Animals
Nine male Wistar rats with a mean body weight of 275 ± 24 g were obtained from Charles River Laboratories, Sulzfeld, Germany. They were housed in groups of three in a Makrolon® cage (Type IV). The cages contained a wooden bedding material (Lignocel select fine, J. Rettenmaier & Söhne GmbH + Co. KG, Rosenberg, Germany) and as cage enrichment a wooden chewing block and two red, transparent plastic tubes were provided. The cages were changed twice per week. Rats were given free access to a commercially available diet (3438 maintenance diet, KLIBA NAFAG, Provimi Kliba AG, Kaiseraugst, Switzerland) and tap water. The animals were acclimated for two weeks prior to surgical implantation of the radiotelemetry transmitter in an animal room maintained at 22 ± 2°C and 55 ± 10% relative humidity, on a twelve-hour light–dark cycle (beginning at 6:00 am) with at least 15 air changes per hour. A radio was turned on during the working hours to reduce possible stress caused by environmental noises. After the implantation surgery and a rehabilitation period of at least two weeks, rats were used in a previous anaesthetic study, which lasted nine weeks [9]. The rats were then given a recovery period of at least three weeks prior to the start of the present study. At the beginning of the present study, the rats had a mean body weight of 434 ± 48 g.
Radiotelemetry transmitter implantation surgery
The radiotelemetry transmitter (DSI PhysioTel™ C50-PXT) was implanted under general anaesthesia. Anaesthesia was induced with MMF (same dosage as used in this study) and was maintained with a second injection of MMF (one third of the initial dosage) after 45 minutes. Analgesics and antibiotics were administered prior to surgery: 50 mg·kg−1 metamizole i.m. (Novalgin®, 500 mg·ml−1, Sanofi Aventis, Frankfurt/Main, Germany), 1 mg·kg−1 meloxicam s.c. (Metacam®, 20 mg·ml−1, Boehringer Ingelheim, Ingelheim/Rhein, Germany), 10 mg·kg−1 enrofloxacin s.c. (Baytril® 2.5% ad us. vet., Bayer, Leverkusen, Germany). After induction of anaesthesia, protective eye lubricant (VitA-POS, Ursapharm, Saarbrücken, Germany) was administered in both eyes, the rat was laid in a supine position in the middle of a water heating pad, shaved on the ventral site and desinfected with Kodan®-spray (Schülke & Mayr, Norderstedt, Germany) and Betaisodona®-solution (Mundipharma GmbH, Limburg (Lahn), Germany). Supplemental Oxygen was provided via a head chamber throughout the anaesthesia. The rat was covered with a sterile foil and an incision was made along the linea alba to open the abdominal cavity. The intestines were moved towards the diaphragm and held in position with a wet swab. The aorta was exposed and blood flow was stopped with two vascular clips. The blood pressure catheter was inserted between the two clips and fixed with one drop of tissue glue (Histoacryl®, B.Braun, Aesculap AG, Tuttlingen, Germany). The clips were carefully opened and the swab was removed. The transmitter was fixed with a permanent suture (Mersilene® 3–0, Ethicon®, Johnson-Johnson Medical GmbH, Norderstedt, Germany) to the abdominal wall. Before the abdominal cavity was closed with a muscle and skin suture (Vicryl® 3–0, Ethicon®, Johnson-Johnson Medical GmbH, Norderstedt, Germany), the ECG leads were exteriorized through the muscle layer. One ECG lead was placed subcutaneously near the sternum and the other was placed on the ventral site of the trachea, both fixed to the nearby muscle tissue with a non-absorbable suture (Mersilene® 3–0, Ethicon®, Johnson-Johnson Medical GmbH, Norderstedt, Germany). Twenty ml·kg−1 warmed lactated ringer’s solution (Ringer-Lactat nach Hartmann B. Braun, B/BRAUN, Melsungen, Germany) were administered subcutaneously and anaesthesia was reversed with a s.c. injection of AFN (same dosage as used in this study). After surgery the rats received analgesics for another two days.
