Twelve, male Wistar rats with a mean body weight of 287 ± 30 g were acquired from a commercial breeder (Charles River Laboratories, Sulzfeld, Germany). These animals were housed in groups of three in a Makrolon® cage (Type IV) containing a wooden bedding material (Lignocel select fine, J. Rettenmaier & Söhne GmbH + Co. KG, Rosenberg, Germany). Cage bedding changes were performed twice weekly. Two red, transparent plastic tubes, nesting material and a wooden chewing block were provided in each cage for animal enrichment. The rats received a commercially available diet (3438 maintenance diet, KLIBA NAFAG, Provimi Kliba AG, Kaiseraugst, Switzerland) and tap water ad libitum. The animal room was maintained at 22 ± 2°C and 55 ± 10% relative humidity and there was an air change of at least 15 cycles/hour. Light was on from 6:00 am to 6:00 pm, starting and ending with a dimmer-period of 30 minutes. Together with the light, a radio was turned on. The rats were allowed to acclimate to the housing conditions and the husbandry procedures for at least two weeks prior to the surgical implantation of the radiotelemetry transmitter (see below).
Implantation of the radiotelemetry transmitter
The implantation of the transmitter (DSI PhysioTel™ C50-PXT) was performed under general anaesthesia using MMF in the same dosage as described below in this study. One third of the initial dosage was administered again after 45 minutes in order to maintain anaesthesia. For analgesia, the rats received prior to surgery 50 mg•kg−1 metamizole i.m. (Novalgin®, 500 mg•ml−1, Sanofi Aventis, Frankfurt/Main, Germany) and 1 mg•kg−1 meloxicam s.c. (Metacam®, 20 mg•ml−1, Boehringer Ingelheim, Ingelheim/Rhein, Germany). Metamizole application was repeated three times until the next day. To prevent a bacterial infection, 10 mg•kg−1 enrofloxacin (Baytril® 2.5% ad us. vet., Bayer, Leverkusen, Germany) was administered subcutaneously after the analgesic injections. As soon as the rats lost their righting reflex, the ventral site of the rat was shaved and then disinfected with Kodan®-spray (Schülke & Mayr, Norderstedt, Germany) and Betaisodona®-solution (Mundipharma GmbH, Limburg (Lahn), Germany). Protective eye lubricant (VitA-POS, Ursapharm, Saarbrücken, Germany) was administered to both eyes, loss of body temperature was minimized using a warm water heating pad and supplemental oxygen was provided through a nose cone. Once a surgical level of anaesthesia was confirmed through the loss of reflexes, an incision was made in the linea alba from below the sternum to the umbilical region to open the abdominal cavity. The intestinal tract was carefully repositioned using a moist swab in a cranial direction to exposure the aorta abdominalis which was dissected free for insertion of the blood pressure catheter of the telemetry unit. Blood flow was temporarily stopped using two vascular clips and the catheter was inserted between them and fixed in place with tissue glue (Histoacryl®, B.Braun, Aesculap AG, Tuttlingen, Germany). Clips and the swab were then removed and the transmitter was sutured to the abdominal wall (Mersilene® 3–0, Ethicon®, Johnson-Johnson Medical GmbH, Norderstedt, Germany). The ECG leads were exteriorized through the abdominal muscle layer and placed subcutaneously, one to the end of the sternum and the other to the ventral region of the trachea. The ECG leads were sutured to the nearby muscle tissue (Mersilene® 3–0, Ethicon®, Johnson-Johnson Medical GmbH, Norderstedt, Germany). The abdominal cavity was closed in layers with first a muscle and then a skin suture (Vicryl® 3–0, Ethicon®, Johnson-Johnson Medical GmbH, Norderstedt, Germany). At the end of the procedure the rats received 20 ml•kg−1warmed lactated ringer’s solution (Ringer-Lactat nach Hartmann B. Braun, B. Braun, Melsungen, Germany) subcutaneously. The anaesthesia was then antagonized with a subcutaneous injection of AFN (same dosage as used in this study, see below). The whole implantation procedure took 90 minutes on average. Meloxicam was administered once-daily for two additional days. The rats were allowed two weeks to recover from this surgical procedure before entering the study.
