Upload
others
View
7
Download
0
Embed Size (px)
Citation preview
Research Collection
Doctoral Thesis
Optimization of anesthetic protocols and their impact onphysiological and behavioral parameters in laboratory mice
Author(s): Ćesarović, Nikola
Publication Date: 2014
Permanent Link: https://doi.org/10.3929/ethz-a-010124123
Rights / License: In Copyright - Non-Commercial Use Permitted
This page was generated automatically upon download from the ETH Zurich Research Collection. For moreinformation please consult the Terms of use.
ETH Library
Diss. ETH No
21673
Optimization of Anesthetic Protocols and their Impact on
Physiological and Behavioral Parameters in Laboratory Mice
A dissertation submitted to
ETH ZURICH
for the degree of
Doctor of Sciences
presented by
NIKOLA ĆESAROVIĆ
Dr. med. vet.
Vetsuisse Faculty, University of Zürich
Date of birth
10.07.1980
citizen of
Belgrade, Republic of Serbia
accepted on the recommendation of
Professor Dr. W. Langhans, examiner
Professor Dr. B. Beck Schimmer, co-examiner
PD Dr. M. Arras, co-examiner
2014
Table of Contents
I
Table of Contents
List of Tables and Figures .................................................................................................... VI
List of Abbreviations ........................................................................................................... VIII
Summary ............................................................................................................................. XII
Zusammenfassung ............................................................................................................. XIV
Chapter 1: General Introduction ...........................................................................................17
General anesthesia in small laboratory rodents .................................................................18
Inhalation anesthesia in laboratory routine .....................................................................19
Isoflurane .......................................................................................................................20
Sevoflurane ....................................................................................................................20
Minimum Alveolar Concentration (MAC)............................................................................21
Balanced anesthesia .........................................................................................................22
Post-operative pain in mice—its detection and treatment challenges ................................23
Methods of gathering physiological data in undisturbed animals .......................................24
Telemetry .......................................................................................................................24
Objectives and thesis outline ................................................................................................27
Chapter 2: Implantation of radiotelemetry transmitters yielding data on ECG, heart rate, core
body temperature and activity in free-moving laboratory mice ..............................................28
Table of Contents
II
Abstract .............................................................................................................................29
Protocol .............................................................................................................................30
Pre-operative considerations .............................................................................................30
Mice: housing requirements, general condition and health monitoring ...........................30
Hair clipping at one day prior to surgery .........................................................................31
Implantation ......................................................................................................................31
Operating environment, preparation of the telemetric transmitter ...................................31
Anesthesia .....................................................................................................................32
Surgery ..........................................................................................................................32
Post-operative care ...........................................................................................................33
Data acquisition .................................................................................................................34
Representative Results .....................................................................................................35
Tables and Figures: titles and legends ...........................................................................35
Discussion .........................................................................................................................41
Acknowledgments: ............................................................................................................42
Disclosures: ......................................................................................................................43
Chapter 3: Isoflurane and sevoflurane provide equally effective anesthesia in laboratory
mice .....................................................................................................................................44
Abstract .............................................................................................................................45
Introduction .......................................................................................................................45
Table of Contents
III
Materials and Methods ......................................................................................................47
Animals ..........................................................................................................................47
Preliminary transmitter implantation ...............................................................................48
Experimental setting ......................................................................................................48
Determination of minimum alveolar concentration ..........................................................49
Anesthesia experiments .................................................................................................50
Telemetric data acquisition and analysis ........................................................................50
Changes in body weight, and food and water intake ......................................................51
Acid-base balance and blood gas concentration ............................................................52
Statistical analysis ..........................................................................................................52
Results ..............................................................................................................................53
Minimum alveolar concentration .....................................................................................53
Acute effects of anesthesia ............................................................................................53
Post-anesthetic effects ...................................................................................................53
Discussion .........................................................................................................................58
Acknowledgement .............................................................................................................61
Chapter 4: Combining sevoflurane anesthesia with fentanyl-midazolam or s-ketamine in
laboratory mice .....................................................................................................................62
Abstract .............................................................................................................................63
Introduction .......................................................................................................................63
Table of Contents
IV
Materials and Methods ......................................................................................................66
Animals and housing conditions .....................................................................................66
Transmitter implantation .................................................................................................67
Experimental setting ......................................................................................................67
Premedications ..............................................................................................................68
Determination of minimum alveolar concentration ..........................................................68
Anesthesia experiments .................................................................................................69
Telemetric data acquisition and analysis ........................................................................70
Changes in body weight .................................................................................................71
Acid-base balance and blood gas concentration ............................................................71
Statistical analysis ..........................................................................................................72
Results ..............................................................................................................................73
Minimum alveolar concentration .....................................................................................73
Induction of anesthesia ..................................................................................................74
Effects during anesthesia ...............................................................................................75
Effects during the first 3 days after anesthesia ...............................................................76
Discussion .........................................................................................................................79
Acknowledgments ..........................................................................................................84
Chapter 5: Impact of inhalation anesthesia, surgery and analgesic treatment on home cage
behavior in laboratory mice ..................................................................................................85
Table of Contents
V
Abstract .............................................................................................................................86
Introduction .......................................................................................................................87
Materials and Methods ......................................................................................................88
Ethics statement ............................................................................................................88
Animals ..........................................................................................................................88
Experimental groups ......................................................................................................89
Experimental treatments and data recording ..................................................................89
Behavioral analysis ........................................................................................................90
Statistical analysis ..........................................................................................................90
Results ..............................................................................................................................91
Effects of treatment on analyzed behaviors: total 18-hour ..............................................93
Observations ..................................................................................................................93
Effects of treatment on analyzed behaviors: 6-hour observations ..................................93
Discussion .........................................................................................................................96
Conclusion ........................................................................................................................99
Acknowledgements ........................................................................................................99
Chapter 6: General Discussion ........................................................................................... 100
Chapter 7: References ....................................................................................................... 107
Appendix 1 ......................................................................................................................... 128
Acknowledgments .............................................................................................................. 130
List of Tables and Figures
VI
List of Tables and Figures
Chapter 1
Figure 1.1) Structural formula of isoflurane
Figure 1.2) Structural formula of sevoflurane
Figure 1.3) Schematic drawing of DSI ETA F-20 implantable transmitter
Figure 1.4) Schematic drawing of the DSI telemetry acquisition system
Chapter 2
Table 2.1) General condition and health monitoring data sheet (Apendix 1)
Figure 2.1) Schedule for establishing telemetric-transmitter-bearing mice
Figure 2.2) Radiograph/sketch showing location of the implanted telemetry
transmitter.
Figure 2.3) Biopotential curves. Raw printout of one-lead ECG curves from a
conscious mouse and of the same animal under inhalation anesthesia
Figure 2.4) Raw data from long-term measurements in healthy and diseased mice
Figure 2.5) Example of presentation of results from long-term telemetry measurements
after an experiment
Chapter 3
Figure 3.1) Heart rate, core body temperature and respiration rate during 50 min of
anesthesia with isoflurane or sevoflurane
List of Tables and Figures
VII
Figure 3.2) Acid–base balance (pH), partial pressure of carbon dioxide ( pCO2) and
standard bicarbonate (HCO3) in arterial blood at 10, 30 and 50 min of
anesthesia.
Figure 3.3) Post-anesthetic measurements of the impact of isoflurane and sevoflurane on
heart rate, core body temperature and locomotor activity.
Chapter 4
Figure 4.1) Chamber for induction and system for maintenance of sevoflurane inhalation
anesthesia.
Figure 4.2) (A) Mean minimum alveolar concentrations for sevoflurane in adult C57BL/6J
female mice. (B) The mean time required until immobilization after mice were
placed in the sevoflurane-filled induction chamber
Table 4.1) Behaviors of mice during induction of anesthesia with sevoflurane.
Figure 4.3) Mean heart rate, core body temperature, and respiratory rate after
premedication in the home cage, in the induction chamber, and during 50-min
sevoflurane anesthesia
Figure 4.4) Mean acid–base balance (pH), pCO2, and pO2 in arterial blood after 10, 30,
and 50 min of anesthesia
Figure 4.5) Mean postanesthetic measurements of the effects of 3 anesthesia protocols
on heart rate and core body temperature
Chapter 5
Table 5.1) Ethogram of home cage behaviors
Figure 5.1) Scatter plot of discriminant scores assigned to individual mice.
Figure 5.2) Effects of anesthesia and surgery with or without analgesic treatment on
duration of 3 spontaneous home-cage behaviors compared to control values
List of Abbreviations
VIII
List of Abbreviations
°C degrees Centigrade
A anesthesia only group
AALAS American Association for Laboratory Animal Science
ACT activity (as measured by telemetry system)
ANOVA analysis of variance
BT body temperature
bpm beats per minute
C control group
cc cubic centimeter
cm centimeter
CNS central nervous system
d day
DSI Data Sciences International
ECG electrocardiogram
ECLAM European College of Laboratory Animal Medicine
ED50 median effective dose
ESLAV European Society for Laboratory Animal Veterinarians
List of Abbreviations
IX
ETA F-20 model of the telemetry transmitter - ECG, Temperature, Activity
EU European Union
FELASA Federation of Laboratory Animal Science Associations
FiO2 inspiratory oxygen fraction
FMS fentanyl-midazolam-sevoflurane
g gram
GLM univariate general linear model
h hour
HCO3 bicarbonate
HEPA high-efficiency particulate absorption
HFIP hexafluoroisopropanol
HR heart rate
Hz Hertz
i.m. intra muscular
i.v. intra venous
IACUC Institutional Animal Care and Use Committee
KS (S)-Ketamine-Sevoflurane
lx Lux
List of Abbreviations
X
MAC minimum alveolar concentration
mg/kg milligrams per kilogram of body weight
min minute
mL/min milliliters per minute
mm millimeter
mmHg millimeter quicksilver
mmol/L millimol per liter
p.o. oral (per os)
PBS phosphate buffered saline
pCO2 partial pressure of carbon dioxide
pH measure of the acidity or basicity of an aqueous solution
pO2 partial pressure of oxygen
S sevoflurane
s second
S- surgery + anesthesia group
s.c. subcutaneous
S+ surgery + anesthesia + analgesia group
SD standard deviation
List of Abbreviations
XI
UK United Kingdom
Vt tidal volume
wk week
μL microliter
μl/g microliter per gram body weight
Summary
XII
Summary
The mouse is the most commonly used laboratory animal species, with populations of
laboratory mice estimated at more than 60 million worldwide. In the course of bio-medical
research projects, many of these animals will undergo anesthesia and/or procedures that
may elicit discomfort or pain. Apart from the obvious impact on animal wellbeing, such states
could also greatly influence the quality of data obtained. Moreover, mice show strong neuro-
physiological reactions when handled, restrained or tethered during a study. Thus, in order to
study changes in the cardio-vascular system or in behavior, one needs to record data in un-
stressed, freely moving animals in their home-cage environment. To obtain data on
physiological parameters such as heart rate (HR), body temperature (BT) and activity (ACT)
in such animals, telemetry transmitters can be implanted. In this work, we first describe an
improved method of transmitter implantation surgery. By adhering to a few simple rules such
as strict asepsis, optimal anesthesia and intra- and post-operative analgesia and care, a
skilled operator can implant telemetry transmitters to obtain high fidelity data on bio-
potentials, BT and ACT. After a convalescence period of 10 days, such animals can be used
for other experiments without encountering implantation-surgery-related artifacts.
Further, motivated by the fact that, in modern bio-medical research, anesthesia and post-
operative analgesia in laboratory mice is important from both ethical and scientific
perspectives, we set out to test the properties, and the effects on vital parameters, of the two
currently most widely used volatile anesthetics: isoflurane and sevoflurane. Both substances
appear to produce equally safe anesthesia up to 50 minutes. Although having little effect on
cardio-vascular parameters during anesthesia in surgical depth, both compounds induce
marked respiratory depression, resulting in respiratory acidosis and hypercapnia. All animals
displayed a marked increase in HR for a period of 12h after anesthesia with either isoflurane
or sevoflurane.
In order to improve respiration and possibly add a built-in sedative and analgesic component
to inhalation anesthesia, we tested fentanyl-midazolam-sevoflurane (FMS) and S(+)-
ketamine–sevoflurane (KS) balanced anesthesia protocols, against previously established
sevoflurane mono-anesthesia. The FMS protocol proved to be superior to other protocols by
Summary
XIII
inducing less severe respiratory depression without negatively influencing other parameters;
hence, animals experienced less acidosis and hypercapnia. Further, after anesthesia of 50
minutes, the FMS group displayed little to no effect on HR, BT, ACT as compared with other
anesthetic regimes.
In contrast to physiological parameters, behavior can be readily observed and used in
welfare assessments in laboratory routine, thus we were naturally interested in finding a
behavioral correspondence to changes observed by telemetry in our studies but also in
studies involving surgery and post-operative pain. Several spontaneous, home-cage
behaviors were changed as a response to short (15 min) sevoflurane anesthesia and minor
surgery with or without pain treatment. Duration of locomotion, self-grooming and resting
behavior were found to be affected for up to 18 h. Effects gradually increased from animals
anesthetized only, to the operated group that received pain treatment, and were greatest in
the group undergoing surgery without pain treatment. The fact that self-grooming behavior
exhibited the most striking changes after treatment, and that it was the only behavior not
influenced by circadian rhythm, places it center stage in the future search for behavioral
indicators of pain and distress in the laboratory mouse.
This thesis aimed to increase current knowledge on anesthetic regimes and post-operative
pain indicators in laboratory mice. The results indicate clearly that isoflurane and sevoflurane
inhalation anesthesia, although quite safe and superior to injectable protocols, have a
substantial impact on the physiology and behavior of the animal. However, by using a
balanced FMS-anesthesia approach, some of those side-effects could be ameliorated. Thus,
although post-operative pain recognition and therapy is still very challenging, indicators
based on spontaneous home-cage behaviors such as self-grooming might be useful and
should be evaluated further.
Zusammenfassung
XIV
Zusammenfassung
Die Maus ist mit mehr als 60 Millionen gehaltenen Tieren weltweit das am häufigsten
eingesetzte Versuchstier. Im Verlauf von biomedizinischen Versuchen werden viele Tiere
anästhesiert und / oder durchlaufen Behandlungsmethoden, die Unbehagen oder
Schmerzen hervorrufen können. Neben dem offensichtlichen Einfluss dieser Prozesse auf
das Wohlbefinden eines Tieres kann die Qualität der erhobenen Daten dadurch erheblich
beeinflusst werden. Darüber hinaus zeigen Mäuse starke neuro-physiologische Reaktionen
beim Handling im Allgemeinen als auch bei notwendigen Fixationsmethoden während einer
Studie. Um insbesondere Veränderungen im kardiovaskulären System oder im Verhalten
näher erforschen zu können, ist es wichtig, die Daten in ungestressten und sich frei
bewegenden Tieren in deren gewohnter Umgebung zu generieren. Hierzu können
Telemetrie-Sender dienen, die Tieren implantiert werden, um anschliessend Daten über
physiologische Parameter wie Herz-Frequenz (HR), Körperkerntemperatur (BT) und Aktivität
(ACT) zu gewinnen.
In dieser Arbeit wird zunächst eine verbesserte Methode über die chirurgische Implantation
von Transmittern beschrieben. Durch die Einhaltung von einigen einfachen Regeln wie
strikter Asepsis, optimaler Anästhesie sowie adäquater intra-und postoperativer Analgesie
und Betreuung kann ein geübter Operateur Telemetrie-Transmitter selbst implantieren, um
anschliessend aussagekräftige Daten über biologische Potentiale, Körperkerntemperatur und
Aktivität zu gewinnen. Nach zehn Tagen Rekonvaleszenz können die Tiere in anderen
Versuchen eingesetzt werden, ohne dass durch die Implantations-Operation bedingte
Artefakte befürchtet werden müssen.
In der modernen biomedizinischen Forschung spielen die Anästhesie und die post-operative
Analgesie bei Versuchstieren – sowohl aus ethischer als auch aus wissenschaftlicher Sicht -
eine wichtige Rolle. Daher testeten wir in einer weiteren Studie die beiden momentan am
häufigsten verwendeten Inhalationsanästhetika Isofluran und Sevofluran hinsichtlich ihrer
Eigenschaften und ihrer Einflüsse auf Vitalparameter. Beide Substanzen bewirkten eine
gleichwertig sichere Anästhesie bis zu einer Dauer von 50 Minuten. Obwohl während der
Anästhesie bis zum Erreichen der chirurgischen Toleranz kaum Veränderungen der
kardiovaskulären Parameter auftraten, erzeugten beide Anästhetika eine signifikante
respiratorische Depression, die schliesslich zu einer respiratorischen Azidose und
Zusammenfassung
XV
Hyperkapnie führte. Alle Tiere zeigten zudem einen deutlichen Anstieg der Herzfrequenz bis
zu 12 Stunden nach der Anästhesie, sowohl mit Isofluran als auch mit Sevofluran.
Um die Atmung zu stabilisieren und um zusätzlich zur Inhalationsanästhesie eine sedative
als auch analgetische Komponente beizusteuern, wurde eine weitere Studie durchgeführt.
Dabei wurden die beiden Kombinationsanästhesien, Fentanyl-Midazolam-Sevofluran (FMS)
und S(+)-Ketamin-Sevofluran (KS), mit der vorgängig etablierten Monoanästhesie mit
Sevofluran verglichen. Die Anästhesie mit FMS erwies sich hierbei als das beste Anästhesie-
Regime. Die schwere respiratorische Depression wurde abgemildert, ohne dabei andere
Parameter negativ zu beeinflussen; folgend waren Azidose und Hyperkapnie weniger stark
ausgeprägt. Nach einer Anästhesiedauer von 50 Minuten waren bei der FMS-Gruppe, im
Gegensatz zu den anderen Anästhesie-Gruppen, zudem kaum bis gar keine Auswirkungen
auf Herzfrequenz, Körperkerntemperatur und Aktivität festzustellen.
Im Gegensatz zu physiologischen Parametern kann das Verhalten relativ einfach beobachtet
werden und wird in der Laborroutine als Beurteilungskriterium für das Wohlbefinden eines
Tieres herangezogen. Daher waren wir sehr daran interessiert herauszufinden, ob es einen
Zusammenhang zwischen bestimmten Verhaltensweisen und Veränderungen gibt, die wir
durch die Telemetrie beobachten konnten. Der Fokus wurde hier sowohl auf unsere
Telemetrie-Studien als auch auf Studien gelegt, die mit chirurgischen Eingriffen und post-
operativen Schmerzen verbunden sind. Nach einer 15-minütigen Sevofluran-Anästhesie in
Verbindung mit einer kleineren Operation (mit oder ohne Schmerzbehandlung) war die
Dauer von Lokomotion, Putzverhalten und Ruheverhalten im Heimkäfig in Abhängigkeit von
der jeweiligen Behandlung bis zu 18 Stunden nach dem Versuch verändert. Tiere mit
Anästhesie ohne Eingriff zeigten am wenigsten Beeinflussung, gefolgt von den Tieren, die
operiert wurden und eine Schmerzbehandlung erhielten. Die grössten Auswirkungen auf das
Verhalten zeigten sich in der Gruppe von Tieren mit Operation ohne jegliche nachfolgende
Schmerzbehandlung. Das Putzverhalten wurde durch die Versuche am deutlichsten
verändert. Da dieses Verhalten nicht durch den zirkadianen Rhythmus beeinflusst wird,
nimmt es eine besonders wichtige Stellung für die zukünftige Erforschung von
Verhaltensindikatoren für Schmerzen und Leiden bei der Versuchsmaus ein.
