 min anaesthesia), where post-anaesthetic or post-operative care has to be considered. Therefore we analysed the potential for the protocol not only to provide reliable anaesthesia but also to ease termination of anaesthesia in order to promote smooth and fast recovery. Thus, antagonization ought to mitigate persistent hypothermia, reduced general condition, inactivity and lack of food and water intake as well as other spontaneous behaviours and signs of hampered recovery, impaired health and reduced well-being.

This study aimed to characterize intraperitoneal injection anaesthesia with fentanyl–midazolam–medetomidine in terms of safety, reliability and post-anaesthetic health and well-being. Since fast and smooth recovery after anaesthesia is desired in many experimental settings, we focused on the impact of the reversal of anaesthesia by investigating the beneficial, as well as any possible adverse, effects of antagonization by recording continuous telemetry and video data for a full 24 h period after anaesthesia.

Materials and methods

Ethics statement

The animal housing and all procedures and protocols were approved by the Cantonal Veterinary Office (Zurich, Switzerland) under the license number 86/2011. Housing and experimental procedures were in accordance with Swiss animal protection law and also conform to Directive 2010/63 EU of the European Parliament and of the Council of 22 September 2010 on the Protection of vertebrate animals used for experimental and other scientific purposes and to the Guide for the care and use of laboratory animals.¹⁰

Animals and standard housing conditions

Thirty-two four-week-old female C57BL/6 J mice were obtained from a commercial supplier (Harlan, Horst, The Netherlands). Mice were housed in groups of two to four animals prior to and after the experiments. All mice were housed individually during the experiments as single housing is a prerequisite for data acquisition and analyses (e.g. obtaining telemetric recordings, filming of behaviour and automated analysis). The mice were free of all viral, bacterial, and parasitic pathogens listed in the Federation of European Laboratory Animal Science Associations (FELASA) recommendations, except for the Helicobacter species. The animals’ health status was monitored by a sentinel program according to the FELASA guidelines throughout the experiments.¹¹

The animals were kept in Eurostandard type III open-top plastic cages (425 mm × 266 mm × 155 mm, floor area 820 cm²; Tecniplast, Indulab, Gams, Switzerland) with autoclaved dust-free sawdust bedding (80–90 g per cage, LTE E-001 Abedd; Indulab). A standard cardboard house (Ketchum Manufacturing, Brockville, ON, Canada) served as a shelter, except during the experiments. Animals were fed a pelleted and extruded mouse diet (Kliba No. 3431; Provimi Kliba, Kaiseraugst, Switzerland) ad libitum and had unrestricted access to sterilized drinking water. Autoclaved hay (8–12 g/cage) and one Nestlet™ per cage (about 5 × 5 cm, consisting of pressed cotton fibres; Indulab) were provided as nesting material.

The light/dark cycle in the animal room consisted of a 12 h/12 h cycle (lights on at 15:00 h, lights off at 03:00 h, according to the experimental set-up and recording sequences) with artificial light of approximately 40 lux in the cage. The mean room temperature was 21 ± 1℃, with a relative humidity of 50 ± 5% and 15 complete changes of filtered air per hour (HEPA H14 filter); the air pressure was controlled at 50 Pa. The animal room was insulated to prevent electronic or other noise.

Experimental design

Treatment groups

Mice were assigned randomly to four groups of eight animals. All animals underwent anaesthesia lasting 50 min. Sixteen mice were implanted with telemetric transmitters prior to the experiments in order to obtain intra- and post-anaesthetic measurements of heart rate (HR) and core BT, either with or without reversal of anaesthesia. Another 16 animals were used for behavioural analysis (video recordings) during the post-anaesthetic recovery period, either with or without reversal of anaesthesia. At four weeks after the initial anaesthesia experiment, eight of these animals underwent a terminal experiment to obtain arterial blood with which to determine intra-anaesthetic acid–base balance and blood gas concentration during a second anaesthesia of 50 min duration.