Experimental design
The following three anaesthetic regimes were performed repeatedly in male Wistar rats: 1) an inhalational anaesthesia with 2–3 Vol% isoflurane (Forene® 100% (V/V), Abbott, Wiesbaden, Germany) at maintenance, terminated after 40 minutes, 2) a combination of 100 mg · kg−1 ketamine (Ketavet®, 100 mg·ml−1, Pfizer, Berlin, Germany) and 5 mg·kg−1 xylazine (Rompun® 2%, 20 mg·ml−1, Bayer, Leverkusen, Germany) administered i.m., which was not reversed, and 3) an anaesthesia consisting of 0.15 mg·kg−1 medetomidine (Domitor, 1 mg·ml−1, Orion Pharma, Espoo, Finland), 2 mg·kg−1 midazolam (Dormicum®, 5 mg·ml−1, Roche, Grenzach-Wyhlen, Germany) and 0.005 mg·kg−1 fentanyl (Fentanyl®-Janssen, 0.05 mg·ml−1, Janssen, Wien, Austria), which was administered i.m. and antagonized after 40 minutes with a s.c. injection of the antagonists 0.75 mg·kg−1 atipamezole (Antisedan®, 5 mg·ml−1, Orion Pharma, Espoo, Finland), 0.2 mg·kg−1 flumazenil (Flumazenil Hexal®, 0.1 mg·ml−1, Hexal, Holzkirchen, Germany) and 0.12 mg·kg−1 naloxone ( Naloxon Inresa, 0.4 mg·ml−1 l, Inresa, Freiburg, Germany). Anaesthesias were performed twice weekly (Monday + Thursday or Tuesday + Friday) over three consecutive weeks. The six anaesthesias were described in the following text as run 1 to run 6. The anaesthetic regimes were assigned randomly to each individual rat. Each rat received only two of the three anaesthetic treatments for six times with a wash out and recovery period of at least two weeks between them. SAP, DAP, MAP, PP, HR and BT were measured continuously before, during and after each anaesthesia. To monitor the anaesthetic depth, the righting reflex and the pedal withdrawal reflex on the fore- and hind limbs were measured (classified with +, ±, and -) 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. One run was divided into different intervals: 1) an acclimatization period prior to the anaesthesia of at least two hours for assessing baseline values, 2) the performance of anaesthesia, and 3) a recovery period. To analyse alterations in the duration of the anaesthetic effect, the anaesthesia period was further divided in different anaesthetic stages: 1) induction time (defined as the time from application to loss of the righting reflex), 2) time of non-surgical tolerance (defined as the time from loss of the righting reflex until loss of all reflexes tested), 3) time of surgical-tolerance (defined as the time from loss of all reflexes tested until regaining of at least one reflex), 4) wake-up period (defined as the time from regaining the first reflex until regaining the righting reflex) , and 5) recovery period (defined as the time from regaining the righting reflex until measurement was terminated). The end of measurement was not earlier than six hours after induction. Sleeping times longer than six hours occurred with KX, therefore data assessment was prolonged and terminated not earlier than two hours after the righting reflex had returned. Additionally, each rat was weighed to calculate the individual dose of injectable anaesthetics and to assess a possible influence of repeated anaesthesia on BW. Daily monitoring of the animals’ general condition and well-being were carried out, especially with regard to inflammation and tissue necrosis at the hind legs after injection of anaesthetics. Only visible changes (open wounds, lameness) were assessed, because the animals were used in further studies.
The Animal Care and Ethics Committee of the Regional Authority in Tuebingen, Baden-Wuerttemberg, Germany approved this study (Approval number: 12–038).
Procedure
The experimental set-up was the same as that used in the previous study [9]. To reduce variability, anaesthesias were performed by the same veterinarian supported always by the same animal care takers, who were familiar with the animals. To ensure a thorough monitoring of each rat during anaesthesia, not more than three rats were anaesthetized per day. The measurements have been carried out in the same laboratory as described below:
The implanted radiotelemetry transmitters of the three rats were switched on using a magnet before starting the measurements. The rats were then placed individually into a Makrolon® cage, each positioned directly on a receiver plate, and continuous data collection of arterial blood pressure and BT was started. The cages contained bedding material (Lignocel select fine, J. Rettenmaier & Söhne GmbH + Co. KG, Rosenberg, Germany), a red, transparent plastic tube and water was provided in a water bottle during the entire measurement. Following recommendations to pre-warm animals prior to anaesthetic induction, a warm water heating pad was placed between the receiver plate and the Makrolon® cage and it was maintained at 38°C throughout the measurement period [15],[16]. To facilitate reaching resting baseline values, rats were housed individually, food was withdrawn, each cage was covered with a cloth, a radio was switched on (because rats were habituated to background music), and the operator left the room for at least two hours. After this acclimatization period the operator started with the anaesthetic induction of the first rat, while the other two rats remained in their covered cages. Performing anaesthesia on one rat did not appear to have an influence on the measured parameters of the other two. All anaesthesias were carried out close to the receiver plate to ensure the continuous capture of the telemetric signal and are described in detail below:
ISO: An anaesthetic chamber was prefilled with 5 Vol% ISO supplemented with oxygen. The rat was placed in the whole body chamber, which was positioned directly on the receiver plate. Loss of righting reflex was tested by tipping over the anaesthetic chamber. When the righting reflex was lost, the animal was laid on its back on the receiver plate and ISO was administered using a nose cone. The concentration of ISO was reduced to 2–3 Vol% to maintain an anaesthetic depth of surgical tolerance. ISO administration was terminated 40 minutes after induction of anaesthesia and the rat was laid back in its cage, which was placed on the receiver plate again. The rat was positioned in a dorsal recumbency to test reflexes until regaining the righting reflex.