Ziel dieser Studie war es, das derzeitige Wissen über Anästhesie und post-operative
Schmerzindikatoren bei der Maus zu vertiefen. Die Ergebnisse weisen klar darauf hin, dass
Zusammenfassung
XVI
Isofluran und Sevofluran, auch wenn beide sehr sicher und den Injektionsanästhesien
vorzuziehen sind, einen wesentlichen Einfluss auf die Physiologie und das Verhalten eines
Tieres haben. Jedoch können durch die Verwendung einer FMS - Kombinationsanästhesie
einige der Nebenwirkungen abgeschwächt werden. Obwohl die Erkennung von post-
operativen Schmerzen und deren Behandlung bei der Maus immer noch eine
Herausforderung ist, können spezifische Verhaltensindikatoren wie das Putzverhalten dabei
sehr hilfreich sein und sollten weiterhin näher untersucht werden.
Chapter 1: General introduction
17
General Introduction
Animal experimentation often plays a major and critical role in modern bio-medical research.
Despite the wide variety of in vitro and in silico tests developed in recent years, there are still
many questions that need to be addressed in animal studies, especially in studies aiming for
a better understanding of human disease, and in the development of new therapeutic
strategies or surgical implants. Due to a whole cluster of factors, including ease of holding
and handling, similarity to human physiology, availability of genetically modified strains and a
magnitude of well-established disease models, the mouse (Mus musculus) is the most
commonly used laboratory animal species. Further, recent improvements in miniaturized
imaging as well as developments in micro-surgical and tissue-engineering techniques have
opened new areas of research in which mice can be used.
In 2008, more than 12 million animals were used in research across the EU, 59.3% of which
were mice [1]. According to the UK Department of Statistics, approximately 40% of all animal
procedures required some kind of anesthesia. Extrapolating these findings to the population
of laboratory mice would mean that more than 2.4 million mice underwent anesthetic
procedures in the EU in 2008 alone. The situation in Switzerland is very similar. In 2012,
more than 250,000 mice were used in experiments in which anesthesia and potentially
painful procedures were applied [2].
Rather than being a goal in itself, anesthesia is often only a tool used while addressing the
scientific question in hand. Most animal experiments requiring anesthesia aim to produce a
carefully defined abnormality or carry out a diagnostic or imaging procedure. Such
procedures assume that there will be no foreign influences on the physiology of the animals
either during or after the procedure [3]. Often, achieving experimental goals will depend
greatly on the ability of the anesthetic protocol to provide stable, relaxed, pain- and
complication-free animals while having little-to-no impact on the experiment itself.
Thus, their decisive role in controlling and diminishing pain and suffering during an
experimental procedure as well as their impact on the subsequent experiment and the data
generated, brings small rodent anesthetic protocols into sharp focus for both the legislative
and scientific communities.
Chapter 1: General introduction
18
Further, research protocols requiring anesthesia for survival of surgery, unavoidably need to
control and ameliorate post-operative pain. Experiments causing mild to moderate pain in
mice, such as skin incision and laparotomy, present difficulties for veterinary staff and
researchers in terms of analgesic treatment. Although there may have been profound effects
on cardio-vascular, endocrine and even immune systems, as prey animals, mice will skillfully
hide any behavioral signs of mild-to-moderate pain, thus rendering post-operative pain
detection and therapy in daily laboratory routine challenging. Finding behavioral changes
indicative of pain might hold the key to the problem of inadequate post-operative analgesia,
and has been the focus of research activity of several groups worldwide.
Although the number of animals used in research continues to decline, the complexity of the
experiments and scientific questions addressed as well as the ethical considerations and
legal requirements are rising rapidly. That being the case, it is of paramount importance to
refine animal experiments. The use of novel and sophisticated general anesthesia methods
to ensure surgical tolerance without influencing the experiment, as well as the search for
precise and ‘user-friendly’ behavioral indicators of post-operative pain enabling individually
tailored analgesic protocols, should contribute greatly to improvement of animal welfare in
bio-medical research.
General anesthesia in small laboratory rodents
Based on the assumption that procedures that elicit pain in humans cause pain in animals,
our modern society confers a fundamental responsibility on individuals using animals in
research, teaching or testing, i.e., to anticipate, and minimize or eliminate any potential pain,
distress or discomfort [4]. By producing a state of unconsciousness, muscle relaxation,
analgesia, diminished motor responses to painful stimulation and suppressed autonomic
responsiveness to noxious stimuli, general anesthesia plays a major role here. According to
the nature and route of drugs administered, either injectable or inhalation anesthesia is used.
In the past, injectable anesthetic protocols were by far the most widely used. However, in the
past decade there has been a clear trend towards the use of inhalation anesthetics in
laboratory routine [5]. The increase in their popularity is due partially to the fact that, in
contrast to injectable drugs, their pharmacokinetic properties enable predictable anesthetic
depth and its rapid adjustment [6]. Moreover, their quick elimination from the body and
Chapter 1: General introduction
19
comparatively low metabolic rate mean that anesthesia-related artifacts in the experiment
occur to a much lesser extent than with injectable drugs.
Inhalation anesthesia in laboratory routine
Inhalation anesthetics are used widely for anesthetic management of animals. They are
administered, and in large part eliminated from the body, via the lungs [6].
The structural diversity of inhaled anesthetics suggests that they do not all interact directly
with a single specific receptor site. Some correlations of the potencies of anesthetics with
their physicochemical properties (lipid solubility) do suggest a common mechanism of
general anesthetic action (Meyer-Overton rule). Current postulates suggest, however, that
general anesthesia results from a constellation of drug effects and that it is unlikely that a
single mechanism of action exists [7].
Inhaled anesthetics may act by altering neuronal activity in several regions of the central
nervous system (CNS). The regions involved include (but are not limited to) the
hypothalamus, thalamus, brain stem and spinal cord as well as the cerebral cortex [7].
Because the brain stem reticular formation plays a role in altering the state of consciousness
and alertness and in regulating motor activity, this structure is often suggested as an
important site of anesthetic action. However, recent findings suggest a crucial role of spinal
cord and other extra-cranial targets in providing certain aspects of surgical anesthesia [8, 9].
Anesthetic vapors can be provided to the animal either by spontaneous respiration via the
nose (nuzzle) mask or by an endo-tracheal tube with assisted positive pressure respiration.
Although necessary in some experimental settings, such as open-chest surgery, insufflation
laparoscopy or intra-pulmonary substance administration [10-12] endo-tracheal intubation
generally tends to be avoided in laboratory rodents. Due to their small size, and the special
materials and skill required for intubation [10, 13] the usual means of maintaining anesthesia
in laboratory mice is by spontaneous respiration via a nose (nuzzle) mask.
The most widely used inhalation anesthesia system for small laboratory rodents in
Switzerland is the modified Bain coaxial breathing (modified Mapleson D) system, which has
an activated coal char filter at the exhaust end. Such machines are usually coupled with
variable bypass, out-of-circuit vaporizers, like the Datex-Ohmeda Tec 5 or Dräger Vapor 19.n
Chapter 1: General introduction
20
used in our facility. Because this system produces very little respiratory resistance, it is well
suited for anesthesia in mice. Furthermore, as it is very easy to operate, it is widely accepted
by the research community.
Isoflurane
Figure 1.1) Structural formula of isoflurane
Isoflurane (2-chloro-2-(difluoromethoxy)-1,1,1-trifluoro-ethane) is a halogenated ether used
for inhalation anaesthesia (Figure 1.1). Its use in human medicine has declined during the
last decade, being replaced mainly with sevoflurane, desflurane and the intravenous
anaesthetic propofol. Nevertheless, isoflurane is generally considered as the most widely
used volatile anesthetic in veterinary medicine today [6]. It has a low blood-gas partition
coefficient, which enables quick induction and recovery. Further, it has a very low
biotransformation rate as only 0.2% of isoflurane appears as metabolites [14], and it is
reported to have a sedative but no-to-weak analgesic effect in humans [15].
Sevoflurane
Figure 1.2) Structural formula of sevoflurane
O
Cl
F2HCH CH CF3
C
F3C
F3C
H OCH2F
Chapter 1: General introduction
21
Sevoflurane (2,2,2-trifluoro-1-[trifluoromethyl]ethyl fluoromethyl ether), also called
fluoromethyl hexafluoroisopropyl ether, is a sweet-smelling, non-flammable, highly
fluorinated methyl isopropyl ether used for induction and maintenance of general anesthesia
(Figure 1.2).
Like isoflurane, sevoflurane has a low blood-gas partition coefficient, but the fact that it is
less irritating to mucosa has made its use very common in human medicine. It is estimated
that 4.9% - 5.6% is biotransformed rapidly in the liver, almost exclusively by the 2E1 isoform
of the cytochrome P450 system. Major metabolites, fluoride and hexafluoroisopropanol
(HFIP)-glucuronide, are eliminated from the body via the kidneys [16, 17]. It is still used only
rarely in veterinary practice.
Minimum Alveolar Concentration (MAC)
Since its description by Merkel and Eger in the early 1960s, the minimum alveolar
concentration (MAC) has become the standard index of anesthetic potency for inhalation
anesthetics [18]. The MAC represents the minimum alveolar concentration of a volatile
anesthetic agent at 1 atmosphere that produces immobility in 50% of individuals subjected to
a noxious stimulus. Thus, MAC corresponds to the median effective dose (ED50), meaning
that, at a given concentration, half of the subjects are anesthetized and the other half have
not yet reached that level.
In animals, the noxious stimulus is usually produced by clamping the tail or by passing an
electrical current through subcutaneous electrodes. The advantage of measuring the alveolar
concentration is that, after a short period of equilibration, this concentration directly
represents the partial pressure of anesthetic in well-perfused areas of the CNS and is
independent of the uptake and distribution of the agent to other tissues [19].
Due to the difficulty of measuring alveolar concentration in small rodents, the inspired
concentration is often used as a proxy [20, 21]. This method is best applied to rapidly
equilibrating (poorly blood-soluble) agents. Only with equilibration can it be assumed that the
partial pressure of the inspired gas equals that at the site of action [19]. However, as it is
strongly desired by clinicians to have all patients (or at least the great majority) that undergo
painful procedures anesthetized, the MAC value needs to be expanded to the anesthetic
Chapter 1: General introduction
22
dose [22]. In this regard, anesthetic dose is commonly defined in terms of MAC multiples and
should correspond to ED95. At least in humans it is 20% - 40% greater than MAC, i.e., 1.2 –
1.4 MAC.
Balanced anesthesia
Anesthesia with a single agent can require doses that produce excessive hemodynamic side-
effects or respiratory depression. For example, the concentrations of isoflurane required to
eliminate movement after skin incision frequently produce hypotension in humans [23, 24],
and induce prominent respiratory depression in mice and man [25, 26].
The concept of balanced anesthesia can trace its roots back to the theory of
anociassociation postulated by Crile in 1910, in which the author described a method for
minimizing pain, shock and post-operative neurosis by combining applied psychology, opioid
pain relief, local and volatile anesthetics [27]. The term ‘balanced anesthesia’ was first
introduced by Lundy in 1926 and was presented as a combined use of different agents
and/or techniques to produce the different components of anesthesia [28].
Currently, balanced anesthesia is defined as the concurrent administration of a mixture of
small amounts of several anesthetic drugs to decrease the adverse effects of each individual
drug [29]. In small animals, it is used mainly to decrease the requirements of inhalant
anesthetics in order to limit the cardiovascular and respiratory depressant effects that each
one induces [30, 31].
Besides the volatile component, balanced anesthesia usually involves co-administration of
an opioid analgesic and/or sedative-hypnotics [29]. As an opioid analgesic component,
fentanyl represents the drug of choice in rodent anesthesia. Compared to buprenorphine (the
opioid most widely used in laboratory animal medicine), although relatively short acting (30-
40 min.), fentanyl does not severely affect respiration when combined with other CNS
depressant agents [3]. In combination with benzodiazepines like midazolam, fentanyl has
been known to produce synergistic effects and induce deep sedation in rodents and humans
[3, 32, 33]. Ketamine is another drug used widely in veterinary medicine. An anesthetic state
is induced by interrupting ascending transmission from the frontal parts of the cortex, rather
than by generalized depression of all brain centers as seen in most general anesthetics [34].
Chapter 1: General introduction
23
The anesthesia produced is a “somnolent state in which the patient does not appear to be
asleep or anesthetized but rather disconnected from his surroundings” [35], thus the term
dissociative anesthesia. The fact that it only marginally affects cardio-vascular and
respiratory systems and at the same time has good analgesic properties, as well as the fact
that it can be applied via virtually all accessible routes (intra muscular, intra peritoneal, intra
venous, intra nasal etc.) renders ketamine a very desirable drug in veterinary medicine [3].
Post-operative pain in mice—its detection and treatment
challenges
Legislation governing the use of animals in biomedical research requires that any
unnecessary pain or distress is avoided or alleviated [36]. Successful implementation of
effective pain management strategies in animals requires accurate assessment of post-
surgical or post-procedural pain [37]. However, pain in animals is difficult to assess, due
mostly to a lack of methods that can validate and objectively measure it. There are no
generally accepted objective criteria for assessing the degree of pain either in humans or in
animals. Species vary widely in their response to pain, and animals of the same species
often show different responses to different types of pain [38].
Post-operative pain in laboratory mice is especially tricky to estimate because its quality and
duration is influenced by a number of factors, e.g., anesthesia, the surgeon’s skills, individual
sensitivity, metabolic disorders, etc. [39]. In addition, as prey animals, mice will generally hide
signs of pain even until high intensity pain or even moribund states are reached, where the
need for therapy is obvious or even already obsolete.
Further, a reason for the underuse of analgesic drugs in the past has probably been fear of
induction of experimental artifacts [5, 40]. Admittedly a number of negative side effects are
associated with nonsteroidal antiinflammatory drugs, including gastrointestinal tract
ulceration, reduction of platelet aggregation, nephrotoxicity, hepatotoxicity, and bone healing
impairment [41]. These drugs have recently been shown to induce apoptosis in cancer cells
as well [42]. Moreover, various opioids have been demonstrated to possess antiinflammatory,
antifibrotic and antitumor abilities as well as to produce emesis, respiratory depression and
bradycardia and to change animal behavior [43-46].
Chapter 1: General introduction
24
This situation often results in the improper use of pain relief measures. Whereas underuse of
painkillers will severely influence animal wellbeing, overuse and general side effects may
have major impacts on the experiment.
Thus, detecting and treating mild-to-moderate post-operative pain in laboratory mice remains
very challenging and relies largely upon defining readily observable behavioral changes
indicative of such states.
Methods of gathering physiological data in undisturbed animals
In the past, in order to obtain data on the physiology or behavior of laboratory animals,
researchers needed to either restrain or anesthetize the animals or rely on inaccurate
measurements on conditioned animals in unnatural environments. It is well known that
anesthesia has a significant effect on heart rate, blood pressure and body temperature, and
that animal behavior is changed by the presence of human observers. Moreover, restraint
influences catecholamines and corticosterone levels and induces stress to such an extent
that it has even been used as a model of hippocampal neuronal damage in rats and
disrupted memory retrieval in mice [47-50]. It is clear that such measurements are inaccurate
and hampered by artifacts [51]. Further, such techniques do not allow the continuous long-
term measurements that are vital for modern physiological and behavioral studies.
Telemetry
Firstly described by Riley in 1970 [52] as a novel method of gathering data of body
temperature in freely moving animals, radio-telemetry found its way into physiology and
cardio-vascular research in the 1990s and has since become an invaluable tool in this field
[51]. However, the size of the radio-telemetry devices developed in the 1970s, and their
inability to be implanted in the body, meant that this technique was of limited use in
laboratory mice [53].
Chapter 1: General introduction
25
Figure 1.3) Schematic drawing of DSI ETA F-20 implantable transmitter
Development of fully implantable radio-telemetry devices has not only enabled chronic
continuous data acquisition from stress-free mice in their ‘home’ environment, but also
minimized the complication rate associated with trans-cutaneous tethering. Since first
described by Kramer in 1993, the implant design and the system itself have changed little
over the years, with the exception of the implants becoming smaller and lighter [54].
Telemetry systems usually consist of an implantable transmitter, a receiver plate(s), data
exchange matrix and a computer to store and subsequently analyze the data (Figure 1.4).
The body of the implantable transmitter used in this work (DSI ETA F-20) consists of a bio-
potential signal amplifier, a temperature probe, radio-frequency electronics, a battery and a
magnetically activated switch that allows the transmitter to be turned on and off while within
the animal's body (Figure 1.3). The transmitter body is hermetically sealed and coated with a
bio-compatible silicone elastomere coating, and occupies a volume of 1.9 cc with a weight of
3.9 grams [55]. A pair of flexible, insulated leads for bio-potential signal acquisition protrude
from the body. The transmitted radio-signal is captured by the receiver plate and digitalized.
After the signal is multiplexed in the data exchange matrix it is transferred to a computer
where it is stored and subsequently analyzed.
Chapter 1: General introduction
26
Figure 1.4) Schematic drawing of the DSI telemetry acquisition system. The mouse is
equipped with the transmitter, which measures ECG, heart rate, core body temperature and
activity data, which are transmitted via radio signals to the receiver plate. The receiver plate
is located nearby (usually underneath) the animal’s cage. A data exchange matrix provides
power to the receivers on the one hand, and on the other multiplexes a number of signals
into one that can be stored on the computer via the Dataquest PCI card. (Illustration adapted
from Weiergräber 2005) [56]
Chapter 1: Objectives and thesis outline
27
Objectives and thesis outline
The goal of this thesis was to detect, further develop and describe inhalation and balanced
anesthesia protocols for use in laboratory mice. Further, we were interested in the impact of
such protocols on physiological and behavioral parameters in the animal, thus demonstrating
their advantages and adverse effects and identifying influences on research read-out. For
this purpose, undisturbed, freely moving mice were monitored during and following
anesthesia in their home cages, thus closely mimicking the situation in laboratory routine.
- Chapter 2
o This chapter describes optimization of implantation procedures for the
telemetry transmitters yielding data on ECG, heart rate, core body
temperature and activity in freely moving mice
- Chapter 3
o The focus of this chapter was to test the differences between two inhalation
anesthetics: isoflurane and the more modern sevoflurane. We were mainly
interested in determining minimum alveolar concentration (MAC) values and
looking into safety (regarding physiological parameters) during and following a
50-min long period of anesthesia.
- Chapter 4
o This study aimed to develop new balanced anesthesia protocols for small
laboratory rodents and to compare these with standard inhalation anesthesia.
- Chapter 5
o To discriminate between the impact of anesthesia, post-operative pain and
analgesic treatment, behavioral effects of a short inhalation anesthesia and
minor surgery with or without pain treatment were addressed in this chapter.
Chapter 2: Implantation of Radiotelemetry Transmitters
28
Implantation of radiotelemetry transmitters yielding
data on ECG, heart rate, core body temperature and
activity in free-moving laboratory mice
Based on Cesarovic N, Jirkof P, Rettich A, Arras M.
Published Nov. 2011 in Journal of visualized experiments : JoVE
The video component of this article can be found at http://www.jove.com/video/3260/
Chapter 2: Implantation of Radiotelemetry Transmitters
29
Abstract
The laboratory mouse is the animal species of choice for most biomedical research, in both
the academic sphere and the pharmaceutical industry. Mice are a manageable size and
relatively easy to house. These factors, together with the availability of a wealth of
spontaneous and experimentally induced mutants, make laboratory mice ideally suited to a
wide variety of research areas.
In cardiovascular, pharmacological and toxicological research, accurate measurement of
parameters relating to the circulatory system of laboratory animals is often required.