The three different anaesthetic treatments were evaluated using a randomized, crossover design with each rat receiving each of the following three anaesthetic treatments on different days: 1) an inhalational anaesthesia with isoflurane for 40 minutes (Forene® 100% (V/V), Abbott, Wiesbaden, Germany), 2) a combination of ketamine (Ketavet®, 100 mg•ml−1, Pfizer, Berlin, Germany) and xylazine (Rompun® 2%, 20 mg•ml−1, Bayer, Leverkusen, Germany) administered intramuscularly and 3) a combination of medetomidine (Domitor, 1 mg•ml−1, Orion Pharma, Espoo, Finland), midazolam (Dormicum®, 5 mg/ml, Roche, Grenzach-Wyhlen, Germany) and fentanyl (Fentanyl®-Janssen, 0.05 mg•ml−1, Janssen, Wien, Austria) administered intramuscularly and reversed after 40 minutes with a subcutaneous injection of atipamezole (Antisedan®, 5 mg•ml−1, Orion Pharma, Espoo, Finland), flumazenil (Flumazenil Hexal®, 0.1 mg•ml−1, Hexal, Holzkirchen, Germany) and naloxone (Naloxon Inresa, 0.4 mg•ml−1, Inresa, Freiburg, Germany). With KX anaesthesia the rats received no treatment to reverse the anaesthesia. The rats were allowed a recovery period of two weeks between the anaesthesias. To allow for a thorough monitoring of each individual rat during anaesthesia, only three anaesthesias were performed per day. SAP, DAP, MAP, PP, HR and BT were continuously measured from the start of measurements (6.00 am) until the end of measurements (~5 pm). A complete measurement consisted of a 2–4 hour pretreatment acclimatization period for reaching baseline values, the time for performing anaesthesia (a duration of 40 minutes was chosen for ISO and MMF anaesthesia; KX anaesthesia lasted in some animals up to 395 minutes, because it was not reversed), a wake-up and a recovery period. All anaesthesias were performed by the same veterinarian to reduce variability.
The experimental procedures were approved by the Animal Care and Ethics Committee of the Regional Authority in Tuebingen, Baden-Wuerttemberg, Germany (Approval number: 12–038).
Before starting the telemetric measurement, each rat was weighed and placed individually into a Makrolon® cage, containing bedding material and a red, transparent plastic tube and covered with a cloth. Single housing and withdrawal of food were required, because rats had come to rest to assess individual baseline values. Tap water was provided in a water bottle during the entire time of data collection. Each cage was placed on a radiofrequency receiver plate. A water heating pad was placed between the cage and the telemetric receiver plate and it was left on (38°C) throughout the study. Data collection started by switching on the radiotransmitter using a magnet, the operator left the room and the animals were given at least two hours to establish resting, baseline conditions. Afterwards, the induction of anaesthesia was started with the first rat. It was of note that this procedure seemed to have no impact on the cardiovascular parameters measured from the other two rats in the room. Their cages were still covered with a cloth and anaesthesia was performed as quiet as possible so that the produced noise did not drown out the radio, which was switched on at the beginning of measurements. Each rat received protective eye lubricant (VitA-POS, Ursapharm, Saarbrücken, Germany) at the beginning of anaesthesia. After induction of anaesthesia, the righting reflex (defined as positive, when a rat, placed on its back/side, immediately turns over to the normal position with all four feet on the ground) and pedal withdrawal reflex on the hind and forelimbs were monitored and classified (+, ±, −) after 2.5, 5, 7.5, 10, 15, 20, 30, 40, 42.5, 45, 47.5, 50, 60, 70… minutes until the righting reflex had returned. The time until loss and later regaining of the righting reflex was determined. Based on the presence or absence of the reflexes assessed, the anaesthesia time was divided into the following intervals: 1) the induction time defined as the time from application of the anaesthetic(s) to loss of the righting reflex, 2) the time of non-surgical tolerance, defined as the time from the loss of the righting reflex until loss of the pedal withdrawal reflexes on the hind and forelimbs, 3) the time of surgical tolerance, defined as the time from the absence of righting and pedal withdrawal reflexes until regaining at least one pedal withdrawal reflex, 4) the wake-up period, describing the time from regaining one pedal withdrawal reflex until regaining the righting reflex and 5) the time of recovery, defined as the time from regaining the righting reflex until the end of measurements. The measurements were terminated not earlier than six hours after induction for all three anaesthesias. Because of long-lasting effects of KX, measurements continued for at least two hours after the rats regained the righting reflex.