KX: Ketamine (100 mg·kg−1) and xylazine (5 mg·kg−1) were mixed in one syringe with a total volume of 1.25 ml·kg−1, which is too much to be injected all at once. Therefore, half of this volume was administered in the caudal part of the thigh muscle of each hind leg. As soon as the righting reflex was lost, the rat was placed directly on the receiver plate in dorsal recumbency. Oxygen was supplemented using a head chamber. Rats received no treatment to reverse the anaesthesia. Therefore, the duration of the sleeping time was variable. Accordingly, rats were laid on their backs on the receiver plate until the righting reflex returned and after that they were put back in their cage for the rest of the measurements. Wake-up and recovery periods lasted several hours and therefore prevented a normal food and water uptake. Therefore, each rat was administered s.c. 5 ml of warmed lactated ringer’s solution (Ringer-Lactat nach Hartmann B. Braun, B. Braun, Melsungen, Germany) one hour after induction of anaesthesia.
MMF: 0.15 mg·kg−1 medetomidine, 2 mg·kg−1 midazolam and 0.005 mg·kg−1 fentanyl were combined in one syringe with a total volume of 0.65 ml · kg−1. The mixture was administered i.m. in the caudal part of the thigh muscle of one hind leg while holding the rat next to the receiver plate. The rat was put back in its cage until loss of the righting reflex. The rat was placed directly on the receiver plate in dorsal recumbency and oxygen was provided using a head chamber. A mixture of antagonists (0.75 mg·kg−1 atipamezole + 0.2 mg·kg−1 flumazenil + 0.12 mg·kg−1 naloxone) was administered s.c. 40 minutes after induction of the anaesthesia and the rat was placed in dorsal recumbency in its cage to determine the time when the righting reflex was regained.
Statistical analysis
For the continuous telemetric data acquisition the software NOTOCORD-hem™ was used and the raw data were further evaluated with MS Excel with subsequent export to SAS 9.3 (SAS Institute Inc., Cary, North Carolina, USA) for the statistical evaluation. The statistical analysis was performed separately for each treatment group (ISO, KX, MMF), parameter (SAP, DAP, MAP, PP, HR, BT, BW) and anaesthetic interval (induction time, time of non-surgical tolerance, time of surgical tolerance, wake-up period, recovery period). The standardized area under the curve (AUC divided by interval length) was calculated for each animal individually using the trapezoidal rule for each anaesthetic interval. Baseline values were calculated for each animal as the standardized AUC including the measurements from 60 to 10 minutes before induction of anaesthesia. The standardized AUC was then used as target variable for the statistical evaluation. The differences between the 1st run and its consecutive runs were analysed. An analysis of covariance (ANCOVA) for repeated measurements was calculated including baseline as covariate. The parameter body weight was analysed by an analysis of variance (ANOVA). In addition, the length of the intervals was analysed using an ANOVA model for repeated measurements. To exclude effects of repeated anaesthesia on baseline values, parameters were analysed during the baseline interval using an analysis of variance (ANOVA). Treatment effects for the telemetry parameters were quantified by mean differences, based on the adjusted mean values and their two-sided 95% confidence interval. Treatment effects for body weight and the length of the intervals were quantified by the differences in mean and their two-sided 95% confidence interval. The level of significance was fixed at α = 5%. A p-value of less than 0.05 was considered to be statistically significant.