Transmitter implantation

Prior to the experiments, at about age 8–9 weeks, 16 mice were instrumented with telemetric transmitters. TA10ETA-F20 transmitters (Data Sciences International, St Paul, MN, USA) – which measure HR and core BT in freely moving mice – were implanted as described previously in detail.¹² Briefly, under anaesthesia with sevoflurane (Sevorane®; Abbott, Baar, Switzerland), the transmitter body was implanted in the abdomen under aseptic conditions. One wired loop electrode was fixed with silk sutures (Perma-Handseide 6–0; Ethicon, Norderstedt, Germany) between the muscles located to the right of the trachea, whereas the other wired loop lead was sutured to the xiphoid process. Muscle layers and skin were closed with resorbable sutures (Vicryl 6–0; Ethicon). Together with pain relief (twice daily: buprenorphine 0.1 mg/kg and meloxicam 5 mg/kg), supportive therapy consisting of 300 µL glucose (5%) and 300 µL saline (0.9%) warmed to body temperature was applied subcutaneously twice daily for four days.¹² After transmitter implantation, the mice had a 5–6 week period of convalescence before the first experiment.

Experimental setting

Data were acquired in the regular housing room using telemetric and behavioural recordings from undisturbed animals in their home cages during the 24 h following anaesthesia. To avoid interference, all necessary husbandry and management procedures were completed in the animal room at least one day before starting each experiment or data recording, and disturbances (such as unrelated experimental procedures) were not allowed.

Prior to the experiment the mice were placed individually in a new cage for three days to habituate. The cage was equipped with clean sawdust bedding and one Nestlet™. The cardboard house and hay were omitted to allow a better view of the animals during the experiments and for video recordings. Throughout the experiments, the mice stayed in the same cage; this cage was not cleaned and the bedding was not changed during the experiments. All experiments were conducted when the mice were aged 16–32 weeks with body weights ranging from 25 to 30 g. To avoid any influences of circadian rhythm, anaesthetic procedures were carried out between 13:00 and 15:00 h (= at the end of the dark phase). Anaesthesia was performed in a separate experimental area within the animal room to avoid transportation and thus to allow continuous telemetric data recording of the animals in their familiar environment.

Anaesthesia experiments

Information on agents and dosages for anaesthesia and antagonization as well as the administration route were taken from the literature,⁷ and from a study employing the same strain, sex and approximate body weight range of mice as our study.⁴

After weighing the animals, an intraperitoneal injection of fentanyl (0.05 mg/kg, Fentanyl; Kantonsapotheke Zurich, Zurich, Switzerland), midazolam (5 mg/kg, Dormicum®; Roche Pharma AG, Reinach, Switzerland) and medetomidine (0.5 mg/kg, Dorbene®; Graeub AG, Bern, Switzerland) mixed in NaCl (0.9%) to a volume of 3 µL/g body weight was applied. Animals were then returned to their home cages and observed for 7 min to allow time for the mice to calm down in a familiar environment and for the drugs to be absorbed. Time to immobility was established by observing the animals’ spontaneous behaviour and posture. Righting reflex was tested every 10 to 15 s after the animals became motionless by carefully turning the mice on their backs; any attempts by the animals to right themselves from dorsal to sternal recumbence were judged as positive reflex responses. At 7 min after injection of the anaesthetics, the mice were transferred to the experimental area of the animal room. Mice were placed in dorsal recumbence on a water-filled warming mat (TP500; Gaymar, Orchard Park, NY, USA) set at 38 ± 1℃, and protective eye ointment (Vitamin A Blache; Interdelta SA, Givisiez, Switzerland) was applied. Anaesthesia was continued for 50 min while the mice breathed room air spontaneously.

For reflex testing during anaesthesia, noxious stimuli in the form of a tail pinch, and cervical as well as abdominal skin pinch were applied at 10 min intervals. Furthermore, the pedal withdrawal reflex (grasping the skin between the toes of one hind leg and pulling the leg away from the animal), toe pinch (forceps placed on the first phalangeal bone of one hind leg and slight pressure applied) and whisker movements (gently brushing whiskers with forceps) were tested at the same time intervals. All reflex tests were induced by the same investigator using blunt forceps with a spacer between its arms. The reflex tests were registered as positive or negative, i.e. whether any motor responses or reactive movements were observable or not.

Respiratory rate (RR) was assessed by counting the movements of the thorax wall and recorded at 10 min intervals. During anaesthesia, mice were observed for respiratory rhythm abnormalities (i.e. irregularities, pauses, etc.). In addition, they were monitored for alterations in heart rhythm using a real-time telemetry electrocardiogram (ECG) read-out.