Determination of heart rate, heart rate variability, and duration of PQ and QT intervals are
based on electrocardiogram (ECG) recordings. However, obtaining reliable ECG curves as
well as physiological data such as core body temperature in mice can be difficult using
conventional measurement techniques, which require connecting sensors and lead wires to a
restrained, tethered, or even anaesthetized animal. Data obtained in this fashion must be
interpreted with caution, as it is well known that restraining and anesthesia can have a major
artifactual influence on physiological parameters [26, 57].
Radiotelemetry enables data to be collected from conscious and untethered animals.
Measurements can be conducted even in freely moving animals, and without requiring the
investigator to be in the proximity of the animal. Thus, known sources of artefacts are
avoided, and accurate and reliable measurements are assured. This methodology also
reduces inter-animal variability, thus reducing the number of animals used, rendering this
technology the most humane method of monitoring physiological parameters in laboratory
animals [54, 58]. Constant advancements in data acquisition technology and implant
miniaturization mean that it is now possible to record physiological parameters and locomotor
activity continuously and in real-time over longer periods such as hours, days or even weeks
[39, 58].
Here, we describe a surgical technique for implantation of a commercially available telemetry
transmitter used for continuous measurements of core body temperature, locomotor activity
and biopotential (i.e. one-lead ECG), from which heart rate, heart rate variability, and PQ and
QT intervals can be established in free-roaming, untethered mice. We also present pre-
Chapter 2: Implantation of Radiotelemetry Transmitters
30
operative procedures and protocols for post-operative intensive care and pain treatment that
improve recovery, well-being and survival rates in implanted mice [39, 59].
Protocol
The animal experiment was approved by the Cantonal Veterinary Office (Zurich,
Switzerland). Housing and experimental procedures were in accordance with Swiss Animal
Protection law and conform to the European Directive on the Protection of Animals Used for
Scientific Purposes (DIRECTIVE 2010/63/EU OF THE EUROPEAN PARLIAMENT AND OF
THE COUNCIL of 22 September 2010).
Pre-operative considerations
Mice: housing requirements, general condition and health monitoring
It is recommended that mice delivered from vendors or transferred from external rodent
colonies should arrive at the housing facility at least two weeks prior to surgery. This period
should allow the animals to adapt to the new environment and facility-specific housing
conditions. Mice, as social living animals, should be housed in compatible groups during this
adaptation period. For monitoring an individual’s level of food and water consumption, each
mouse is housed singly from 3 days before surgery until 10 days after surgical transmitter
implantation. The time line for establishing telemetric-transmitter-bearing mice is shown in
Figure 2.1. It is crucial that the animals come to surgery in good health and condition.
Therefore, before surgery, animals should be monitored once per day for 2-3 days
concerning general condition (appearance, posture, spontaneous behavior) as well as for
body weight, food and water consumption. These data are documented on a medical record
(general condition and health monitoring data sheet, appendix 1) to establish individual
baseline levels of general condition and overall health and wellbeing. Any animals showing
symptoms of disease or impaired general condition before surgery should be excluded from
the experiment.
Chapter 2: Implantation of Radiotelemetry Transmitters
31
Hair clipping at one day prior to surgery
The day prior to implantation, in order to shave the animals for surgery, mice are
anesthetized briefly in a small (8x8x8cm) Perspex chamber using sevoflurane (8%) or
isoflurane (5%) in pure oxygen (600 mL/min). After loss of the righting reflex, the mouse is
taken out of the chamber and the anterior neck and abdominal hair is clipped with the animal
lying in dorsal recumbence; anesthesia is maintained for approximately 5 minutes with a
nose mask with sevoflurane 3-4% or isoflurane 1.5-3% in pure oxygen at a flow rate of 600
mL/min. After clipping the hair, the animals are allowed to awaken and are then brought back
to their home cage.
Implantation
Operating environment, preparation of the telemetric transmitter
On the day of implantation, all procedures regarding transmitter preparation and surgery are
carried out on a work bench with a laminar flow hood equipped with a surgical microscope.
Aseptic conditions are assured by the use of autoclaved instruments and sterilized materials
and by disinfecting the work bench [60]. Prior to implantation, the telemetric transmitters
(ETA-F10, Data Sciences International, St. Paul, MN, USA) are first prepared. After removing
from their sterile package, the leads of the transmitter are shortened to a length appropriate
for the size of the mouse to be implanted. In the majority of adult outbred or inbred mice, the
red electrode may be shortened to approximately 42 mm and the white/colourless electrode
to a length of approximately 55 mm. Insulation tubing is removed from the distal (sensory)
part of the leads: approximately 20 mm of tubing is removed from the red electrode,
approximately 10 mm of tubing is removed from the white/colourless electrode. The distal
part of each electrode (which is now without tubing) is formed into a loop by fixing the end
with thin silk sutures (PERMA-Handseide, 6-0, Ethicon, Norderstedt, Germany). After
preparing the electrodes, the transmitter is placed in warm sterile saline ready to be
implanted when the animal is anesthetized and surgically prepared.
Chapter 2: Implantation of Radiotelemetry Transmitters
32
Anesthesia
At 5–10 minutes before induction of inhalation anesthesia, a mixture of midazolam (4 mg/kg)
and fentanyl (0.04 mg/kg) are administered subcutaneously as premedication, thus providing
sedation and pre-emptive analgesia. General inhalation anesthesia is induced by placing the
animal in the induction chamber and introducing the volatile anesthetic agent (sevoflurane
8% or isoflurane 5% in pure oxygen 600 ml/min). When the animal shows loss of the righting
reflex it is transferred to the work bench under the laminar flow hood, and placed in dorsal
recumbence on a specially designed metal plate fitted with a nose mask and tubing from the
anesthesia apparatus. Anesthesia is maintained by spontaneous breathing (sevoflurane 3-
4% or isoflurane 1.5-3% in pure oxygen at a flow rate of 600 mL/min). During anesthesia, the
animal’s eyes are protected with ointment (Vitamin A, Baush & Lomb, Steinhausen,
Switzerland). While lying on the metal plate the animal is warmed by the water-bath heated
surface (39°C +/-1) of the work bench.
Surgery
The skin of the anterior neck and abdominal region is disinfected with 70% ethanol. A 1- to
1.5-cm-long incision in the skin is made from the lower thorax along the midline to the
abdomen. The negative (white/colorless) lead is tunneled subcutaneously from the thorax to
the neck, where a small incision (≤0.5 cm) is made in the longitudinal direction. The skin and
underlying tissues are prepared to make space for the fixation of the wire loop of the
electrode. The wire loop is fixed between the muscles located to the right of the trachea,
using two thin silk sutures (PERMA-Handseide, 6-0, Ethicon, Norderstedt, Germany). The
wound in the neck is then closed with absorbable sutures (VICRYL 6-0, Ethicon, Norderstedt,
Germany) in layers. The abdominal wall is then opened at the linea alba and the body of the
telemetric transmitter is placed into the abdominal cavity of the mouse. The wire loop of the
positive (red) electrode is sutured to the xiphoid process with silk sutures in such a way that it
lies between the liver and the diaphragm in the left upper abdominal region (Figure 2.2).
Then, the muscle layers of the abdominal region are closed with absorbable sutures
(VICRYL 6-0, Ethicon, Norderstedt, Germany). Before finally closing the abdominal wall, a
mixture of Sulfadoxin and Trimethoprim [(30 mg/kg and 6 mg/kg, respectively; dissolved in 1
mL of saline (0.9%) and at approximately body temperature (38-39°C)] is injected into the
abdominal cavity for the purposes of anti-infective prophylaxis and to support fluid
Chapter 2: Implantation of Radiotelemetry Transmitters
33
homeostasis. Finally, the skin of the abdominal region is restored with staples (Precise®, 3 M
Health Care, St. Paul, MN, USA).
Post-operative care
After completion of surgery and anesthesia, 0.1 mg/kg of buprenorphine (Temgesic®, Essex
Chemie AG, Lucerne, Switzerland) and 5 mg/kg of meloxicam (Metacam®, Boehringer
Ingelheim, Basel, Switzerland) is administered subcutaneously for pain treatment, and the
animals are left on the warm (39°C +/-1) surface of the work bench to recover for
approximately 2h. 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, is applied subcutaneously twice daily for 4 days. For
further recovery support, it is worthwhile providing the animals with an additional drinking
bottle containing 15% glucose solution. During the recovery period of 4-10 days, it is
recommended that the animals are kept warm. Therefore, in our case, the mice are housed
in a warming cabinet (30°C +/- 1). Monitoring of general condition and body weight, as well
as food and water consumption, is performed once daily according to the general condition
and health monitoring data sheet (Appendix 1) for 10 days post-operatively. Humane
endpoints, i.e. the sacrifice of an animal to avoid unnecessary suffering and pain if
progression of recovery is unsatisfactory, are realized under the following conditions:
i.) If in poor general condition, i.e. the animal is substantially apathetic (no movement after
being touched/pushed) and its body surface feels cold despite warming, the animal should
be euthanatized immediately.
ii.) If, on day 4 after transmitter implantation, the animal shows clear signs of apathy, is
extremely aggressive or does not show any food intake, it should be euthanatized
immediately.
iii.) On day 8 after transmitter implantation, the animal has to display a clear increase in body
weight in comparison to the preceding post-operative days. Moreover, it has to consume at
least 80% of the pre-operative daily food intake. If one of these conditions is not met, the
animal should be euthanatized immediately.
Chapter 2: Implantation of Radiotelemetry Transmitters
34
At 10 days after implantation, the animal is transferred back to the animal room under
standard housing conditions. Mice should be housed in compatible groups to allow social
interaction and to prevent the adverse effects of long-term individual housing, which can
have substantial impacts on the read-out of subsequent experiments [61, 62]. Mice should
have a period of at least 4 weeks convalescence after transmitter implantation before the first
experiment is conducted and data acquisition begins.
Data acquisition
Data collection is initiated by touching the animal with a magnet, whereupon the transmitter
is switched on. Dataquest A.R.T. Software (Data Sciences International, St. Paul, MN, USA)
coordinates the detection, collection, analysis and graphical presentation (in the form of wave
forms) of signals from one or more animals. The Acquisition program collects data signals
sent to the computer from the converters and receivers via a Data Exchange Matrix (Data
Sciences International). This program can either collect data for a specific length of time at
regular intervals or sample continuously and save the data on the computer’s hard drive. As
the range and the quality of the emitted signal depends strongly on the material composition
of the cage and surrounding equipment (e.g. metal vs. plastic), it is suggested that the
receiver plate is placed as close to the animal as possible, e.g. under the animals’ cage or
above the experimental area, e.g. laboratory bench or treadmill. It is recommended that the
correct configuration of the recording and data transmission system be checked by making a
short examination of real-time measurements in continuous sampling mode. After the data
have been gathered and stored, they can be plotted, listed and analyzed for a variety of
different parameters using the Analysis program. Details of the configuration of the recording
system (e.g. defining the sampling modus), and analysis software (e.g. for heart rate
variability parameters, PQ interval and QT interval established from biopotential/ECG curves)
can be found in the manufacturer’s manuals. Valuable hints for biometric planning and
statistical methods useful for telemetric data acquisition and interpretation are published
elsewhere [58].
Chapter 2: Implantation of Radiotelemetry Transmitters
35
Representative Results
An overall scheme of the described procedure is shown in Figure 1. The position of the
implanted transmitter, including the location of the electrodes for obtaining biopotentials from
the heart (one-lead ECG) is shown in Figure 2.2. Examples of raw data from short term
biopotential curves (one-lead ECG), and long-term heart rate, core body temperature and
locomotor activity recordings of individual mice are given in Figure 2.3 and Figure 2.4,
respectively. Figure 2.5 gives an example of published data from long-term measurements in
groups of mice after an experiment. Several other parameters can be established from the
biopotential curves. Examples for presentation of heart rate variability parameters [63], QT
interval and PQ interval [64, 65] are published elsewhere.
Tables and Figures: titles and legends
Table 2.1) General condition and health monitoring data sheet (Appendix 1)
This template facilitates monitoring of an individual mouse’s general condition and health.
Baseline examination of an animal’s appearance, posture, and spontaneous behavior, as
well as determination of body weight, and food and water consumption must be established
before implantation surgery once per day for 3 days. Comparison of baseline determinations
with those obtained daily for 10 days after surgery serve to assess the progression of post-
operative recovery. In addition, post-operative care and pain treatment are well documented
in the form of a medical record. Instructions on humane endpoints are given in order to
facilitate decisions on whether a mouse should be sacrificed to prevent unnecessary pain
and suffering if the animal does not meet the criteria for fast recovery after implantation.
Chapter 2: Implantation of Radiotelemetry Transmitters
36
Figure 2.1) Schedule for establishing telemetric-transmitter-bearing mice
Chronological order of procedures relating to the implantation of a transmitter showing the
time points at which a mouse can be used for experiments and data acquisition.
Figure 2.2) Radiograph/sketch showing location of the implanted telemetry transmitter. The
body of the transmitter is positioned in the abdominal cavity. The positive lead is formed into
a wire loop and fixed to the xiphoid process with sutures. The negative lead is tunneled
subcutaneously from the thorax to the neck and fixed as a wire loop between the muscles
directly next to the trachea. The radiograph is taken from the authors' previous publication in
Laboratory Animals [62].
day -3
implantation
day 10
convalescence (3-4 weeks)
daily monitoring of general condition and health
baseline postoperative recovery phase data acquisition phase
Chapter 2: Implantation of Radiotelemetry Transmitters
37
Figure 2.3) Biopotential curves. Raw printout of one-lead ECG curves from a conscious
mouse and of the same animal under inhalation anesthesia with sevoflurane. Heart rate is
calculated automatically by the telemetry system. The 3-second sequence recorded under
anesthesia indicates a heart rate of 440 bpm. The curve recorded in the conscious mouse
shows a heart rate of 660 bpm, which falls within the expected range for heart rate during
moderate physical activities such as grooming or eating. From biopotential/one-lead ECG
curves, heart rate variability parameters, interbeat interval, and PQ and QT intervals can be
established with use of the manufacturer’s software.
conscious
anesthetized
Chapter 2: Implantation of Radiotelemetry Transmitters
38
Figure 2.4) Raw data from long-term measurements in healthy and diseased mice. Heart
rate (bpm), core body temperature (°C) and locomotor activity (counts) are measured while
mice are housed individually in their home cage without any disturbance from man or
experimental procedures. Heart rate is recorded for 30 seconds every 5 minutes (sampling
frequency 1000 Hz). Core body temperature is sampled for 10 seconds every 5 minutes.
Locomotor activity is recorded continuously and stored at 5 minute intervals. Five-minute
data points are traced for 6.5 days. The telemetric measurements are recorded from three
mice with differing bodily conditions. The healthy mouse shows a clear circadian rhythm with
normal increases in physiological values and locomotor activity behavior during the dark
(night) phase. In contrast, after major surgery, heart rate is increased, particularly in the
daylight phase, and locomotor activity is depressed. The third mouse suffered from chronic
tumor disease—its circadian rhythm of heart rate and core body temperature appears
flattened, and locomotor activity is diminished. Representative data of heart rate
measurements (normal values and after major surgery) are taken from the authors' previous
publication in Altex [39].
6d tumor disease
he
art
ra
te [
bp
m]
900
800
700
600
500
400
300
200
chronic tumor disease
6 d, tumor disease
bo
dy t
em
pe
ratu
re [
°C]
39
38
37
36
35
34
6d, tumor disease
acti
vity
12
10
8
6
4
2
0
1 day
major surgery
6 days after transmitter implantation
he
art
ra
te [
bp
m]
900
800
700
600
500
400
300
200
6 d after transmitter implantation
bo
dy t
em
pe
ratu
re [
°C]
39.0
38.0
37.0
36.0
35.0
34.0
6 d after transmitter implantation
acti
vity
12
10
8
6
4
2
0
1 day
normal
1day
6d, control
he
art
ra
te [
bp
m]
900
800
700
600
500
400
300
200
6d, control
bo
dy t
em
pe
ratu
re [
°C]
39
38
37
36
35
34
6d, normal
acti
vity
12
10
8
6
4
2
0
1 day
Heart
Rate
[bpm]
Locomotor
activity
[counts]
Body
temp.
[°C]
Chapter 2: Implantation of Radiotelemetry Transmitters
39
-1
-0.8
-0.6
-0.4
-0.2
0
0.2
0.4
0.6
0.8
1
-60
-40
-20
0
20
40
60
80
100
120
[°C
][b
eats
per
min
ute
]
heart rate
[% c
ou
nts
]
core body temperature
day 2day 1 day 3 day 4
Isoflurane
Sevoflurane
-100
-50
0
50
100
150
200 locomotor activity
*
*
*
*
Chapter 2: Implantation of Radiotelemetry Transmitters
40
Figure 2.5) Example of presentation of results from long-term telemetry measurements after
an experiment. The figure 2.5 is taken from the authors' previous publication in Laboratory
Animals [26]. As an exemplary experiment, a 50-minute isoflurane or sevoflurane anesthesia
was performed. The long-term impact of the anesthetics on heart rate, core body
temperature and locomotor activity after the animals were awake was compared. Using 16
transmitter-implanted mice, telemetric data were recorded in eight mice per anesthetic while
the animals were single-housed and allowed to roam freely in their home cages. For analysis
of long-term postanesthetic effects, we took into account that values vary greatly during a 24-
h cycle since mice are active mainly at night. Therefore, the means of the telemetric values
for each animal were calculated separately for the night (12 h dark) and day (12 h light)
phases. An individual’s normal values were established by calculating means from the three
days prior to anesthesia. For each day after anesthesia, the mean of the dark and light phase
was compared with the individual’s normal values, resulting in delta values. Thus, delta
values represent deviation from normal values (established prior to anesthesia) at the
corresponding 12 h day and night time. Columns represent the mean from eight mice; bars
indicate standard deviation. Asterisks indicate significance at P ≤ 0.05 (One-way analysis of
variance for comparison of group means at each of the four days after anesthesia with
normal values).
Chapter 2: Implantation of Radiotelemetry Transmitters
41
Discussion
Radiotelemetry is a powerful alternative to conventional methods of measurement of
physiological parameters in biomedical research. High-quality telemetry systems consisting
of implantable transmitters, receivers and data acquisition and analysis hardware and
software are now commercially available, even for animals as small as mice. Telemetry
represents the only technique currently available for data collection from unrestrained, freely
moving mice. By using this method, it is now possible to gather data continuously and/or for
longer periods of time from animals residing in their own familiar environment, thus
minimizing the stress to the animals and consequent experimental artifacts. The form and
position of the leads has been optimized in order to obtain signals even during fast
movements (e.g. struggling, running, fighting) or in an upright posture [62]. Thus, accurate
measurements can be obtained during experiments, e.g. during anesthesia, stress induction,
while running on a treadmill, during behavioral experiments, during infection experiments,
and many other experimental situations.
However, in order to obtain reliable, reproducible and artifact-free data, it is crucial to exclude
environmental influences, and we draw particular attention to the importance of standardized
conditions. It is recommended that the room is isolated from electronic and acoustic noise,
including ultrasonic sound, to which mice are particularly sensitive. In addition, no
disturbances, such as visitors or unrelated experimental procedures, should be allowed when
conducting measurements. To avoid interfering influences (particularly in case of home cage
measurements), all necessary husbandry procedures should be completed in the room
before starting each measurement. In addition, the housing of mice — particularly if males
are used — in groups or individually can have an impact on the measurements and must be
considered when planning experiments [62]. Also, the mice must be healthy and free of
murine pathogens, since latent or manifest infections, as well as diseases or any other health
impairments, can have considerable influence on physiological
parameters and activity behavior. Accordingly, mice should recover fully after implantation
and be given sufficient time to adapt to bearing the transmitter before starting any
experiments.