The different anaesthetic treatments were performed as described below.
A whole body chamber was prefilled with 5 Vol % ISO in 100% oxygen. The chamber was placed on the transmitter receiver plate and the rat was positioned into the chamber and the time was measured till loss of the righting reflex. To assess the righting reflex, the chamber with the animal was tipped over. The rat was then placed in dorsal recumbency in the middle of the water heating pad and 5 Vol % ISO was further administered using a nose cone. The concentration of ISO was then individually regulated and reduced to 2–3 Vol % for producing a depth of anaesthesia suitable for surgical procedures, which meant that all reflexes tested had to disappear. Forty minutes after induction of anaesthesia, ISO administration was stopped and the rat was put back in its cage on the receiver. The rat was positioned on its back, so that return of the righting reflex could be determined, when the animal turned around to a ventral recumbency.
Ketamine (100 mg•kg−1) and xylazine (5 mg•kg−1) were mixed together in one syringe. The volume of the KX injection was too large (1.25 ml•kg−1) to be administered in one hind leg, therefore, it was divided in half and injected intramuscularly in the caudal parts of the femoral musculature of both hind legs. For a continuous telemetric measurement even during this procedure, the injection was performed next to the receiver plate to assure capture of the telemetric signals. After loss of the righting reflex, the rat was placed on its back in the middle of the heating pad and supplied with 100% oxygen using a nose cone. Due to the fact that the duration of this anaesthesia could not be accurately predicted, the rat stayed on the heating pad until its righting reflex returned. Thereafter, the animal was immediately placed back in its cage located on the receiver. The long sleeping and recovery time in KX anaesthesia necessitated the administration of fluids and therefore all rats received 5 ml of warmed lactated ringer’s solution (Ringer-Lactat nach Hartmann B. Braun, B. Braun, Melsungen, Germany) subcutaneously one hour after induction.
Medetomidine (0.15 mg•kg−1), midazolam (2.0 mg•kg−1) and fentanyl (0.005 mg•kg−1) were mixed in one syringe (total volume: 0.65 ml•kg−1) and were administered intramuscularly in the caudal part of the femoral musculature of one hind leg. After loss of the righting reflex the cage was removed and the rat was placed in dorsal recumbency in the middle of the heating pad and 100% oxygen was provided using a nose cone. The antagonists atipamezole (0.75 mg•kg−1), flumazenil (0.2 mg•kg−1) and naloxone (0.12 mg•kg−1) were mixed in one syringe and administered subcutaneously 40 minutes after induction of the anaesthesia. Thereafter, the rat was returned to its cage in dorsal recumbency to determine the moment of righting.
NOTOCORD-hem™ was used for telemetric data acquisition and data were further evaluated using MS Excel. Values for each parameter over 10 minutes were summarized by the assessment of mean baseline values. Baseline values were calculated as the mean of the measurements starting 60 up to 10 minutes before the induction of anaesthesia. Mean values measured during and after anaesthesia were calculated with medians based on 20 second intervals. For data import from MS Excel the software package SAS 9.2 was used. The statistical evaluation was done using the software package SAS 9.3 (SAS Institute Inc., Cary, North Carolina, USA). The statistical evaluation was done for each parameter measured (SAP, DAP, MAP, PP, HR and BT) and each of the defined intervals (induction time, time of non-surgical tolerance, time of surgical tolerance, wake-up and recovery period). The area under the curve (AUC) was calculated for each animal individually using the trapezoidal rule for the intervals. The different intervals were compared to the baseline interval by an analysis of variance (ANOVA) for repeated measurements for every anaesthesias separately. Effects were quantified by mean differences and their two-sided 95% confidence interval. Additionally, the parameters were analysed by a one-factorial analysis of covariance (ANCOVA) with heteroscedastic variances and the fix factor treatment including the baseline as covariate. The following comparisons were performed by two sided t-tests:
ISO vs. KX,
ISO vs. MMF,
KX vs. MMF.
Treatment effects were quantified by mean differences, based on the adjusted mean values, and their two-sided 95% confidence interval. The level of significance for both analysis was fixed at α = 5%. A p-value less than 0.05 was considered to be statistically significant.