In half of the animals, anaesthesia was stopped after 50 min by intraperitoneal injection of naloxone (1.2 mg/kg, NaloxonOrPha; OrPha Swiss GmbH, Kusnacht, Switzerland), flumazenil (0.5 mg/kg, Anexate®; Roche Pharma AG) and atipamezole (2.5 mg/kg, Alzane®; Graeub AG) mixed in 0.5 mL of NaCl (0.9%) to a volume of 9 µL/g body weight. In the remaining animals, 9 µL/g body weight of saline was injected, i.e. anaesthesia was not antagonized. All animals were returned immediately to their respective home cages where either telemetric or video recordings were performed continuously for the next 24 h. The time elapsed to the regaining of righting reflex and spontaneous locomotion was determined from the video recordings.

Telemetric data acquisition and analysis

Telemetric data were recorded using the Dataquest LabPRO program (Data Sciences International). Data acquisition started three days before anaesthesia and continued for 24 h following the 50 min period of anaesthesia. To establish normal values (in the three days before anaesthesia) and to investigate the post-anaesthetic effects (during the 24 h following anaesthesia), HR and core BT were measured in the free-ranging mouse in its home cage every 5 min for 30 s and 10 s, respectively.

To determine the intra-anaesthetic effects of anaesthesia, bipolar ECG and core BT were monitored continuously. ECG tracings were used both to monitor heart rhythm for alterations and to calculate HR (one recording every 10 s). From these data, the mean values of HR and core BT were calculated for each minute for each animal.

Acid–base balance and blood gas concentration

Arterial blood was sampled at 50 min of anaesthesia to investigate the intra-anaesthetic side-effects of anaesthesia on the partial pressure of oxygen (PaO₂, mmHg) and carbon dioxide (PaCO₂, mmHg) as well as on acid–base balance (pH), haematocrit and blood glucose concentration.

Blood sampling and analyses were carried out as described previously.¹³ Briefly, following incision of the anterior neck, dissection of the right common carotid artery, and cutting of a small hole in the artery using a fine-bladed pair of scissors, arterial blood was collected in a heparinized syringe. Acid–base balance, PaCO₂, PaO₂, haematocrit and concentration of blood glucose were determined immediately using a blood gas analyzer (Epoc; Alere Inc, Ottawa, ON, Canada). These animals died immediately from the subsequent rapid blood loss under anaesthesia.

Video recordings and behavioural analysis

In order to establish baseline values as well as to analyse behaviour in mice with or without reversal of anaesthesia, the animals were recorded digitally in the absence of a human observer, 24 h before and after the anaesthesia experiment, using infrared-sensitive cameras suspended 1.5 m above the cages. The recorded material (24 h of continuous footage) was subsequently analysed using EthovisionXT 7 software (Noldus, Wageningen, The Netherlands) measuring the distance moved (in cm) from the animals’ centre point.

In order to determine the temporal distribution of changes in general activity, the 24 h period was divided into four consecutive 6 h periods according to the light–dark cycle in the animal room. Data were summed and analysed for the following time frames: 00:00–06:00 h (light phase), 06:00–12:00 h (light phase), 12:00–18:00 h (dark phase) and 18:00–24:00 h (dark phase).

Latency to drink, eat and build nests was also determined, using ObserverXT™ software (Noldus). These time points were defined as the first purposeful contact with a water nipple, food-pellet or Nestlet™.

Statistical analysis

Data are presented as mean ± standard deviation, except for the post-anaesthetic telemetry recordings, which are presented as boxplots for every hour during the 24 h observation period.

Statistical analyses were performed using SPSS 22 software (IBM, Armonk, NY, USA). All data were tested for normal distribution and homogeneity of variance (Shapiro–Wilks, Levene’s test).

One-way analysis of variance (ANOVA) was performed to compare group means of latency times and activity levels defined as distance moved. Post hoc analysis with Bonferroni tests was carried out to identify significant differences between groups; P values ≤ 0.05 were considered significant.

Results

Anaesthesia period

Following intraperitoneal injection of the anaesthetics, animals were motionless at 210 ± 42 s and lost their righting reflex at 330 ± 47 s.

Seven minutes after injection (i.e. at the beginning of the 50 min anaesthesia), reflex testing (Table 1) was able to induce motor responses in 100% of the animals for toe pinch, and in 31.2% and 25% of the animals for tail pinch and pedal withdrawal reflex, respectively. During the course of anaesthesia, responses diminished subsequently; however, toe and tail pinch responsiveness were not excluded reliably at any time during anaesthesia.