Chapter 2: Implantation of Radiotelemetry Transmitters
42
Data collection by radiotelemetry in mice requires preliminary surgical implantation of the
telemetry transmitter. This should be performed only by trained personnel with surgical skills
in order to minimize tissue trauma and subsequent pain and distress. For experimenters
holding basic or even advanced (micro-) surgical skills, it is recommended to perform the first
trials in fresh mouse cadavers using training implants (i.e., dummies, provided by the
manufacturer) to establish the procedures and become familiar with the specifics of this kind
of surgery. After such training, most experimenters would be capable to implant this type of
transmitters with success and would reach a useful proficiency after a few implantations.
Aseptic conditions should be maintained during surgery to keep the microbiological burden
and the risk of infections low. However, complete sterility cannot be provided because of
some specific, sterility conflicting conditions in mice (e.g., cooling effect of extensive hair
clipping and disinfection, impracticality of bandages to protect the wounds). Thus, anti-
infective prophylaxis is administered during the implantation. Well-tailored analgesic
treatment and a clearly defined monitoring plan as well as adequate post-operative care play
a crucial role in the satisfactory outcome of the experiment.
Overall, the surgical implantation of a telemetric transmitter in mice will be stressful for the
animal. In particular, if genetic modification in specific mouse lines influences the phenotype
and impairs the animals’ bodily condition, complications in the peri-operative time frame and
increased death rates after implantation might be a risk. To avoid unnecessary suffering,
individuals exhibiting unsatisfactory recovery or prolonged convalescence should be
released from the experiment and sacrificed before reaching a moribund stage. For this
purpose, a data sheet (Apendix 1: general condition and health monitoring data sheet)
facilitating the systematic monitoring of critical symptoms and providing advice on humane
endpoints has been established. Thus, recovery is documented in the style of a medical
record or a laboratory journal, which makes the conducting of this methodology (i.e.
implantation procedure and post-operative recovery) transparent to the relevant authorities
and animal welfare bodies responsible for animal experimentation (e.g., IACUC).
Acknowledgments:
The authors would like to thank Charles River Germany for providing CD-1 mice. We also
thank Robin Schneider and the staff of the central biological laboratory for support in housing
Chapter 2: Implantation of Radiotelemetry Transmitters
43
mice. We kindly thank Flora Nicholls for excellent technical assistance and Professor Kurt
Burki for generously providing research facilities and resources.
Disclosures:
We have nothing to disclose.
Chapter 3: Inhalation Anesthesia
44
Isoflurane and sevoflurane provide equally effective
anesthesia in laboratory mice
Based on Cesarovic N, Nicholls F, Rettich A, Kronen P, Hässig M, Jirkof P, Arras M.
Published Oct 2010 in Laboratory animals
Chapter 3: Inhalation Anesthesia
45
Abstract
Isoflurane is currently the most common volatile anesthetic used in laboratory mice, whereas
in human medicine the more modern sevoflurane is often used for inhalation anesthesia.
This study aimed to characterize and compare the clinical properties of both anesthetics for
inhalation anesthesia in mice. In an approach mirroring routine laboratory conditions
(spontaneous breathing, gas supply via nose mask, preventing hypothermia by a warming
mat) a 50-minute anesthesia was performed. Anesthetics were administered in oxygen as
carrier gas at standardized dosages of 1.5 minimum alveolar concentrations, which was
2.8% for isoflurane and 4.9% for sevoflurane. Both induction and recovery from anesthesia
proceeded quickly, within 1-2 minutes. During anesthesia, all reflex testing was negative and
no serious impairment of vital functions was found; all animals survived. The most prominent
side effect during anesthesia was respiratory depression with hypercapnia, acidosis, and a
marked decrease in respiration rate. Under anesthesia, heart rate and core body
temperature remained within the normal range, but were significantly increased for 12 hours
after anesthesia. Locomotor activity, daily food and water consumption and body weight
progression showed no abnormalities after anesthesia. No significant difference was found
between the two anesthetics. In conclusion, isoflurane and sevoflurane provided an equally
reliable anesthesia in laboratory mice.
Introduction
Rodents are usually anaesthetized by injection of hypnotic, analgesic and muscle relaxant
liquid agents [40]. Since continuous intravenous, target-controlled infusion, or so-called total
intravenous anesthesia, with short acting drugs such as propofol (e.g. [66]), is hard to master
in mice, the intraperitoneal or subcutaneous application route is normally chosen in this
species [67-69]. Although it would seem easy and practical to induce general anesthesia with
an injection of a single (e.g. pentobarbital) or mixed (e.g. ketamine/xylazine,
medetomidine/midazolam/fentanyl) long-acting drug(s), this type of anesthesia is hard to
control. Once the initial dose has been administered, the duration and depth of anesthesia
cannot be adjusted to the specifics of the mouse (strain, age, gender, individual variation,
Chapter 3: Inhalation Anesthesia
46
etc.) or the surrounding conditions (time of day, housing conditions, etc.), all of which
influence the animals’ response to the anesthetic [70-74]. Thus, despite prior dosage testing,
managing injection anesthesia often remains difficult, i.e. on the one hand anesthesia is
shallow in some individuals, and on the other, the death rate can be unexpectedly high [67,
68, 75].
Such failures are rarely encountered with inhalation anesthesia. Modern, commercially
available volatile anesthetics such as isoflurane, sevoflurane, desflurane and others are
vaporized in dedicated vaporizers, added to a carrier gas and supplied to the animal via the
respiratory tract. Due to their low blood:gas partition coefficients, these compounds provide
rapid induction of anesthesia, are short-acting, and are removed from the body in a short
time, mostly by respiration [76]. The dosage can be adapted easily and can be titrated to
effect for an individual animal. Thus, provided that the animals’ vital functions and depth of
anesthesia are monitored, cases of death are unusual because the course of anesthesia can
be easily controlled. Thus, recently developed volatile anesthetics are used increasingly in
laboratory rodents [40, 77] especially since ready-to-use inhalation anesthesia devices
tailored for small rodents are commercially available. The most up-to-date anesthesia
equipment [78-80] normally includes active scavenging systems to prevent the release of
waste gas, which is mandatory for protecting personnel and which has been a problem in the
past [81-84].
The most common and well-known volatile anesthetic in laboratory rodents is isoflurane [40].
Sevoflurane, a more modern inhalation anesthetic, is used in human medicine [85], but is
uncommon in veterinary medicine due to its higher cost. To date, the clinical impacts of
isoflurane and sevoflurane have been described mostly in man or in animal species other
than mice. This led us to investigate the possible advantageous properties and drawbacks of
isoflurane and sevoflurane anesthesia in laboratory mice. These two substances were
compared in a practical setting for their effects during and after anesthesia from a clinical
viewpoint, with the aim of determining their impact on animal physiology and general post-
anesthetic condition.
Chapter 3: Inhalation Anesthesia
47
Materials and Methods
Animals
Sixty-four, 6-week-old female C57BL/6J mice were obtained from our in-house breeding
colony. The mice were free of all viral, bacterial, and parasitic pathogens listed in FELASA
recommendations [86]. Health status was monitored by a sentinel program throughout the
experiments.
Animals were kept in type 3 open-top plastic cages (425 mm x 266 mm x 150 mm, floor area
820 cm2) with autoclaved aspen bedding (80–90 g/cage) (LTE E-001 Abedd, Indulab, Gams,
Switzerland). Autoclaved hay (8–12 g/cage) and 2 Nestlets™ (each 5 x 5 cm), consisting of
cotton fibers (Indulab, Gams, Switzerland) were provided as nesting materials. A standard
cardboard house (Ketchum Manufacturing, Brockville, Canada) was provided. Animals were
fed a pelleted mouse diet (Kliba No. 3431, Provimi Kliba, Kaiseraugst, Switzerland) ad libitum
and had unrestricted access to sterilized drinking water. The light/dark cycle in the room
consisted of 12/12 h with artificial light (approximately 40 Lux in the cage) from 03:00 h to
15:00 h. The temperature was 21±1°C, with a relative humidity of 50±5%, with 15 complete
changes of filtered air per hour (HEPA H 14 filter); the air pressure was controlled at 50 Pa.
Mice were housed in groups, except during the 4 days before and 4 days after anesthesia,
when they were housed individually. The first day of single housing served for adaptation to
the change in housing conditions; from the second day onwards, the individuals’ normal
values for heart rate, core body temperature, locomotor activity, body weight, food and water
consumption were recorded. To avoid interfering influences, all necessary husbandry and
management procedures were completed in the room before starting single housing of mice,
and disturbances (e.g. visitors or unrelated experimental procedures) were not allowed. The
animal room was insulated to prevent electronic noise.
The study was approved by the Cantonal Veterinary Office (Zurich, Switzerland) under the
license number 111/2007. Housing and experimental procedures were in accordance with
Swiss animal protection law and conform to the European Convention for the protection of
Chapter 3: Inhalation Anesthesia
48
vertebrate animals used for experimental and other scientific purposes (Council of Europe
nr.123 Strasbourg 1985).
Preliminary transmitter implantation
Prior to the experiments, at age 10 weeks, 16 mice were instrumented with telemetric
transmitters. TA10ETA-F20 transmitters (Data Sciences International, St. Paul, MN, USA) --
which measure heart rate, core body temperature and locomotor activity in freely moving
mice -- were implanted as previously described in detail [63]. Briefly, under anesthesia 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
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. Post-operatively, buprenorphine (Temgesic™, Reckitt and Colman Products Ltd.,
Hull, England) was given at a dose of 0.1 mg/kg body weight, injected subcutaneously twice
per day for 4 days [59]. After transmitter implantation, mice had a period of 6 weeks
convalescence before the first experiment.
Experimental setting
All experiments were conducted when the mice were aged 16--36 weeks, with body weights
ranging from 25 to 30 g. All experiments and weighing procedures were carried out between
15:00 h and 18:00 h. Anesthesia was performed in a separated operating area within the
animal room.
Anesthesia was provided by a commercially available rodent inhalation anesthesia apparatus
(Provet, Lyssach, Switzerland), which was equipped with interchangeable vaporizers for
isoflurane (Ohmeda Isotec 5, Abbott, Baar, Switzerland) and sevoflurane (Ohmeda Sevotec
5, Abbott, Baar, Switzerland). As carrier gas, 100% oxygen was used at a flow rate of 400
mL/minute. The anesthetic gas was introduced into the nose mask (inner diameter 1.2 cm)
through a thin tube (outer diameter 0.4 cm, inner diameter 0.3 cm). The opening of the thin
tube was at a distance of exactly 0.5 cm from the latex membrane, which had a hole in the
Chapter 3: Inhalation Anesthesia
49
center that fit around the nose of the mouse. The nose of the mouse was placed 2 mm in
front of the opening of the inner tube. The nose mask merged into a thick outer tube
(surrounding the thin inner tube), which allowed waste anesthetic gas to be eliminated from
the nose mask by a pump-driven filter system (flow rate 400 mL/minute). The same principle
was utilized for the induction chamber (inflow and outflow 400 mL/minute). The
concentrations of anesthetic gases in the nose mask (at 2 mm in front of the opening of the
inner tube) and in the induction chamber were measured at the beginning of anesthesia and
then every 5 minutes using a side-stream monitoring device employing infrared technology
(Datex-Ohmeda AS/3, Anandic Medical, Deissenhofen, Switzerland). The device was
calibrated just before use using the proprietary standard reference gas supplied by the
manufacturer.
Determination of minimum alveolar concentration
Forty-eight non-transmitter-implanted mice underwent anesthesia three to four times in order
to standardize anesthesia by establishing minimum alveolar concentration. Care was taken
that animals had a break of at least two weeks between tests.
Minimum alveolar concentration was determined with a protocol modified from published
methods [87-89]. Briefly, after inducing anesthesia in the chamber for two minutes at
maximum concentration of anesthetic gases (5% isoflurane, 8% sevoflurane), the mouse
was taken out of the chamber and placed in dorsal recumbence on a warmed mat (see
below). Anesthetic gas was then applied at the desired concentration via a nose mask, with
the mouse breathing spontaneously. After an equilibration time of 10 minutes, painful stimuli
in the form of pinching the tail, the interdigital webbing (pedal withdrawal reflex), the
abdominal skin, or neck skin were applied every 5 minutes for the next 30 minutes. All stimuli
were induced by the same investigator by using a blunt forceps containing a spacer between
its arms. The motor response to a painful stimulus was evaluated as positive or negative, i.e.
whether a motor response was visible or not.
Using this protocol, rough minimum alveolar concentration was first estimated within the
concentration window (1%-3% and 2%-4% for isoflurane and sevoflurane, respectively) in
Chapter 3: Inhalation Anesthesia
50
which we empirically expected the minimum alveolar concentration to lie. Concentrations
were graded in 0.25% steps and 10 animals per concentration were used.
Finally, four concentrations were chosen, and 25 animals per concentration were then tested.
Minimum alveolar concentration was then calculated as the mean of the two partial
pressures bracketing the response or lack of response in our tested population.
Anesthesia experiments
Anesthesia was induced by placing the mouse in a clear Perspex induction chamber (8x8x8
cm, volume 512 mL), into which either 5% isoflurane (Isoflo™, Abbot, Baar, Switzerland) or
8% sevoflurane (Sevorane™, Abbott, Baar, Switzerland) was then introduced. After 2
minutes, the animal was quickly transferred to a nose mask, where anesthesia was
maintained with 2.8% isoflurane or 4.9% sevoflurane, equivalent to 1.5 minimum alveolar
concentrations (as established above). Mice breathed spontaneously while lying in dorsal
recumbence on a water-filled warming mat (Gaymar, TP500, Orchard Park, NY, USA) set at
38°C±1°C.
Tail pinch, pedal withdrawal, and abdominal skin pinch reflexes were applied at 5 minutes
intervals. All reflex tests were induced by the same investigator by using a blunt forceps
containing a spacer between its arms. The reflex tests were registered as positive or
negative, i.e. whether any motor response was observable or not. Respiration rate was
counted from the movement of the thorax wall.
Anesthesia was stopped after 50 minutes by removing the nose from the mask and letting
the mouse breath room air. Two minutes later, when the mice had righted themselves from
dorsal to ventral recumbence and were able to move, they were placed back in their home
cage.
Telemetric data acquisition and analysis
Using the 16 transmitter-implanted mice, telemetric data were recorded in eight mice per
anesthetic. Mice were allocated randomly to the two groups. Telemetric data were recorded
Chapter 3: Inhalation Anesthesia
51
with the Dataquest LabPRO program (Data Sciences International, St. Paul, MN, USA). Data
collection was initiated by switching on the transmitter with a magnet. Data acquisition
started 3 days before anesthesia and continued for the 4 days following anesthesia.
To establish normal values (3 days before anesthesia) and to investigate the post-anesthetic
effects (4 days following anesthesia), heart rate and core body temperature were measured
every 5 minutes for 30 seconds and 10 seconds, respectively. Locomotor activity was
recorded continuously and stored at 5 minute intervals.
To estimate the acute effects of anesthesia (i.e. data measured during the 50-minute
anesthesia experiment), heart rate and core body temperature were recorded for 4 seconds
every 15 seconds (four measuring points of 4 seconds per minute) while administering
anesthesia. From these data, the mean values of heart rate and core body temperature were
calculated for each minute for each individual. Normal values represent means from the time
period 15:00 h to 18:00 h (i.e. the congruent time frame in which anesthesia was carried out)
during the 3 days prior to the experiment.
For analysis of long-term post-anesthetic effects, we took into account that values vary
greatly during a 24h-cycle since mice are active mainly at night. Therefore, the means of the
telemetric values for each animal were calculated separately for the night (12 h dark) and
day (12 h light) phases. An individuals’ normal values were established by calculating means
from the 3 days prior to anesthesia. For each day after anesthesia, the mean of the dark and
light phase was compared with the individual’s normal values, resulting in delta values.
Changes in body weight, and food and water intake
Body weight progression and daily food and water consumption were established from
transmitter-implanted mice for 3 days before and 3 days after anesthesia. Weights (animal,
food pellets, water bottle) were recorded with a precision balance (PR 2003 Delta Range,
Mettler-Toledo AG, Greifensee, Switzerland) especially adjusted for use with moving
animals. Body weights recorded in transmitter-bearing mice were corrected to take into
account the weight of the transmitter (3.6 g). The mean normal weights (from 3 consecutive
Chapter 3: Inhalation Anesthesia
52
daily measurements prior to the experiment) were calculated for each mouse, and compared
to the weights recorded on each of the 3 days afterwards.
Acid-base balance and blood gas concentration
Three to four weeks after minimum alveolar concentration determination, the same 48 non-
transmitter-implanted mice were used to obtain arterial blood with which to assess the acute
side effects of the anesthetics used on respiration and acid-base balance.
Arterial blood was taken under anesthesia at time points 10, 30, and 50 minutes of
anesthesia from 8 mice per anesthetic and time point. Following incision of the anterior neck,
dissection of the right common carotid artery, and cutting a small hole in the artery using a
fine-bladed pair of scissors, arterial blood was collected in a heparinized syringe. Acid-base
balance (pH), partial pressure of carbon dioxide (pCO2, mmHg) and standard bicarbonate
(HCO3, mmol/L) were determined immediately using a blood gas analyzer (AVL Compact 3,
AVL List, Graz, Austria). These animals died immediately from the subsequent rapid loss of
blood under anesthesia. The normal values of pH, pCO2, and HCO3 used for comparison had
been established previously from the arterial blood of 20 HanRcc:NMRI mice of similar age
as those used in the present study [68].
Statistical analysis
All data are presented as mean ± standard deviation. Statistical analysis using SPSS for
Windows, version 13.0 was carried out to validate the results of the post-anesthetic effects of
isoflurane and sevoflurane. One way ANOVA was performed to compare group means of
heart rate, core body temperature and locomotor activity at each of the 4 days after
anesthesia in both anesthetics with normal values. Post hoc analysis with Bonferroni tests
was carried out to identify significant differences between groups; p-values ≤0.05 were
considered significant.
Chapter 3: Inhalation Anesthesia
53
Results
Minimum alveolar concentration
Mean minimum alveolar concentration was established as 1.85% (±0.15%) for isoflurane and
3.25% (±0.14%) for sevoflurane in the adult female C57BL/6J mice used here. All anesthesia
experiments were conducted with 2.8% isoflurane or 4.9% sevoflurane, under which none of
the mice showed a motor response to testing of the pedal withdrawal reflex, tail pinch or
abdominal skin pinch.
Acute effects of anesthesia
All mice were clearly immobilized within one minute after placing in the induction chamber.
Monitoring of heart rate, core body temperature, and respiration rate during anesthesia
revealed no deviation from normal values in heart rate and core body temperature. In
contrast, the respiration rate decreased markedly below the normal values of the resting
mouse (Figure 3.1).
Acid-base balance and blood gas measurement in the arterial blood showed acidosis (i.e.
decrease of pH) and hypercapnia (i.e. increase of pCO2) at 10, 30 and 50 minutes of
anesthesia (Figure 3.2).
When anesthesia was completed, i.e. when the nose mask was removed, animals showed
increasing respiration rate and muscle rigor within one minute. Mice turned to sternal
recumbence and showed spontaneous movement at 1-2 minutes after anesthesia was
withdrawn.
Post-anesthetic effects
Telemetric measurements revealed a significant increase in heart rate and core body
temperature for 0-12 hours after anesthesia compared to normal values (i.e. before
anesthesia) with both anesthetics. In this time frame, locomotor activity also showed a
Chapter 3: Inhalation Anesthesia
54
tendency to increase, but this was not significant for either anesthetic (Figure 3.3). Statistical
comparison of isoflurane vs. sevoflurane regarding their long-term effects on heart rate, core
body temperature and locomotor activity revealed no difference between these two
anesthetics.
Body weight with both anesthetics was constant in the post-anesthetic phase, i.e. mean body
weights varied with ≤0.5% (±0.01%) compared to the normal body weight before anesthesia.
The mean daily food intake showed a decrease of 10% (±0.2%) at the first post-anesthetic
day in both anesthetics. At the second post-anesthetic day, mean food consumption was
almost unchanged, with a decrease of 1.5% (±0.2%) in both anesthetics. At the third post-
anesthetic day, food intake was decreased with 5% (±0.1%) in isoflurane and 9% (±0.1%) in
sevoflurane. Water consumption showed high interindividual variability. The mean water
consumption in both anesthetics showed an increase of 6% (±19% for isoflurane; ±25% for
sevoflurane) at the first day after anesthesia. Mean water intake ranged from a decrease of
1.5% to an increase of 8.7% with standard deviations ranging from 12% to 19% at the
second and third post- anesthetic days in both anesthetics. The alterations in body weight,
food and water intake were not statistically significant.
Chapter 3: Inhalation Anesthesia
55
Figure 3.1) Heart rate, core body temperature and respiration rate during 50 min of
anesthesia with isoflurane or sevoflurane. The grey areas indicate normal values at the
corresponding time of day in conscious animals. Data represent the mean values of eight
mice; bars represent standard deviation
[bre
ath
s p
er
min
ute
][°
C]
[bp
m]
induction chamber
400
450
500
550
600
650
700
750
800
35
36
37
38
39
350
duration of anaesthesia [min]
respiratory rate
core body temperature
heart rate
maintenance with nose mask (spontaneous breathing) in dorsal recumbance on the warmed plate
0
20
40
60
80
100
120
140
160
180
0 5 10 15 20 25 30 35 40 45 50
Isoflurane
Sevoflurane
Normal
Chapter 3: Inhalation Anesthesia
56
Figure 3.2) Acid–base balance (pH), partial pressure of carbon dioxide ( pCO2) and
standard bicarbonate (HCO3) in arterial blood at 10, 30 and 50 min of anesthesia. Hatched
areas indicate the normal values established in a previous study (68). Data represent the
mean values of eight mice; bars indicate standard deviation
pH
7.1
7.15
7.2
7.25
7.3
7.35
7.4
7.45
0
10
20
30
40
50
60
70
80
90
10 min 30 min 50 min
pC
O2 [m
mH
g]
10 min 30 min 50 min
HC
O3 [m
mo
l/L
iter]
0
5
10
15
20
25
30
10 min 30 min 50 min
Isoflurane
Sevoflurane
Normal
Chapter 3: Inhalation Anesthesia
57
Figure 3.3) Post-anesthetic measurements of the impact of isoflurane and sevoflurane on
heart rate, core body temperature and locomotor activity. Delta (D) values represent
deviation from normal values (established prior to anesthesia) at the corresponding 12 h day
and night time. Data represent the mean from eight mice; bars indicate standard deviation.
Asterisks indicate significance at P ≤ 0.05
-1
-0.8
-0.6
-0.4
-0.2
0
0.2
0.4
0.6
0.8
1
-60
-40
-20
0
20
40
60
80
100
120[°
C]
[bp
m]
heart rate
[% c
ou
nts
]
core body temperature
day 2day 1 day 3 day 4
Isoflurane
Sevoflurane
-100
-50
0
50
100
150
200 locomotor activity
**
*
*
anaesthesia
Chapter 3: Inhalation Anesthesia
58
Discussion
Prior to these investigations, the anesthetic procedure and dosage of anesthetics were
standardized based on the specifics of the equipment (e.g., calibrating the gas concentration)
and mouse population (e.g., strain, age, gender) used. Therefore, minimum alveolar
concentrations were established following the widely accepted method of determining the
concentration of anesthetic gas at which 50% of the animals fail to respond with purposeful
movements to the testing of reflexes. The minimum alveolar concentration determined for
sevoflurane in female C57BL/6J mice was almost identical to that described for male outbred
mice (3.25% in female C57BL/6J vs. 3.22% in male CD-1) [90]. The minimum alveolar
concentration is known to vary considerably between mouse strains [88], but other factors
such as age and gender can also influence anesthetic potency. Gender differences may
explain why the minimum alveolar concentration for isoflurane was found to be higher
(1.85%) than values described in the literature for C57BL/6J mice (1.30% [89] and 1.34%
[91]). In former reports, male mice [89], or a mixture of both sexes [91] were used, whereas
in our experiments only female mice were investigated. On the other hand, the suggestion
that anesthetic requirements decrease with age, and literature reports of lower minimum
alveolar concentrations in younger mice (7--9 weeks [89]), 6--12 weeks [91]) are in contrast
to the 0.5% increase found in our study, in which older mice (16--36 weeks) were used.
However, when comparing minimum alveolar concentrations established in different
laboratories, technical aspects of how the gas was provided must be considered. The easy-
to-use open anesthesia system used here might include an uncertainty in the absolute
values of the gas concentration measurements.
After preliminary standardization of the dosages by establishing the minimum alveolar
concentrations, the anesthetics were then compared with dosages of isoflurane and
sevoflurane representing 1.5 minimum alveolar concentrations. For this dosage it is generally
postulated that 99.9% of animals will not react to painful stimuli [22, 92], i.e. that the animals
have reached surgical tolerance. However, since analgesia was not proved with
sophisticated methods such as measurements of the reaction in heart rate, blood pressure or
respiration upon a substantially painful stimulus (e.g. skin incision), surgical tolerance cannot
be definitively claimed from our study. However, motor reflex responses were suppressed in
Chapter 3: Inhalation Anesthesia
59
all animals and none died from the 50-minute inhalation anesthesia with 1.5 minimum
alveolar concentrations (i.e. 2.8% isoflurane or 4.9% sevoflurane).
In the early induction phase of anesthesia, heart rate peaked at the upper normal level of
700-800 bpm (beats per minute), which we suggest as a normal reaction to removing the
animal from its cage and placing it in a foreign environment. During anesthesia, heart rate
was stable within the normal values of the resting mouse (490-550 bpm). This was in
agreement with recent publications indicating only slight depression of heart rate for
isoflurane anesthesia [64, 77, 93-95].
During 50 minutes of inhalation anesthesia, core body temperature could be maintained with
a simple, water-bath-driven warming mat. That core body temperature falls due to any kind of
anesthesia is well-known, and mice are especially sensitive to hypothermia due to their small
size and high body surface area (e.g. a drop to 30-31°C was shown following isoflurane
anesthesia in mice [64]). Obviously, such hypothermia should be prevented, because it
influences physiology and the course of anesthesia and can ultimately lead to the death of
the animal. Thus, warming the animal has been common practice for years, and is
particularly worthwhile in long-term anesthesia in mice [96-98].
In contrast to the almost normal levels of heart rate and core body temperature, respiratory
depression, as evidenced by decreased respiration rates (far below the values of the resting
mouse), marked hypercapnia and acidosis, were seen to an equal extent with both isoflurane
and sevoflurane anesthesia. Hypercapnia and acidosis associated with isoflurane has also
been found by others [77], but was less intense than with injection anesthesia with
pentobarbital [95]. Other studies also report a marked decrease in respiration rate upon
isoflurane anesthesia, but values reported in the literature [96, 98] were not as low as found
here with both anesthetics.
Thus, respiratory depression was the major adverse effect observed with both isoflurane and
sevoflane. In general, the influence of inhalational anesthetics on respiratory function is well-
known. By inhibiting the control systems of respiration (e.g. feed back control of central
respiratory centers, various chemoreceptors, pulmonary reflexes and neuronal input)
inhalational anesthetics alter oxygen supply and CO2 elimination [99]. This is mirrored by
Chapter 3: Inhalation Anesthesia
60
aberrations in arterial blood gas levels (e.g. increase of pCO2, decrease of pO2) and thus
hampers the ability of the organism to maintain cellular homeostasis. Respiratory depression
is often accompanied by acid-base imbalance (e.g. alterations of pH and HCO3), and
changes in the depth and frequency of respiration. Since respiratory depression is the most
probable emergency situation when using isoflurane or sevoflurane anesthesia, it may be
useful to monitor the respiration rate as an indirect indicator of impaired respiration and thus
prevent fatal outcomes under routine conditions. In addition, administering oxygen instead of
room air in the recovery phase (i.e. after the volatile anesthetic ceased) may be beneficial for
animals with impaired respiration.
The time required for induction and recovery were almost the same for both anesthetics (1-2
minutes). The short recovery time from both inhalational anesthetics should be highlighted as
a distinct advantage compared to injection anesthesia. The benefits of fast recovery include
reducing postoperative complications associated with prolonged inability to correct
physiological impairment (e.g. hypothermia, hypoglycemia, dehydration) that may induce
suffering and hamper the rapid return of the animal to its normal state.
Heart rate and core body temperature increased significantly in the 12 hours following the
50-minute anesthesia. Locomotor activity showed a tendency to increase only in the first
hours after anesthesia, suggesting that physical activity is not the reason for the elevated
heart rate and core body temperature. Alterations in body weight progression as well as daily
food and water consumption are known to hint at pain, distress or impaired well-being in
laboratory animals [100, 101]. Post anesthetic determination of these indices revealed no
relevant aberrations, indicating the negligible impact of a 50-minute anesthesia with
isoflurane or sevoflurane on body weight, food and water intake. Although it is unclear why
the post anesthetic elevation in heart rate and core body temperature occurred, it appeared
to be of no long-term detriment to the animals.
Considering the effects of isoflurane and sevoflurane on physiology and behavior, we found
that both agents exerted a similar impact on the normal state established in the un-
anesthetized mouse. Whereas several publications describe the usefulness of isoflurane,
there is limited description of sevoflurane in mice in the literature. Henke and co-authors
compared induction and recovery times, and respiration rates of sevoflurane with those of
Chapter 3: Inhalation Anesthesia
61
isoflurane in the gerbil. Although they found prolonged recovery from isoflurane compared to
sevoflurane, they concluded that there is no overall preference for one of these two volatile
anesthetics over the other [96]. Another study compared blood glucose and some specific
parameters required in functional PET investigations in mice; sevoflurane was considered
superior compared to isoflurane, and the former was consequently recommended for
physiologic imaging by Flores and co-authors [102]. From our results, neither anesthetic was
clearly superior over the other.
In summary, we conducted inhalation anesthesia in a routine, cost-effective setting, using
commercially available equipment. The anesthesia experiments were standardized by
establishing minimum alveolar concentrations, i.e. the dosage was adjusted to the
characteristics of the animals used (female C57BL/J mice, aged 16–36 weeks). Both volatile
anesthetics tested showed short induction and recovery times in an easy-to-manage,
standard inhalation anesthesia procedure. During anesthesia, the most prominent adverse
effect was respiratory depression. Hypothermia, which generally occurs under anesthesia,
was prevented by placing the animal on a warmed mat. After completion of anesthesia,
altered physiological functions, such as elevated heart rate and core body temperature,
persisted for approximately half a day. In conclusion, both isoflurane and sevoflurane
provided an equally effective anesthesia with acceptable adverse effects.
Acknowledgement
This work was sponsored by the ECLAM and ESLAV Foundation. The authors would like to
thank Robin Schneider and the staff of the central biological laboratory for support in housing
mice. We thank Professor Kurt Burki for generously providing research facilities and
resources.
Chapter 4: Balanced Anesthesia
62
Combining sevoflurane anesthesia with fentanyl-
midazolam or s-ketamine in laboratory mice
Based on Cesarovic N, Jirkof P, Rettich A, Nicholls F, Arras M.
Published Mar. 2012 in Journal of the American Association for Laboratory Animal Science
(JAALAS)
Chapter 4: Balanced Anesthesia
63
Abstract
Laboratory mice typically are anesthetized by either inhalation of volatile anesthetics or
injection of drugs. Here we compared the acute and postanesthetic effects of combining both
methods with standard inhalant monoanesthesia using sevoflurane in mice. After injection of
fentanyl–midazolam or S-ketamine as premedication, a standard 50-min anesthesia was
conducted by using sevoflurane. Addition of fentanyl–midazolam (0.04 mg/kg–4 mg/kg)
induced sedation, attenuation of aversive behaviors at induction, shortening of the induction
phase, and reduced the sevoflurane concentration required by one third (3.3% compared
with 5%), compared with S-ketamine (30 mg/kg) premedication or sevoflurane alone. During
anesthesia, heart rate and core body temperature were depressed significantly by both
premedications but in general remained within normal ranges. In contrast, with or without
premedication, substantial respiratory depression was evident, with a marked decline in
respiratory rate accompanied by hypoxia, hypercapnia, and acidosis. Arrhythmia, apnea, and
occasionally death occurred under S-ketamine–sevoflurane. Postanesthetic telemetric
measurements showed unchanged locomotor activity but elevated heart rate and core body
temperature at 12 h; these changes were most prominent during sevoflurane
monoanesthesia and least pronounced or absent during fentanyl–midazolam–sevoflurane. In
conclusion, combining injectable and inhalant anesthetics in mice can be advantageous
compared with inhalation monoanesthesia at induction and postanesthetically. However,
adverse physiologic side effects during anesthesia can be exacerbated by premedication,
requiring careful selection of drugs and dosages.
Introduction
Laboratory mice frequently are anesthetized by subcutaneous or intraperitoneal injection of
hypnotic, analgesic, and muscle relaxing agents [40]. Although easy, practical and cost-
effective, this method has its drawbacks. After injection of relatively long-acting drugs
through the subcutaneous or intraperitoneal route, the course and depth of anesthesia is
nearly uncontrollable once the initial dose has been administered. In addition, due to the
considerable variability in dose requirements for mice of different age, strain, sex, and other
specifics (for example, circadian rhythm, sociophysiologic conditions), the margin between
reaching a state of anesthesia sufficiently deep to provide surgical tolerance and a lethal
Chapter 4: Balanced Anesthesia
64
outcome is usually narrow [68]. Moreover, most injection anesthesia protocols induce a
prolonged recovery period accompanied by hypothermia and compromised physiologic
function.
Such problems rarely are encountered with inhalation anesthesia, because this method has
a short recovery phase and accommodates control of the duration and depth of anesthesia,
including expeditious adjustment of the dosage of inhalation anesthetics tailored to the
requirements of the individual animal. Therefore, in terms of survival rate, inhalation
anesthesia generally is suggested to be safe in mice. However, negative effects on the
cardiovascular system combined with depression of respiration are well-known side effects of
halogenated volatile anesthetics [77, 103, 104]. This situation, coupled with the fact that the
analgesia provided by monoanesthesia with volatile anesthetics is still controversial [105,
106], justifies a continued search for improvement.
By taking advantage of the well-known synergistic and additive interactions between
injectable drugs (analgesics or sedatives) and volatile anesthetics, the dosages of each
component can be decreased (relative to its use as a sole agent) while inducing general
anesthesia of sufficient depth with fewer side effects [107-109]. This approach, sometimes
referred to as ‘balanced’ or ‘modular’ anesthesia [110] is used widely in human and
veterinary medicine—but only recently has it has begun to be used in mice. Therefore, in the
present study, 2 protocols of combined injection and inhalant anesthesia in laboratory mice
were established and compared with a standard protocol of inhalant monoanesthesia with a
commonly used volatile anesthetic.
Isoflurane and sevoflurane are the 2 of the volatile anesthetics most widely used in human
and veterinary anesthesia. We decided to use sevoflurane to provide rapid induction and
recovery. Because we considered that volatile anesthetics offer suboptimal analgesia, we
focused on injectable agents that could provide sufficient analgesia to complement inhalant
anesthesia. Ketamine is known for its ability to cause profound analgesia, which can occur
even at subhypnotic dosages—particularly if the S(+)-enantiomer of ketamine is administered
[33]. Therefore, we chose S-ketamine for injection in one protocol. We calculated the dosage
based on literature reports [33, 111-113], with the aim of minimizing side effects such as
catalepsy, slight respiratory depression, and stimulation of locomotor activity (restlessness)
while inducing analgesia and taking advantage of the hypnotic and cardiovascular
stimulatory effects of ketamine [34, 111]. Our second approach to combining inhalation
Chapter 4: Balanced Anesthesia
65
anesthesia with injectable agents in mice was based on drugs that are used widely in human
medicine, namely fentanyl and midazolam. Midazolam, which often is applied as
premedication to anesthesia, belongs to the benzodiazepines, which typically induce
sedation, anxiolysis, and muscle relaxation [114]. Antinociceptive effects of midazolam have
been reported in mice8 and rats [115]. In humans, benzodiazepines frequently are
administered with opioids to improve pain relief. Therefore, we combined midazolam with
fentanyl - a potent synthetic opioid analgesic. Among the typical side effects of opioids [41],
sedation, hypothermia, respiratory depression, and hypercapnia could be of relevance for the
use of fentanyl during anesthesia in mice. Although opioids can cause bradycardia,
vasodilation, and hypotension, they have mostly only mild effects on cardiovascular function.
In addition, their effects on the genitourinary system and gastrointestinal tract [41] (for
example, constipation) are suggested to be tolerable side effects, which may be of only
minor relevance for establishing an anesthesia protocol in mice. Fentanyl often is
administered as an intravenous constant-rate infusion in the context of anesthesia in humans
[116], but this technique is complex and difficult to manage in mice. Therefore, we attempted
to achieve preemptive analgesia with subcutaneous injection of fentanyl with midazolam as
premedication, with dosages selected on the basis of anecdotal evidence, clinical
experience, and hints from the literature [117].
To compare the 3 anesthesia protocols, 50 min of sevoflurane inhalant anesthesia was
conducted either alone (S) or with subcutaneous injection of S-ketamine (KS) or a mixture of
fentanyl and midazolam (FMS). Injections were administered as premedication, and their
effects on behavior during induction of anesthesia and on the sevoflurane concentration
required were noted. During anesthesia, heart rate, core body temperature, respiratory rate,
arterial blood gases, and arterial pH were monitored. The long-term effect of the 3 protocols
on recovery from anesthesia was investigated through telemetric measurements of heart
rate, core body temperature, and locomotor activity for 3 d after anesthesia.
Chapter 4: Balanced Anesthesia
66
Materials and Methods
Animals and housing conditions
Female C57BL/6J mice (n = 98; age, 6 wk) were obtained from our in-house breeding
colony. The 72 mice used for determination of minimal alveolar concentrations were later
euthanized to obtain arterial blood for measuring acid - base balance and blood gas
concentrations. The remaining 26 mice were implanted with telemetric transmitters prior to
the experiments to allow measurement of heart rate, core body temperature, and locomotor
activity. The mice were free of all viral, bacterial, and parasitic pathogens listed in the
FELASA recommendations [86]. Health status was monitored by a sentinel program
throughout the experiments.
Mice generally were housed in pairs; each transmitter-implanted mouse was housed with a
non-implanted companion of the same strain, sex, and age. Mice 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 aspen bedding (80 to 90 g per cage;
LTE E-001 Abedd, Indulab). Autoclaved hay (8 to 12 g per cage) and 2 cotton nesting pads
(each 5 × 5 cm; Nestlets, Indulab) were provided as nesting materials. A standard cardboard
house (Ketchum Manufacturing, Brockville, Canada) served as a shelter. Mice were fed a
pelleted mouse diet (3431, Provimi Kliba, Kaiseraugst, Switzerland) ad libitum and had
unrestricted access to sterilized drinking water provided in a water bottle. The 12:12-h
light:dark cycle in the room was established with artificial light (approximately 40 lx in the
cage; lights on, 0300 to 1500). The temperature was 21 +/- 1 °C, with a relative humidity of
50% +/- 5% and 15 complete changes of HEPA-filtered air hourly. To avoid interfering
influences, all necessary husbandry and management procedures were completed in the
room at least a d before the start of an experiment or data recording, and disturbances (for
example, visitors or unrelated experimental procedures) were not allowed. The animal room
was insulated to exclude electronic noise.