Table 1.

Reflex responsiveness during 50 min anaesthesia (n = 32 mice).

	Anaesthesia duration (min)
	0	10	20	30	40	50
Righting reflex	0/32	0/32	0/32	0/32	0/32	0/32
Tail pinch	10/32	8/32	4/32	6/32	4/32	6/32
Pedal withdrawal reflex	8/32	2/32	2/32	0/32	0/32	0/32
Toe pinch	32/32	32/32	32/32	32/32	28/32	28/32
Abdominal skin pinch	0/32	0/32	0/32	0/32	0/32	0/32
Neck skin pinch	0/32	0/32	0/32	0/32	0/32	0/32
Whiskers reflex	0/32	0/32	0/32	0/32	0/32	2/32

Reflex testing starting at time point 0 of anaesthesia (i.e. 7 min after injection) was performed every 10 min. Fraction of the responsiveness (i.e. motor reaction present) over all animals tested is presented. Responsiveness upon toe and tail pinch was not reliably excluded at any time point during anaesthesia.

During anaesthesia HR averaged 225 ± 23 beats/min and core BT reached steady state at 34.3 ± 0.25℃ (Figure 1). RR ranged from 96 to 180 breaths/min with a mean of 131 ± 15 breaths/min in all the animals. Analysis of arterial blood from eight mice at 50 min of anaesthesia showed the following values: PaO₂ 83.6 ± 8.1 mmHg, PaCO₂ 43.9 ± 6.1 mmHg, pH 7.16 ± 0.04, concentration of blood glucose 22.1 ± 2.5 mmol/L, haematocrit 46.3 ± 3.9%.

Figure 1.

Heart rate (HR) and core body temperature (BT) during induction and anaesthesia. Mean ± SD (n = 16 mice) for HR and core BT after injection of fentanyl–midazolam–medetomidine in the home cage and during 50 min of anaesthesia while mice breathed spontaneously and lay in dorsal recumbency on the warming mat (38℃). Grey areas indicate mean baseline values (measured for 3 days before anaesthesia) at the corresponding time of day in conscious mice.

Post-anaesthetic period

Upon completion of the 50 min anaesthesia period, animals received an injection of either antagonists or saline and were returned to their respective home cages. After reversal of anaesthesia, HR and core BT returned to baseline values within 2 h, whereas mice that received no reversal went through a 16–24 h period of acute hypothermia and bradycardia, reaching average values of 22 ± 1.9℃ and 119 ± 69 beats/min, respectively (Figure 2a). After antagonization, animals regained physiological posture after 1.9 ± 0.3 min and locomotor activity returned after 4.3 ± 0.5 min. However in mice without antagonization, a further 726 ± 36 min and 792 ± 42 min passed before these animals were able to resume their physiological posture and be ambulant (Figure 3). All the animals survived.

Figure 2.

Post-anaesthetic period (24 h). (a) Telemetric recordings of heart rate (HR) and core body temperature (BT) for 24 h during baseline and after anaesthesia with and without reversal are presented as boxplots for each hour (n = 8 mice). (b) Activity shown as distance moved during 24 h. A single representative data-set of locomotor activity levels (represented as distance of animals’ centre point moved in centimetres) during baseline as well as after anaesthesia with or without reversal as analysed by Ethovision® software is presented.

Figure 3.

Duration until recovery of physiological posture (righting reflex) and spontaneous locomotion. Time elapsed after anaesthesia is depicted in minutes per animal (n = 8 mice per group).

Activity levels were measured (Figure 2b) and analysed during four consecutive 6 h periods (00:00–06:00 h and 06:00–12:00 h during the light phase; 12:00–18:00 h and 18:00–24:00 h during the dark phase) (Figure 4). In the first 6 h after anaesthesia, all mice displayed a significant reduction in locomotor activity (measured as distance moved) when compared with baseline values (P = 0.014 compared with animals with antagonization and P = 0.001 compared with animals without antagonization). During the next 6 h period (6–12 h after treatment, light period), the group without antagonization displayed the lowest levels of activity, whereas the group that received antagonization was even more active than the corresponding baseline (P = 0.0001 compared with animals with antagonization and P = 0.007 compared with animals without antagonization). During the first 6 h of the dark period (12–18 h after anaesthesia), the activity levels of both experimental groups were significantly lower than the corresponding baseline (P = 0.0001). This trend continued to the end of the dark phase (18–24 h after anaesthesia), with activity levels of experimental groups below baseline levels (P = 0.015 compared with animals with antagonization and P = 0.138 compared with animals without antagonization).