The study and all procedures and protocols were approved by the Cantonal Veterinary Office
(Zurich, Switzerland) under license number 111/2007. Housing and experimental procedures
were in accordance with Swiss animal protection law and conformed to the European
Chapter 4: Balanced Anesthesia
67
Convention for the protection of vertebrate animals used for experimental and other scientific
purposes (Council of Europe nr.123 Strasbourg 1985) [118]. Housing and experimental
procedures also were in accordance with the Guide for the Care and Use of Laboratory
Animals22 and conformed to the AALAS position statement on the humane care and use of
laboratory animals.
Transmitter implantation
Prior to the experiments, at age 10 wk, 26 mice were instrumented with telemetric
transmitters (TA10ETA-F20, Data Sciences International, St Paul, MN) to measure heart
rate, core body temperature, and locomotor activity in freely moving mice [62, 63]. Briefly,
mice were anesthetized with sevoflurane (Sevorane, Abbott, Baar, Switzerland), and the
transmitter body was implanted in the abdomen under aseptic conditions. One wire-loop
electrode was fixed with silk sutures (6-0 Perma-Handseide, Ethicon, Norderstedt, Germany)
between the muscles located to the right of the trachea, and the other loop lead was sutured
to the xiphoid process. Muscle layers and skin were closed with absorbable sutures (6-0
Vicryl, Ethicon). Postoperative pain was treated with flunixine (5 mg/kg SC twice daily;
Biokema Flunixine, Biokema SA, Crissier-Lausanne, Switzerland) for 4 d [59]. After
transmitter implantation, mice were allowed to recover for 6 wk before the first experiment.
Experimental setting
All experiments were conducted when the mice were 16 to 36 wk of age, with body weights
ranging from 25 to 30 g. To avoid any influence of circadian rhythm, all experiments and
weighing procedures were done between 15:00 and 18:00. The study was designed for the
experiments and anesthesia to be performed at the beginning of the dark phase for these
mice. Anesthesia was performed in a separate operating area within the animal room to
avoid transportation of the mice and to ensure stable conditions of humidity, air pressure,
and room temperature and sufficient removal of gases and smells through the ventilation
system.
The method of delivering inhalation anesthesia was modified slightly from that described
elsewhere [26]. Briefly, sevoflurane was provided by using a commercially available rodent
inhalation anesthesia apparatus (Provet, Lyssach, Switzerland), which was equipped with a
Chapter 4: Balanced Anesthesia
68
sevoflurane vaporizer (Ohmeda Sevotec 5, Abbott, Baar, Switzerland) and a pump-driven
filter system to eliminate waste anesthetic gas. As carrier gas, pressurized air was used at a
flow rate of 600 mL/min. The anesthetic gas was introduced into the induction chamber or
nose mask (Figure 4.1).
Premedications
As injectable drugs, we used fentanyl (0.04 mg/kg s.c; Kantonsapotheke Zurich, Zurich,
Switzerland) mixed with midazolam (4 mg/kg s.c; Dormicum, Roche Pharma Schweiz AG,
Reinach, Switzerland) in one protocol and Sketamine (30 mg/kg s.c; Keta-S, Graeub AG,
Bern, Switzerland) in another. All drugs were dissolved in PBS immediately before injection
in such a manner that dosing could be achieved by application of an injection volume of 2
μL/kg body weight. Injections were provided as premedication, that is, 5 to 7 min before
sevoflurane anesthesia was induced.
Determination of minimum alveolar concentration
Sevoflurane inhalation anesthesia was standardized by establishing minimum alveolar
concentrations during sevoflurane monoanesthesia and after the premedications described
earlier. To this end, we anesthetized 72 nontransmitter-implanted mice 2 to 4 times each;
care was taken that mice had a break of at least 2 wk between tests.
Minimal alveolar concentration was determined according to commonly accepted procedures
used in mice [26, 87-89, 119, 120]. For each protocol, 4 consecutive sevoflurane
concentrations differing by 0.25% were tested; 25 mice were tested per concentration. The
bracketed study design [121] was adapted to our anesthesia protocols to measure minimal
alveolar concentration at a defined time point of anesthesia, that is, at 12 min after inducing
inhalant anesthesia (equivalent to 17 to 19 min after subcutaneous injection of
premedication). Thus, after inducing sevoflurane anesthesia in the induction chamber (Figure
4.1 A) for 1.5 min at a maximal concentration of 8% sevoflurane, the mouse was taken out of
the chamber and placed in dorsal recumbence on a warmed mat. Anesthetic gas then was
applied at the test concentration by using a nose mask, with the mouse breathing
spontaneously (Figure 4.1 B). After an equilibration time of 10 min, 3 noxious stimuli were
applied sequentially: pinching of the tail (tail pinch reflex), interdigital webbing (pedal
Chapter 4: Balanced Anesthesia
69
withdrawal reflex), and abdominal skin (abdominal skin pinch reflex). All stimuli were
induced by the same investigator by using blunt forceps with a spacer between its arms to
allow uniform application of pressure. Any motor response (for example, movement of the tail
or an extremity, head jerking) to one or more of the 3 noxious stimuli was judged as
purposeful movement, indicating that sevoflurane at the concentration applied did not induce
anesthesia in the mouse evaluated. After testing the response to the 3 noxious stimuli (that
is, after 12 to 13 min of inhalant anesthesia), administration of the anesthetic gas ceased,
and the mouse was allowed to recover. By using the responses to the noxious stimuli, the
mouse’s minimal alveolar concentration was calculated as the average of the 2 partial
pressures bracketing the positive response (that is, purposeful movement) or lack of
response in the animal.
Anesthesia experiments
Mice were allocated randomly to 1 of 3 anesthesia protocols. The 3 protocols consisted of
fentanyl (0.04 mg/kg) and midazolam (4 mg/kg) as premedication and 3.3% sevoflurane
(FMS); S-ketamine (30 mg/kg) as premedication and 5% sevoflurane (KS); and 5%
sevoflurane as monoanesthesia (S). After premedication in the FMS and KS protocols, the
mice were examined for 5 to 7 min in their home cage for behavioral aberrations. Inhalant
anesthesia then was induced by placing each mouse in a clear induction chamber (8 × 8 × 8
cm; volume, 512 mL) into which 8% sevoflurane (Sevorane, Abbott, Baar, Switzerland) was
introduced. The mouse’s behavior in the induction chamber and the time point at which it
became immobile were observed and noted. After 1.5 min, the mouse was transferred
rapidly to the nose mask, through which anesthesia was maintained with sevoflurane. Mice
breathed spontaneously while lying in dorsal recumbence on a water-filled warming mat
(Gaymar, TP500, Orchard Park, NY) set at 39 °C +/- 1 °C.
Tail pinch, pedal withdrawal, and abdominal skin pinch reflexes each were tested at 5-min
intervals. All reflex tests were induced by the same investigator by using blunt forceps with a
spacer between its arms to allow uniform application of pressure. The reflex tests were
registered as positive or negative (that is, whether any motor response was present or not).
Respiratory rate was counted from the movement of the thorax wall and recorded at 5-min
intervals. During anesthesia, mice were observed for any abnormality in their respiratory
Chapter 4: Balanced Anesthesia
70
rhythm. In addition, heart rhythm alterations were monitored by using real-time telemetric
electrocardiograms.
Anesthesia was stopped after 50 min by removing the nose from the mask and letting the
mouse breathe room air. Mice were left on the warming mat and allowed to recover from
anesthesia for 10 min before being placed back in their home cages.
Telemetric data acquisition and analysis
Telemetric data were recorded from 8 mice per anesthetic protocol by using the Dataquest
LabPRO program (Data Sciences International). Data collection was initiated by switching on
the transmitter by using a magnet. Data acquisition started 3 d before anesthesia and
continued for 3 d after anesthesia.
To estimate the acute effects of anesthesia (that is, after premedication and during
anesthesia), heart rate and core body temperature were recorded for 4 s every 15 s (4 data
points of 4 s per minute). From these data, mean values of heart rate and core body
temperature were calculated for each minute for each mouse. Baseline values represent
means from 1500 to 1800 (that is, the same time frame during which anesthesia occurred)
during the 3 d prior to the experiment.
To establish baseline values (3 d before anesthesia) and to investigate postanesthetic effects
(3 d after anesthesia), heart rate was measured for 30 s every 5 min, and core body
temperature was measured for 10 s every 5 min. Locomotor activity was recorded
continuously and stored at 5-min intervals.
For analysis of long-term postanesthetic effects, we took into account that values vary greatly
during a 24-h cycle because mice are active mainly at night. Therefore, the means of the
telemetric values for each mouse were calculated separately for the 12-h dark (night) and 12-
h light (day) phases. A mouse’s baseline values were established by calculating means from
the 3 d prior to anesthesia. For each day after anesthesia, a mouse’s baseline value was
subtracted individually from its daytime and nighttime means; the differences are reported as
delta (Δ) values.
Chapter 4: Balanced Anesthesia
71
Changes in body weight
Body weight in transmitter-implanted mice was monitored for 3 d before and 3 d after
anesthesia. Weights were obtained by using a precision balance (PR 2003 Delta Range,
Mettler-Toledo AG, Greifensee, Switzerland) that specifically was adjusted for use with
moving animals. Body weights were corrected to account for the weight of the transmitter
(3.6 g). Mean baseline weight (from 3 consecutive daily measurements prior to the
experiment) was calculated for each mouse and compared with that recorded on each of the
3 d after the experiment.
Acid-base balance and blood gas concentration
At 3 to 4 wk after determination of minimal alveolar concentration determination, arterial
blood was collected from the same 72 non-implanted mice to assess acute effects of
anesthesia on respiration and acid–base balance. Arterial blood was obtained after 10, 30,
and 50 min of anesthesia from 8 mice per anesthetic protocol and time point.
Blood sampling and analyses were carried out as described previously [26, 68]. Briefly, the
anterior neck was incised, the right common carotid artery was dissected out, a small hole in
the artery was created by using fine-blade scissors, and arterial blood was collected in a
heparinized syringe. Acid–base balance (pH), pCO2 (mm Hg), and pO2 (mm Hg) were
determined immediately by using a blood-gas analyzer (Compact 3, AVL List, Graz, Austria).
These mice died immediately due to the subsequent rapid loss of blood under anesthesia.
Reference values of pH, pCO2, and pO2 for comparison had been established by using
arterial blood from 20 HanIbm:NMRI mice that were similar in age to those in the current
study [68].
Chapter 4: Balanced Anesthesia
72
Figure 4.1) (A) Chamber for induction of sevoflurane inhalation anesthesia. (B) Nose mask
for maintaining sevoflurane anesthesia. The mask was equipped with a latex membrane,
which had a hole in the center that fit around the nose of each mouse, with the dual purpose
of preventing both withdrawal of environmental air into the nose mask and leakage of
anesthetic gas from it.
Statistical analysis
All data are presented as mean +/- 1 SD. Statistical analysis (version 17.0, SPSS for
Windows, SPSS, Chicago, IL) was done to validate the results. All data were tested for
gas inlet
exhaustion
latex membrane
gas inlet
A
B
Chapter 4: Balanced Anesthesia
73
normal distribution and homogeneity of variance and met the necessary assumptions for
parametric analyses. One-way ANOVA was performed to compare group means of minimal
alveolar concentrations and time until immobilization as well as heart rate, core body
temperature, and locomotor activity at each of the first 3 d after anesthesia. Post hoc
analysis with Bonferroni tests was done to identify significant differences between groups.
For comparison of baseline values with corresponding experimental group means of heart
rate, core body temperature, and locomotor activity during and at each of the 3 d after
anesthesia, a dependent t test for paired samples was used. P values less than or equal to
0.05 were considered significant.
Results
Minimum alveolar concentration
The minimal alveolar concentration (mean +/- 1 SD) for sevoflurane monoanesthesia and
with premedication using S-ketamine in adult female C57BL/6J mice was 3.3% +/- 0.18%
(Figure 4.2 A). Premedication with fentanyl – midazolam significantly (P = 0.0005) decreased
the mean minimal alveolar concentration for sevoflurane to 2.2% +/- 0.27% compared with
that for the other 2 protocols. This decrease represents a gas savings of 33%.
We considered that providing sevoflurane at 1.5 times the minimal alveolar concentration
would prevent mice from responding to noxious stimulation (that is, surgical tolerance is
achieved). Therefore all subsequent anesthesia experiments were conducted by using 3.3%
sevoflurane after fentanyl–midazolam premedication but by using 5% sevoflurane after S-
ketamine premedication and during sevoflurane monoanesthesia.
Chapter 4: Balanced Anesthesia
74
Figure 4.2) (A) Mean (n = 50 mice; bar, 1 SD) minimum alveolar concentrations for
sevoflurane in adult C57BL/6J female mice. The gas-saving effect is evident from the
decrease in minimum alveolar concentration seen after fentanyl–midazolam premedication
with sevoflurane (FMS) compared with S-ketamine premedication with sevoflurane (KS) and
sevoflurane alone (S). *, P ≤ 0.05 between values. (B) The mean time (n = 8 mice; bar, 1 SD)
required until immobilization after mice were placed in the sevoflurane-filled induction
chamber differed between all protocols. *, P ≤ 0.05 between values.
Induction of anesthesia
Approximately 2 min (107.5 +/- 18.3 s) after injection with fentanyl–midazolam, all mice
showed signs of sedation (for example, absence of locomotion and stationary activity, sleep-
like posture). Approximately 5 min (306 +/- 55.8 s) after injection with S-ketamine, all mice
exhibited symptoms of tremor, ataxia, and dizziness.
When placed in the induction chamber, most nonpremedicated mice (that is, the sevoflurane
monoanesthesia group) showed behaviors including defecation, urinating, shaking the head
or limbs, jumping, and locomotion (Table 4.1). These behaviors were less frequent after S-
ketamine premedication and were nearly totally absent after fentanyl–midazolam
0
0.5
1
1.5
2
2.5
3
3.5
4
sevo
flu
ran
e [
%]
FMS KS S
0
10
20
30
40
50
60
70
80
tim
e [
sec]
FMS KS S
*
*
*
* *4.5 90
A B
Chapter 4: Balanced Anesthesia
75
premedication. One transmitter-implanted mouse died after S-ketamine premedication when
the animal was exposed to sevoflurane in the induction chamber; this animal was replaced.
The time until immobilization differed among all 3 protocols. The shortest time was
associated with the FMS protocol and the longest with sevoflurane monoanesthesia (FMS
compared with S, P = 0.0005; FMS compared with KS, P = 0.0005; KS compared with S, P =
0.004; Figure 2 B).
Anesthesia
(8 mice per
protocol)
Percentage of animals showing these reactions
Locomotion
with or
without ataxia
Jumping
Shaking
head
and/or
limbs
Urination Defecation Apnea/death
FMS 12.5 0 0 0 0 0
KS 100 0 0 37.5 0 12.5
S 100 50 100 100 62.5 0
Table 4.1) Behaviors of mice (n = 8 per protocol) during induction of anesthesia with
sevoflurane.
Effects during anesthesia
During anesthesia, none of the mice showed any motor response to testing of the pedal
withdrawal reflex, tail pinch, or abdominal skin pinch.
During the 50-min anesthesia period in all 3 protocols, heart rate and core body temperature
remained within the general physiologic boundaries for this species (350 to 800 bpm, 35 to
38 °C; Figure 4.3). Heart rate was 446 +/- 51 bpm during FMS anesthesia, 470 +/- 59 bpm
during KS anesthesia, and 519 +/- 60 bpm during sevoflurane monoanesthesia. Compared
with the mean baseline heart rate at the corresponding time of day (525 +/- 80 bpm), the
decreases in heart rate during FMS (P = 0.001) and KS (P = 0.030) were significant.
Compared with the baseline core body temperature at the same time of day (36.8 +/- 0.7 °C),
core body temperature was decreased significantly during FMS (35.4 +/- 0.6 °C; P = 0.0005)
Chapter 4: Balanced Anesthesia
76
and KS (35.4 +/- 0.4 °C; P = 0.0005) anesthesia. Core body temperature showed a tread
toward a decrease during sevoflurane monoanesthesia (36.1 +/- 0.7 °C; P = 0.058).
During all 3 protocols, the respiratory rate declined immediately after the onset of sevoflurane
anesthesia and remained markedly depressed during the 50-min anesthesia session
compared with baseline respiration in resting mice at the same time of day (150 +/- 10
breaths per minute; Figure 4.3). The respiratory rate during anesthesia was 68.5 +/- 7.7
breaths per minute for FMS, 48.8 +/- 5.4 breaths per minute for KS, and 44 +/- 5.1 breaths
per minute for sevoflurane monoanesthesia. After 10, 30 and 50 min of anesthesia in all 3
protocols, blood gas and pH measurements of arterial blood showed prominent acidosis,
hypercapnia, and hypoxia, with values markedly exceeding the physiologic range (Figure
4.4).
During anesthesia with KS, all mice displayed cardiac arrhythmia and episodes of apnea
followed by tachypnea. None of these events occurred during either of the other 2 protocols.
One transmitter-implanted mouse died at 15 min into KS anesthesia and was replaced.
Mice began showing increasing respiratory rate and muscle rigor within 1 min after
sevoflurane was discontinued. In all 3 protocols, the mice had turned to ventral recumbence
and were able to move within approximately 2 min after sevoflurane withdrawal.
Effects during the first 3 days after anesthesia
Compared with baseline values, telemetric measurements revealed a significant (P = 0.0005)
increase in heart rate during the first 12 h after anesthesia in all 3 protocols (Figure 4.5).
Comparing between protocols, the increase in heart rate after sevoflurane monoanesthesia
was significantly (P = 0.0005) higher than that after FMS anesthesia, whereas heart rate after
KS anesthesia did not differ significantly from that in the other 2 protocols.
Compared with baseline values, core body temperature increased significantly during the first
12 h after anesthesia with KS anesthesia (P = 0.005) and sevoflurane monoanesthesia (P =
0.0005) but not after FMS anesthesia (Figure 4.5). Core body temperature was significantly
higher after sevoflurane monoanesthesia compared with KS and FMS (S compared with
FMS, P = 0.0005; S compared with KS, P = 0.006). Locomotor activity and body weight were
unchanged in all groups after anesthesia relative to baseline values before anesthesia.
Chapter 4: Balanced Anesthesia
77
Figure 4.3) Mean (n = 8 mice; bar, 1 SD) heart rate, core body temperature, and respiratory
rate after premedication in the home cage, in the induction chamber, and during 50-min
sevoflurane anesthesia while mice breathed spontaneously and lay in dorsal recumbency on
the warming mat. Dashed lines indicate mean baseline values (measured before anesthesia)
at the same time of day in conscious mice. The baseline respiratory rate was established by
counting the movement of the thorax wall in resting mice before anesthesia.
350
400
450
500
550
600
650
700
750
800
35
36
37
38
39
0
20
40
60
80
100
120
140
160
180
-5 0 5 10 15 20 25 30 35 40 45 50
duration of anaesthesia [min]
resp
irato
ry r
ate
[bre
ath
s
per
min
ute
]co
re b
od
y t
em
pera
ture
[°C
]
heart
rate
[beats
per
min
ute
]
induction c
ham
ber
maintenance with nose mask (spontaneous breathing)
in dorsal recumbency on the warmed mathom
e c
age
FMS fentanyl-midazolam-sevoflurane
KS S-ketamine-sevoflurane
S sevoflurane
normal
Chapter 4: Balanced Anesthesia
78
Figure 4.4) Mean (n = 8 mice; bar, 1 SD) acid–base balance (pH), pCO2, and pO2 in arterial
blood after 10, 30, and 50 min of sevoflurane anesthesia. Dotted lines indicate baseline
levels established from HanIbm:NMRI mice in a previous study.1 Dashed lines indicate
published values from conscious C57BL/6J mice [24].
0
10
20
30
40
50
60
70
80
90
10 min 30 min 50 min
duration of anesthesia
pC
O2
(mm
Hg
)7
7.05
7.1
7.15
7.2
7.25
7.3
7.35
7.4
7.45
10 min 30 min 50 min
duration of anesthesia
pH
0
10
20
30
40
50
60
70
80
90
100
10 min 30 min 50 min
duration of anesthesia
pO
2(m
m H
g)
110
FMS fentanyl-midazolam-sevoflurane
KS S-ketamine-sevoflurane
S sevoflurane
Chapter 4: Balanced Anesthesia
79
Figure 4.5) Mean (n = 8 mice; bar, 1 SD) postanesthetic measurements of the effects of 3
anesthesia protocols on heart rate and core body temperature. Delta (Δ) values represent
deviations from baseline values (established prior to anesthesia) during the corresponding
12-h day and night periods. *, P ≤ 0.05 compared with baseline values and between
protocols.
Discussion
All 3 protocols tested provided a reliable 50-min period of anesthesia in laboratory mice, with
short induction and recovery phases and lack of motor response to noxious stimuli.
Subcutaneous injection of fentanyl–midazolam prior to sevoflurane inhalant anesthesia
induced a gas-saving effect and had the advantage of inducing immediate sedation and
day 2day 1 day 3
-40
-20
0
20
40
60
80
100
120
140
160
-1
-0.5
0
0.5
1
1.5
2
h
ea
rt r
ate
(beats
per
min
ute
)
c
ore
bo
dy t
em
pe
ratu
re
(°C
)
FMS fentanyl-midazolam-sevoflurane
KS S-ketamine-sevoflurane
S sevoflurane
*
*
*
*
*
*
*
*180
Chapter 4: Balanced Anesthesia
80
preventing aversive reactions as well as extensive movements at the time of induction with
sevoflurane. Injection of S-ketamine, the S(+)-enantiomer of ketamine, initially induced
behavioral aberrations suggestive of excitation but attenuated aversive behaviors when mice
were exposed to sevoflurane. In contrast, when sevoflurane anesthesia was induced without
premedication, mice responded with defecation, urination, and locomotion including jumping
and abnormal stationary movements. Compared with sevoflurane monoanesthesia, both
premedication regimens shortened the time required to reach immobilization after exposure
to sevoflurane; this effect was most pronounced with fentanyl– midazolam.
During anesthesia, while mice were warmed by a water-filled mat, core body temperature
and heart rate were depressed compared with baseline values obtained at the time of day
but before anesthesia. Both premedications intensified these effects, but all values during all
3 protocols remained within the ranges considered to be normal for mice. The most important
adverse side effect that occurred during anesthesia was marked respiratory depression, as
indicated by respiratory rates that were far below those of normal resting mice. This
respiratory depression was accompanied by pronounced hypoxia, hypercapnia, and acidosis,
all of which increased with time during anesthesia. Such changes in acid–base balance and
blood gasses are well-known side effects of inhalant as well as injectable anesthesia [26, 77,
122]. The degree of respiratory depression was nearly equal among all protocols, but apnea,
tachypnea, and cardiac arrhythmia occurred with KS anesthesia, and 2 mice in this group
died.
During the first 12 h after anesthesia, heart rate increased in all protocols; this increase was
most pronounced during sevoflurane monoanesthesia and least apparent during the FMS
protocol. Core body temperature was increased at 12 h after sevoflurane monoanesthesia
and to a lesser extent after KS anesthesia. Because locomotor activity was unchanged after
anesthesia regardless of protocol, physical activity is unlikely to be the reason for these
effects. Postanesthetic measurements, including monitoring of body weight, indicated that all
3 protocols had only a short-term effect on the physiology and general condition of the mice.
The minimal alveolar concentration of sevoflurane was determined according to standard
principles [121], including generally accepted adaptations for the particular species-specific
conditions of mice [87-89]. These modifications mainly concern the fact that constant-rate
infusions and mechanical ventilation are not performed during determination of minimal
alveolar concentration in mice. Furthermore, measurements of minimal alveolar
Chapter 4: Balanced Anesthesia
81
concentration in mice were based on the inspired concentration of the inhalant, instead of on
the end-tidal value, as is typical for larger animal species. In addition to the common single
noxious stimulus induced by pinching the tail of the mouse [119], we applied 2 other noxious
stimuli. The hind limb withdrawal reflex has been shown to be useful for estimating depth of
anesthesia in mice [123]. Because applying a clamp between the toes was described as
useful during the determination of minimal alveolar concentration of isoflurane in mice [120],
we incorporated this stimulus in the form of pinching the interdigital webbing of the paw
(pedal withdrawal reflex) in a reproducible manner. As a third noxious stimulus, the
abdominal skin pinch reflex was applied as described earlier [68]. For determination of the
minimal alveolar concentration of sevoflurane, we applied these 3 noxious stimuli only once
at a predefined time point of inhalant anesthesia to standardize the experimental conditions
in regard to sevoflurane concentration and the single injection of fentanyl–midazolam or S-
ketamine, with a view to determining the pharmacokinetics of the injected agents. Therefore,
minimal alveolar concentration was determined at 12 min of sevoflurane anesthesia, which is
congruent with 17 to 19 min after subcutaneous injection of the premedication.
The minimal alveolar concentration determined for sevoflurane monoanesthesia (3.3%) for
the female C57BL/6J mice we tested here was similar to values in from the literature [26,
124]. Analgesic substances are known to reduce the minimal alveolar concentration during
inhalant anesthesia in many animal species [125, 126]. In humans, both fentanyl and
midazolam induce a gas-saving effect when combined with volatile anesthetics [127-129]. In
the current study, applying 0.04 mg/kg fentanyl in combination with 4 mg/kg midazolam as a
subcutaneous bolus injection prior to anesthesia reduced the requirement for sevoflurane
gas by one third. A similar gas-saving effect with isoflurane has been described for ketamine
in dogs [130], but combination of S-ketamine with sevoflurane did not have this effect in our
mice. The most probable explanation for this lack is that we could not administer S-ketamine
as a target-controlled intravenous infusion (as is possible in large animals and humans) but
rather as a single subcutaneous bolus injection. Therefore, from a pharmacokinetic
viewpoint, the effects of S-ketamine might already have been decreasing when we
determined the minimal alveolar concentration (that is, at 17 to 19 min after subcutaneous
injection of 30 mg/kg S-ketamine) [33].
After standardization of the dosages by establishing minimal alveolar concentrations, we
then compared the 3 protocols at dosages of sevoflurane representing 1.5 times the minimal
Chapter 4: Balanced Anesthesia
82
alveolar concentrations. At this dosage, it is generally postulated that 99.9% of animals will
not react to noxious stimuli [92, 131], that is, that the animals have reached surgical
tolerance. However, because we did not confirm analgesia by, for example, measuring heart
rate, blood pressure, or respiration in response to a substantially noxious stimulus (for
example, skin incision), we cannot claim definitively that surgical tolerance was achieved in
the current study. However, motor reflex responses to noxious stimuli were suppressed in all
mice for the entire duration of anesthesia (that is, 50 min).
Shortly (within approximately 2 min) after injection with fentanyl–midazolam, all mice
exhibited reduced physical activity and a sleep-like posture, likely due to the sedative effect
of these agents. In contrast, injection of S-ketamine gave rise to muscle tremors and ataxia.
The spike (up to 800 bpm) in heart rate that we noted in the early phase of induction during
sevoflurane monoanesthesia may be a normal reaction to removal of the mouse from its
cage and placing it in a foreign environment (that is, induction chamber). The markedly lower
heart rate during the induction phase of the FMS protocol suggests bradycardia due to
fentanyl but also indicates the potential benefits of sedation, through stress reduction, during
the initial phase of anesthesia.
During the 50-min anesthesia, mice anesthetized with FMS and KS displayed lower heart
rate and core body temperature than did those anesthetized with S alone. This result can be
explained by the known influences of fentanyl and ketamine on thermoregulation [117, 132].
In addition, the typical cardiovascular effects of the opioid might have potentiated the well-
known cardiopulmonary depression caused by the volatile anesthetic sevoflurane. However,
all heart rate and core body temperature measurements remained within the normal
physiologic ranges of mice in all 3 protocols tested. Given that many other anesthetic
regimens can decrease in core body temperature by more than 4 °C in just a few minutes,
we consider the changes in the current study to be acceptable [64, 68]. Nevertheless, these
findings underline the necessity for thermal support (as supplied in our experiments) during
anesthetic procedures in small laboratory rodents.
In all protocols, the respiratory rate declined far below baseline values in resting mice. Blood
gasses and pH in the arterial blood were impaired by all protocols to a similar extent and to
values clearly different from published reference values from conscious HanIbm:NMRI and
C57BL/6J mice [68, 133]. Therefore, respiratory depression as evidenced by the marked
decrease of respiratory rate in the presence of acidosis, hypoxia, and hypercapnia was the
Chapter 4: Balanced Anesthesia
83
most prominent side effect observed and it was present in all of the anesthetic protocols we
tested. These symptoms may mainly reflect the effect of sevoflurane on cardiopulmonary
function but also may be potentiated by fentanyl and - to a lesser extent - through S-
ketamine. Although surgical stimulation can restore ventilation toward a less deleterious level
[134, 135], surgery was not performed in our experiments. Therefore, lack of constant
surgical stimulation may have exacerbated the respiratory depression associated with
duration of anesthesia. The oxygen content of the air used as a carrier gas likely is
insufficient to prevent hypoxia. Therefore, increasing the inspiratory oxygen fraction (FiO2)
above 0.3 by mixing oxygen into the carrier gas would be useful to minimize hypoxia due to
anesthetic-induced respiratory depression.
The time required for recovery (that is, resposture and motion) after anesthesia was similar
for all groups (1 to 2 min). This fact should be highlighted as a distinct advantage over most
injectable anesthetic regimens in mice [68]. The avoidance of postoperative complications
associated with a prolonged recovery period and the resulting inability to correct physiologic
impairment (for example, hypoxia, hypothermia, hypoglycemia, dehydration) is a key
challenge in developing novel anesthesia methods for small laboratory rodents.
Heart rate and core body temperature increased in all 3 protocols during the first 12 h after
the 50-min anesthesia, but these changes were attenuated or absent in mice that received
FMS anesthesia. Therefore, the influence of our tested anesthesia protocols on physiology
seemed to be of only short-term duration. It should also be noted that, during the 3 d after
anesthesia, none of the anesthetic protocols showed an adverse effect on body weight, thus
suggesting their negligible effects on the animals’ general condition.
In the current study, we tried to apply the most useful and efficient substances and dosages
of injectable drugs and volatile anesthetic available for the adult female C57BL/6J mice we
used. Because dosages vary greatly depending on the specific characteristics of the animals
used (for example, strain, age, sex) as well as the ambient conditions of the laboratory, our
protocols should carefully be adapted for use in other circumstances. Depending on the
animals’ anesthetic needs and the severity of the intervention that they will undergo, higher
or lower dosages particularly of the volatile anesthetic sevoflurane may be required. In
addition, we suggest that the use of S-ketamine as a premedication might be improved by
administering it at a lower dose or in combination with a minor tranquilizer. Such optimization
Chapter 4: Balanced Anesthesia
84
could decrease the frequency, duration, and severity of side effects such as apnea,
arrhythmia (including fatal abnormalities), and excessive excitation.
In conclusion, premedication with subcutaneous injection of fentanyl in combination with
midazolam improved standard sevoflurane monoanesthesia of mice in our laboratory setting.
Advantages included a short and quiet induction phase and decreased negative
postanesthetic side effects on heart rate and core body temperature. A gas-saving effect was
evident in the FMS treatment, corroborating the analgesic potential of the opioid component
(fentanyl) in this modular anesthesia protocol.
Although all 3 protocols used here may be useful for anesthesia in mice, the combination of
injection anesthesia with inhalation anesthesia could be superior to the widely used standard
inhalation monoanesthesia, provided that appropriate drugs are combined and dosages are
adapted to the requirements of the specific animals and laboratory. However, the choice of a
specific anesthetic regimen should always be based on careful deliberation, considering
arguments of animal welfare, feasibility, and any potential interference with the research
project for which the anesthesia is required.
Acknowledgments
This work was sponsored by the ECLAM and ESLAV Foundation. The authors would like to
thank Robin Schneider and the staff of the central biological laboratory for support in housing
mice. We thank Professor Kurt Burki for generously providing research facilities and
resources.
Chapter 5: Behavioral Impact of Anesthesia and Minor Surgery
85
Impact of inhalation anesthesia, surgery and
analgesic treatment on home cage behavior in
laboratory mice
Based on Cesarovic N, Rettich A, Arras M, Jirkof P.
Submitted Nov. 2013, in Applied Animal Behaviour Science
Chapter 5: Behavioral Impact of Anesthesia and Minor Surgery
86
Abstract
Anesthesia and analgesia are used frequently in laboratory routine to ensure animal welfare
and good scientific outcomes in experiments that may elicit pain or require immobilization of
the animal. However, there is concern regarding the effect of these procedures on animal
behavior in subsequent experiments. Our study determined the impact of short inhalation
anesthesia (sevoflurane, 15 min, 4.9%) and minor surgery with or without pain treatment
(carprofen, 5 mg/kg, bid) on spontaneous species-specific home cage behaviors in inbred
mice. Analysis of 18-hour continuous video recordings showed clear post-procedural
changes in spontaneous home cage behaviors, with changes of a moderate level after
anesthesia, being marked after surgery. Self-grooming, resting and locomotion were the
most important behaviors for group separation. Analysis of the temporal distribution of
behavioral changes revealed that resting behavior was altered contradictory to its circadian
rhythm as it was decreased in the light phase and increased in the dark phase. Also,
locomotion was decreased in the dark phase at 12 to 18 hours after surgery and anesthesia.
In contrast, self-grooming was increased independently of circadian rhythm, being increased
for up to 18 hours after surgery and anesthesia. Following surgery, there was no significant
difference in duration of behaviors between animals that were treated with carprofen or left
without pain relief. In conclusion, it can be assumed that the changes observed in home cage
behaviors hint at reduced animal well-being. Pain or the efficacy of post-operative pain
treatment, however, could not be discriminated reliably from the impact of the surgical
procedure including inhalation anesthesia by observing animals’ home cage behavior. For
the interpretation of behavioral research data, the distinct impact of anesthesia, surgery, pain
treatment and other experimental procedures have to be considered. Our results highlight
the requirement for knowledge of species-specific circadian rhythms of behaviors as well as
the importance of determining the appropriate time of day for behavioral and welfare
assessment.
Chapter 5: Behavioral Impact of Anesthesia and Minor Surgery
87
Introduction
Laboratory mice are currently the most widely used animal species in biomedical research.
Due to their manageable size, a wealth of inbred or genetically modified strains and plenitude
of established experimental protocols, mice are used increasingly in complex investigations.
These often require induction of general anesthesia for performing special diagnostic
manipulations (e.g., imaging procedures, endoscopy, blood collection), or surgical
procedures that in turn require peri- and/or post-operative pain treatment. Analgesic
treatment would seem necessary after invasive procedures like laparotomy, but has been
omitted frequently in the past [5, 40]. Reasons may vary from concern that analgesic use
may compromise the data obtained from the proven model to the difficulties of detecting and
interpreting signs of pain after minor surgery in mice (e.g. [40]).
Recently, it has become apparent that the physiological and behavioral changes induced by
minor or moderate surgery can last up to 24 - 48 hours [63, 136]. Moreover, it has been
shown that changes induced by anesthesia, and possibly also by treatment-related
procedures (e.g. handling, transport to operating theatre etc.), may affect physiology and
animal well-being for several hours [26, 137]. Thus it can be assumed that, in some
situations, the effects of anesthesia may overlap and to some extent mask the post-operative
effects of pain. In addition, although the impact of volatile anesthetic agents on learning,
memory, solving of spatial tasks and activity has been studied recently [138-140], the effects
of anesthesia, as an integral part of standard surgical procedures, on spontaneous home
cage behaviors have been described only rarely [141]. Because anesthesia, in particular
inhalation anesthesia, is used increasingly in the laboratory routine of biomedical research,
questions regarding the duration and persistence of long-lasting anesthetic or procedural
effects come into focus [5].
There is concern regarding not only animal welfare but also the reliability of data obtained
from research using animals that have undergone procedures that may elicit pain and/or
involve analgesic and/or anesthetic treatment. This study aimed to determine the effects of
minor surgery with or without pain treatment, as well as the impact of standard, short
inhalation anesthesia alone on spontaneous and species-specific home cage behaviors in
two common inbred mice strains. To this end, the overall temporal distribution of the animals’
Chapter 5: Behavioral Impact of Anesthesia and Minor Surgery
88
natural behaviors was investigated according to their circadian rhythmicity in order to identify
whether specific behaviors are altered significantly after surgery or inhalation anesthesia.
Materials and Methods
Ethics statement
The animal housing and experimental protocols were approved by the Cantonal Veterinary
Department, Zurich, Switzerland, under license no. ZH 120/2008, and were in accordance
with Swiss Animal Protection Law. Housing and experimental procedures also conform to
European Directive 2010/63/EU of the European Parliament and of the Council on the
Protection of Animals used for Scientific Purposes and to the Guide for the Care and Use of
Laboratory Animals (Institute of Laboratory Animal Resources, National Research Council,
National Academy of Sciences, 2011).
Animals
A total of 64 C57BL/6J and DBA/2J mice of both sexes were obtained from our in-house
breeding facility at the age of 6–8 weeks. The health status of the animals was monitored by
a health surveillance program according to FELASA guidelines throughout the experiments.
The mice were free of all viral, bacterial, and parasitic pathogens listed in FELASA
recommendations [86], except for Helicobacter species.
All animals were housed in groups of three to eight animals of the same sex for at least 3
weeks prior to testing in our animal room. Animals were kept in type 3 clear-transparent
plastic cages (425 mm × 266 mm × 155 mm) with autoclaved dust-free sawdust bedding and
two nestlets™ (each 5 cm × 5 cm) consisting of cotton fibers (Indulab AG, Gams,
Switzerland) as nesting material. Additionally, animals were provided with a transparent
plastic shelter (Mouse house™, Indulab, Gams, Switzerland). They were fed a pelleted and
extruded mouse diet (Kliba No. 3436, Provimi Kliba, Kaiseraugst, Switzerland) ad libitum
(provided in the food hopper continuously throughout the entire duration of the experiment)
and had unrestricted access to sterilized drinking water. The light/dark cycle in the room
consisted of 12/12 h with artificial light (approximately 40 Lux in the cage). The temperature
Chapter 5: Behavioral Impact of Anesthesia and Minor Surgery
89
was 21 ± 1°C, with a relative humidity of 55 ± 10%, and the air pressure was controlled at 50
Pa with 15 complete changes of filtered air per hour (HEPA H 14 filter). The animal room was
insulated to prevent electronic and other noise. Disturbances, e.g., visitors or unrelated
experimental procedures, were not allowed.