Figure 4.

Home cage activity represented as distance moved in centimetres for 0–6 h, 6–12 h, 12–18 h and 18–24 h. Mean ± SD (n = 8 mice). Activity was in general decreased on the first day after anaesthesia, except at 6–12 h after antagonization. P values are shown for the comparison with baseline; *P ≤ 0.05; **P = 0.0001.

Latency to engage in spontaneous home cage behaviours such as eating, drinking and nest building were not altered significantly when anaesthesia was reversed (Figure 5). Animals that received no reversal of anaesthesia displayed significant latency prolongation in eating (P = 0.0001 and P = 0.0001 compared with baseline and animals with antagonization, respectively, while there was no significant difference between baseline and animals with antagonization, P = 0.51). The same effect was seen in drinking (P = 0.0001 and P = 0.0001 compared with baseline and animals with antagonization, respectively, while there was no significant difference between baseline and animals with antagonization, P = 0.051), and nest building (P = 0.0001 and P = 0.0001 compared with baseline and animals with antagonization, respectively, while there was no significant difference between baseline and animals with antagonization, P = 0.081).

Figure 5.

Latency to eat, drink and build nests during the post-anaesthetic period. Mean ± SD for baseline and time elapsed after the 50 min anaesthesia period until animals started eating, drinking and building nests (n = 8 mice); *P ≤ 0.05.

Discussion

Intraperitoneal injection of fentanyl–midazolam–medetomidine induced anaesthesia for up to 50 min in adult female C57BL/6 mice. Induction of anaesthesia was rapid; however a decrease in response to reflex testing was delayed in a few animals, as with the pedal withdrawal reflex (often considered to be a sign of deep anaesthesia). Whereas in most animals the pedal withdrawal reflex was negative at 10 min, it was still positive at 20 min in two out of 32 mice, which probably resulted from individual variability of pharmacodynamics and sensitivity to the agents.¹³ In general, during anaesthesia no response could be induced from most of the reflexes tested, with the exception of the pedal withdrawal reflex, toe pinch and tail pinch. The reason for the latter reactions could be that anaesthesia was not deep enough as a result of a low dosage. The textbook dosages used⁷ were identical to those in another study using the same strain, sex and approximate body weight of mice.⁴ Because both anecdotal evidence and clinical experience support the suggested dosages, no preliminary dose testing was deemed necessary to adjust the commonly proposed dosages to the conditions in our laboratory setting (e.g. strain, sex, age, etc.) as generally recommended for injection anaesthesia in mice.¹³ Moreover, clinical experience with widespread use of this kind of anaesthesia (sometimes called ‘neuroleptanalgesia’) suggests that, due to the properties of the drugs, a specific reflex might not in fact be indicative of the depth of anaesthesia. However, no further focused investigations were carried out to ascertain the exact stage of anaesthesia reached in our study. Thus, given our findings, we conclude that, due to the inconsistent loss of reflexes and elimination of response upon noxious stimulation, the dosages might need to be adapted, particularly if the protocol is used for invasive or painful interventions.

During anaesthesia, stable, albeit depressed, HR and RR were observed. Arterial blood gas analysis detected hypoxia, hypercapnia, acidosis and hyperglycaemia. These characteristics should be considered when measurements for cardiovascular studies or of glucose metabolism are performed under anaesthesia. In mice, such typical intra-anaesthetic alterations of physiological equilibria can hardly be prevented except hypoxia, which could be treated by providing additional oxygen in breathing air.

Intraperitoneal injection of the antagonists naloxone–flumazenil–atipamezole resulted in a clear reversal of anaesthesia within a few minutes. Animals regained all reflexes, turned in sternal recumbence and showed normal posture, movement and locomotion. Spontaneous home cage behaviours such as food and water consumption as well as nest building were not altered significantly after reversal of anaesthesia compared with unanaesthetized mice. While locomotion appeared within a few minutes of antagonization, monitoring of locomotor activity with continuous 24 h infrared-sensitive video recordings revealed an overall decrease in locomotor activity, i.e. a shorter distance was covered in the day following anaesthesia with antagonization. In addition, the circadian rhythmicity of locomotor activity was clearly disturbed, as shown by the abnormal distribution of activity bouts in contrast to the regular distribution of activity bouts relative to daytime (i.e. high activity during the light phase and low activity in the dark phase). The aberrant locomotion periods during daytime hinted at reduced sleep times and possible restlessness. After antagonization, HR as well as core BT quickly regained normal values. A slight decline in core BT to values of 33–34℃ was found at one hour after antagonization. Such a decrease in BT after reversal of anaesthesia was also reported previously by Thal and Plesnila⁴ from measurements using a rectal probe. With telemetric measurements we were able to confirm that this decrease in core BT occurred for less than an hour and was thus transient and probably of minor influence on animal well-being.

By contrast, if antagonization was omitted, severe hypothermia and bradycardia occurred for about 16 h or more after the 50 min period of anaesthesia. Locomotor activity was almost absent for about 12 h. Locomotor activity bouts appeared from 12 h onwards but overall activity was reduced and did not reach normal levels even by 24 h. In addition, the time until animals resumed spontaneous home cage behaviours such as food and water consumption as well as nest building was markedly prolonged to 15 and 20 h, respectively. Such extension and aggravation of anaesthesia-related side-effects were probably in part due to the methodology used, i.e. the experimental set-up, which aimed to treat both groups identically with continuous sampling of telemetry and video recording data for one day. Therefore, animals were transferred back to their familiar home cage environment without further heat support, which was of negligible relevance if anaesthesia was reversed but was clearly disadvantageous for those that did not receive antagonization. This underlines the well-known aspects and importance of perioperative care in mice, particularly the provision of warmth during recovery from injection anaesthesia, which is routinely applied in laboratory work.

In summary, reversal of anaesthesia was advantageous and is recommended, as the recovery phase was shortened markedly by antagonization and the adverse effects of the anaesthesia were reduced to an acceptable level. In this study, all substances administered in the anaesthesia mixture were antagonized hence the analgesic properties of these agents were completely eliminated when the anaesthesia was reversed. Since no invasive or painful intervention was conducted in this experiment, the antagonization of analgesic effects was justified from animal welfare and methodological considerations. However, if surgery or otherwise persistent pain-inducing interventions are performed under anaesthesia, post-operative pain must be considered. Consequently, we suggest that either (i) the opioid fentanyl should not be antagonized with naloxone, or (ii) buprenorphine should be administered at antagonization to provide post-operative pain alleviation for several hours.¹⁴ Since the presented protocol (fentanyl–midazolam–medetomidine, antagonized with naloxone–flumazenil–atipamezole) comprises reversal of all anaesthetic and analgesic components, it might be useful in situations where no post-anaesthetic pain is present and rapid recovery is aimed for. Therefore, for minor interventions, for less invasive measurements, and for investigations needing immobilization such as imaging procedures, this protocol might be preferable to other injection anaesthesia protocols (e.g. well-known ketamine-based mixtures with alpha-two agonists).

Conclusions

Intraperitoneal injection of fentanyl–midazolam–medetomidine provides reliable anaesthesia for up to 50 min in laboratory mice, with stable although depressed cardiovascular parameters together with slight hypoxia, hypercapnia, acidosis and hyperglycaemia. Dosages of all components of anaesthetic mixtures should be adjusted to the specific conditions of the experiment (e.g. strain, sex, age of mice, etc.), particularly if anaesthesia is applied for conducting painful interventions (e.g. major surgery) requiring a sufficient depth of anaesthesia and a combination with analgesics. Anaesthesia should be reversed with flumazenil–atipamezole whenever the experimental approach allows. Where this is not the case, extra care should be taken, especially regarding heat support and possibly oxygen support.

Footnotes

Acknowledgements

The authors would like to thank Robin Schneider, Hugo Battaglia, and the staff of the Central Biological Laboratory for their valuable support.

Declaration of conflicting interests

The author(s) declared no potential conflicts of interest with respect to the research, authorship, and/or publication of this article.

Funding

The author(s) received no financial support for the research, authorship, and/or publication of this article.

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