Experimental groups
In order to distinguish between the effects of inhalation anesthesia and surgery with or
without analgesic treatment, 64 animals (4 per sex and strain) were allocated randomly to
one of four treatment groups: 1) the “anesthesia” group (A), which received inhalation
anesthesia only; 2) the “surgery + anesthesia + analgesia” group (S+), which underwent
inhalation anesthesia and minor surgery with analgesic treatment; 3) the “surgery +
anesthesia” group (S-), which underwent anesthesia and minor surgery without analgesic
treatment; and 4) a control group, which received no treatment (C).
Experimental treatments and data recording
For acclimatization, animals were housed individually for 3 days as described in detail above.
The experimental treatment began 2 hours before light phase with a subcutaneous injection
of 2 μl/g body weight of phosphate buffered saline (PBS) for groups S and A. In the S+
group, 5 mg/kg body weight of the non-steroidal anti-inflammatory drug (NSAID) carprofen
(Rimadyl™, Pfizer Inc., NY, USA) was diluted in PBS and injected as 2 μl/g body weight.
Forty-five minutes later, the animals were transferred in individual transport cages to the
operating theatre, which was located nearby. Mice were anesthetized with sevoflurane
(Sevorane™, Abbott, Baar, Switzerland) as a mono-anesthesia. The anesthetic gas was
provided with a rodent inhalation anesthesia apparatus (Provet, Lyssach, Switzerland);
oxygen (100%) was used as carrier gas. After induction of anesthesia in a Perspex induction
chamber (8% sevoflurane, 600 ml/min gas flow) for 2 minutes, animals were transferred to a
warming mat (Gaymar, TP500, Orchard Park, NY, USA) set at 39°± 1°C to ensure constant
body temperature, and anesthesia was maintained via a nose mask (4.9% sevoflurane, 600
ml/min oxygen flow). The fur was clipped and the operating field disinfected with ethanol in
all animals. Male and female mice of both surgery groups underwent a one-side sham
vasectomy or a one-side sham embryo transfer, respectively. The incision in the abdominal
muscle wall was closed with absorbable sutures (Vicryl™, 6/0 polyglactin 910, Ethicon Ltd,
Chapter 5: Behavioral Impact of Anesthesia and Minor Surgery
90
Norderstedt, Germany) and the skin was closed using skin staples (Precise™, 3M Health
Care, St Paul, MN, USA). Surgery was completed within 6–8 min in both surgery groups.
Anesthesia lasted 14–16 min in all 3 treatment groups. Animals were allowed to recover for
15–20 min on the warming mat before being transferred back to the animal room for
subsequent video recording. All experimental and control recordings began at the start of the
light phase shortly after returning the mouse from its transport cage to its home cage.
Behavioral analysis
Behavior was recorded digitally in the absence of a human observer with infrared sensitive
cameras. The recorded material (18 hours of continuous footage) was subsequently
analyzed by trained and trial-blinded personnel using ObserverXT™ software (Noldus,
Wageningen, Netherlands). The duration of locomotion, self-grooming, resting, eating,
drinking and nest building behavior was measured (Table 5.1, [142]).
Data were initially summed for the whole 18-hour period. In order to determine the temporal
distribution of behavioral changes, the 18 hours were divided into three consecutive 6-hour
periods according to the light-dark cycle in the animal room. Data were summed and
analyzed for the following time frames: 0–6 hours (light phase), 6–12 hours (light phase), and
12–18 hours (dark phase).
Table 5.1) Ethogram of home cage behaviors according to Van Oortmerssen [142].
Statistical analysis
Statistical analyses were performed with SPSS 20.0 software (IBM, Armonk, USA). All data
were tested for normal distribution and homogeneity of variance. If necessary, data were log
(X+1) transformed to meet the assumptions of statistical tests.
home cage behaviours
restingmotionless state, no activity (sitting or lying flat, sometimes with the eyes closed or
nearly closed, includes sleeping)
locomotion oriented movement including walking, running, jumping and grid climbing
self groomingwiping, licking and nibbling the fur with forepaws and tongue, but also scratching and claw cleaning
eating consumption of food
drinking consumption of water from the water bottle e
nest building carrying and shredding of the nestlet, arrangement of cotton fibres, creation of a nest
Chapter 5: Behavioral Impact of Anesthesia and Minor Surgery
91
No significant effect of animal gender was detected with any of the measures. Therefore, a
combined data set of males and females was used. Mean and standard deviation (SD) of
total durations of home cage behaviors were calculated.
Discriminant analysis was used to determine the effects of surgery, anesthesia and analgesic
treatment on home cage behavior; behaviors mainly responsible for group separation were
determined. The total durations of determined behaviors were further analyzed using a
univariate general linear model (GLM) with experimental group as a fixed factor. Post hoc
tests (Bonferroni) were used for comparisons between experimental groups. Significance for
all statistical tests was established at p ≤ 0.05.
Results
Contribution to group separation was analyzed with discriminant analysis of the summed
data, initially for the whole 18 hours observation period and subsequently for the three 6-hour
periods; 0–6h; 6–12h; 12–18h (Figure 5.1). Behaviors that contributed most to group
separations were locomotion, self-grooming and resting. These behaviors also represented
the largest part of the overall behavioral time-budgets, by far. Based on this observation, only
the results for locomotion, self-grooming and resting behavior were analyzed with a GLM and
presented in further detail (Figure 5.2).
Chapter 5: Behavioral Impact of Anesthesia and Minor Surgery
92
Figure 5.1) Scatter plot of discriminant scores assigned to individual mice. The significance
of each function in separating groups, and their percentage contribution to between-group
variance are shown on each axis. A) 18h observation period. Duration of self-grooming and
locomotion behavior contributed most to group separation. B) During the first observation
sequence (0-6h post treatment) locomotion and self-grooming behavior were mainly
responsible for group separation. C) During 6–12h post treatment, duration of self-grooming
behavior was mainly responsible for group separation. D) Locomotion, self-grooming and
resting contributed significantly to group separation during the 12–18h post-treatment
observation period.
Function 1 (p ≤ 0.0001, 93%)
Fun
ctio
n2
(n
on
-sig
., 5
%)
Function 1 (p ≤ 0.0001, 87%)
Fun
ctio
n2
(n
on
-sig
., 1
0%
)
4
2
0
- 2
- 4
- 4 - 2 0 2 4
- 4 - 2 0 2 4
4
2
0
- 2
- 4
A
Fun
ctio
n2
(n
on
-sig
., 9
%)
Function 1 (p ≤ 0.0001, 89%)
- 4 - 2 02
4
4
2
0
- 2
- 4
B
Fun
ctio
n2
(n
on
-sig
., 2
6%
)
4
2
0
- 2
- 4
- 4 - 2 0 2 4
C D
Surgery without analgesiaSurgery with analgesiaAnesthesiaControl
Function 1 (p= 0.0012, 78%)
total 18 hours 0 – 6 hours
6 – 12 hours 12 – 18hours
Chapter 5: Behavioral Impact of Anesthesia and Minor Surgery
93
Effects of treatment on analyzed behaviors: total 18-hour
Observations
During the total 18-hour observation period, the control group (C) (230 min +/- 60) and the
anesthesia group (A) (197 min +/- 60) displayed significantly (p ≤ 0.0001, each comparison)
longer durations of locomotion compared to animals that underwent surgery with (S+) (112
min +/- 63) and without (S-) (89 min +/- 30) pain treatment (Figure 5.2 A 1).
Self-grooming behavior was prolonged significantly after all experimental procedures
compared with the untreated C group (199 min +/- 37) ranging from a high level in group S
(404 min +/- 71; p ≤ 0.0001) to an intermediate level in group S+ (351 min +/- 62; p ≤
0.0001),with the shortest durations in group A (278 min +/- 54; p = 0.002). Additionally,
significant differences between the anesthesia and surgery groups were seen: S- (p ≤
0.0001) and S+(p = 0.004) (Figure 5.2 A 2).
No significant differences in resting durations were observed between any treatments: C
(568 min +/- 83), S- (523 min +/- 71), S+ (539 min +/- 71) and A (507 min +/- 67) (Figure 2A
3).
Effects of treatment on analyzed behaviors: 6-hour observations
By dividing the observations into 6-hour sequences, circadian differences in the effects
became apparent (Figure 5.2 B 1-3). During the first 6h observation period (0–6h),
locomotion durations did not show any significant differences between groups S- (26.7 min
+/- 18.7) S+ (26.8 min +/- 19.1), A (51.7 min +/- 48.1) and C (47.9 min +/- 19.9). Between 6
and 12 hours post treatment, groups S- (15.9 min +/- 8.4), S+ (11.8 min +/- 7.2) and A (20.9
min +/- 10.5) displayed no significant differences when compared to group C (17.2 min +/-
9.7), while locomotion duration of S+ was significantly shorter than that of group A (p =
0.036). In the last observation period (12–18h), corresponding to the first 6 hours of dark
phase, all groups showed significantly shorter durations of locomotion compared to the
untreated group C (185.7 min +/- 52): S- group (46.7 min +/- 28 ; p ≤ 0.0001), S+ (73 min +/-
63; p ≤ 0.0001) and A (121.6 min +/- 57; p = 0.006). Further, duration of locomotion was
significantly shorter (p = 0.001) in group S- than in group A. Total duration of self-grooming
during the first 6h observation period (0–6h) was significantly longer in experimental groups
Chapter 5: Behavioral Impact of Anesthesia and Minor Surgery
94
S- (162 min +/- 33; p ≤ 0.0001), S+ (141 min +/- 45; p ≤ 0.0001) and A (111 min +/- 43; p ≤
0.0001) compared to the control group (50 min +/- 20). Additionally, there was a significant
difference between groups S- and A (p = 0.001). A comparable tendency was seen in the
second time period (6–12h). Animals that underwent surgery groomed themselves for
significantly longer in groups S- (106.4 min +/- 43; p = 0.004) and S+ (96.3 min +/- 33; p =
0.042), compared to group C (62 min +/- 22), while in group A self-grooming was prolonged
only insignificantly (79.4 min +/- 34, n.s.). In the last observation period (12–18h), animals
that received surgery without pain treatment (S-) spent the most time grooming (136.2 min
+/- 33; p ≤ 0.0001) followed by animals that received surgery with pain treatment S+ (113.3
min +/- 27; p = 0.014). Compared to the untreated group C, differences were significant (80
min +/- 25).
Animals that received anesthesia only (A) (91.3 min +/- 30) showed significantly shorter
grooming durations compared to group S- (p ≤ 0.0001). Animals that underwent anesthesia
or surgery spent significantly less time resting in the first observation period (0–6h) compared
to the untreated group C (224 min +/- 44): S- (154 min +/- 47; p = 0.002), S+ (165 min +/- 61;
p = 0.013) and group A (160 min +/- 55; p = 0.006).In the second observation period (6–12h),
resting duration was significantly shorter in group S- (220.9 min +/- 49; p = 0.011) compared
to the untreated group C (267.8 min +/- 27), with somewhat shorter resting duration in groups
S+ (237.4 min +/- 37) and A (239.7 min +/- 45). Animals that underwent surgery and
anesthesia spent significantly more time resting compared to untreated controls (60.1 min +/-
35) in the last observation period (12–18h): S- (148.1 min +/- 49; p ≤ 0.0001), S+ (137 min
+/- 59; p ≤ 0.0001) and A (108.7 min +/- 50; p = 0.045).
Chapter 5: Behavioral Impact of Anesthesia and Minor Surgery
95
Figure 5.2) Effects of anesthesia and surgery with or without analgesic treatment on duration
of 3 spontaneous home-cage behaviors compared to control values. * Significant (p ≤ 0.05)
differences between experimental groups. A) Total duration of locomotion (A1) was
decreased post experiment, self-grooming (A2) increased, and resting (A3) behavior
100
300
200
0
100
200
300
400
500
600
self
gro
om
ing
du
rati
on
[min
]
0
50
100
150
200
250
0
surgerywithout
analgesia
0
200
600
400
800
rest
ing
du
rati
on
[min
]
100
200
400
300
0
0 – 6h 6 – 12h 12 – 18h
time after treatment
200
300
400
loco
mo
tio
nd
ura
tio
n[m
in]
100
0
surgerywith
analgesia
anaesthesia control
A1
A2
A3
B1
B2
B3
Surgery without analgesiaSurgery with analgesiaAnesthesiaControl
*
*
*
*
*
*
**
*
*
**
**
**
** *
**
*
*
**
*
* **
*
Chapter 5: Behavioral Impact of Anesthesia and Minor Surgery
96
remained unchanged when the whole 18h observation period was analyzed. B) Temporal
distribution of behavioral effects became apparent when dividing observations into 3
consecutive 6-hour sequences. Duration of locomotion behavior (B1) was unchanged during
the lights-on period but shortened dramatically during the dark period. The increase in
duration of self-grooming (B2) was distributed equally in all time sequences, whereas resting
(B3) was decreased during the first (0–6h) and increased during the last (12–18h) sequence.
Discussion
To assess the impact of inhalation anesthesia and surgery with or without pain treatment in
mice, we used non-invasive behavioral observations that can be applied in the animals’
home cage without disturbing the animal or provoking additional stress. Using this system we
were able to analyze each animal’s behavior continuously for 18 hours following
experimental treatments. In contrast to most physiological and clinical parameters, behavior
can be observed easily in a non-invasive manner and can provide a sensitive correlate of the
internal state of an animal. Alterations in the frequency of or in the latency to display,
spontaneous and species-specific behaviors (e.g., rearing, sniffing, walking or burrowing
behavior) [143, 144], as well as the quality of nest construction and structuring of cage
territory, [63, 137] are recent examples of such behavioral indicators.
The results of this study showed that minor surgery with short inhalation anesthesia, either
with or without pain treatment, induced alterations in spontaneous home-cage behaviors
such as self-grooming, resting and locomotion. These changes persisted for up to 18 hours.
When interpreting the data summed over the whole 18-hour observation period, we found
that locomotion and self-grooming behaviors were most affected by the experimental
procedures. After surgery, animals displayed a marked decrease in locomotion and a strong
increase in self-grooming. Self-grooming showed a clear stepwise increase from baseline
over anesthesia only, to surgery with pain treatment, to surgery without pain treatment.
Differences in self-grooming between groups were significant except for the difference
between surgery with pain treatment and surgery without pain treatment. In contrast to
locomotion and self-grooming, there was no effect on the overall duration of resting behavior
if the 18-hour post treatment period was observed as a whole.
Chapter 5: Behavioral Impact of Anesthesia and Minor Surgery
97
Pain treatment with carprofen had no statistically significant effect on alterations of
spontaneous home cage behaviors. However, animals that received pain treatment during
surgery readily assumed intermediate levels of locomotion and self-grooming compared to
the group in which pain treatment was not administered during surgery and the group that
underwent anesthesia only. Therefore, it could be speculated that some, but not complete,
amelioration of post-operative pain is achieved by administering the non-steroidal anti-
inflammatory drug (NSAID) carprofen at a dosage of 5 mg/kg body weight. However,
previous studies using physiological investigations and behavioral testing demonstrated that
carprofen provided sufficient relief from post-operative pain [63, 144]. Thus, it might be that
the alterations of spontaneous home cage behaviors analyzed in this study are not ideal
parameters for estimating pain alleviation by NSAIDs.
When dividing the observation period into 3 consecutive 6-hour-long time segments
according to the light cycle in the animal room, circadian-dependent display patterns could
be observed.
For the first 12 hours after treatment - corresponding to the complete light phase in the
animal room - all animals displayed general low levels of locomotor activity, and no
difference could be observed between treated and untreated groups regarding this behavior.
However, in post-treatment hours 12–18 (first 6 hours of the dark phase) locomotion of both
surgery groups was reduced by 75% and that of the anesthesia group by 35% compared to
that of the control group. In contrast, the duration of self-grooming behavior was influenced
strongly by treatment in all 3 time segments, showing a marked increase in treated animals
as compared to the untreated control group. Remarkably, the effects on duration of self-
grooming seemed not to be influenced by the light-dark cycle in the animal room.
Furthermore, the duration of resting behavior, which displayed no differences between
groups in the summed 18-hour analysis period, showed clear and gradual treatment-related
effects when analyzed according to the time progression, i.e. in separate 6-hour periods.
Effects were most significant during the first 6 hours after treatment, when resting decreased,
and at 12–18 hours after treatment, when it increased markedly in comparison to control
animals.
In recent years, volatile anesthetics (e.g., sevoflurane, isoflurane) have been used
increasingly in laboratory animal practice due to their safety and association with rapid
recovery. Further, inhalation anesthesia is used in diverse procedures, including
Chapter 5: Behavioral Impact of Anesthesia and Minor Surgery
98
neurobiology research in which animals are studied subsequently in behavioral tests and
where their performance may be influenced by the persistent effects of anesthetic drugs
[138].
Previous studies have demonstrated that inhalation anesthesia can induce changes in heart
rate, core body temperature and fecal corticosterone levels that last for several hours after
treatment [26, 145]. In this study, we have shown that non-painful, short (15 minutes)
sevoflurane inhalation anesthesia can cause post-anesthetic alterations in locomotion, self-
grooming and resting behaviors that are noticeable for up to 18h after treatment. Therefore,
for accurate interpretation of behavioral research data the distinct individual impacts of
anesthesia, surgery, pain treatment and other experimental procedures have to be
considered. However, as our study protocol did not determine the extent of effects caused by
treatment-related actions (like transport, handling etc.), this question requires further
investigation.
Studying behavior in sufficient detail to detect post-surgery-related changes in spontaneous,
home-cage based behavior patterns, and the effects of drugs upon such behaviors, is quite
tedious and time consuming, thus studies are often confined to analyzing only a limited range
of behaviors or performing assessments only over a very short time frame [143]. Determining
an optimal observation time-point is one of the major difficulties in behavioral research. For
example, the display of signs of pain as well as pain tolerance itself is dependent on
circadian rhythm, and thus the need to determine the appropriate time of day for
observations [136, 146, 147] renders post-operative pain assessment even more challenging
in mice. In this study, we were able to show that all 3 behaviors studied in detail (locomotion,
self-grooming and resting) displayed different temporal effect patterns. Whereas the effects
on self-grooming were distributed evenly over the whole period analyzed, locomotion was
changed only during ‘dark’ and resting first decreased and then increased markedly in
treated groups. We believe that these data suggest strongly that the effects on spontaneous,
home-cage based behaviors caused by anesthesia and minor surgery are not displayed
uniformly throughout the day. Our results highlight the requirement for knowledge of species-
specific circadian rhythms of behaviors as well as the importance of determining the
appropriate time of day for behavioral and welfare assessment.
Chapter 5: Behavioral Impact of Anesthesia and Minor Surgery
99
Conclusion
Spontaneous home cage behaviors, locomotion, self-grooming and resting, were altered for
up to 18 hours in all treatment groups, and a graduation between untreated control,
anesthesia and surgery groups was found. Short inhalation anesthesia induced moderate
changes whereas the impact of surgery was considerable. Thus it can be assumed that the
observed changes in home cage behaviors hint at reduced animal well-being. Pain therapy
only partially ameliorated the aforementioned effects, leading to the conclusion that either the
chosen dosage was too low or that alterations in the spontaneous home cage behaviors
analyzed in this study do not allow NSAID efficiency to be estimated reliably.
While self-grooming behavior changed post experimentally independently of circadian
rhythm, changes in locomotion and resting behavior were distinctly affected by the time of
day.
In conclusion, for proper interpretation of behavioral research data, the distinct impacts of
anesthesia, surgery, pain treatment and other experimental procedures have to be
considered. Our results highlight the requirement for knowledge of species-specific circadian
rhythms of behaviors as well as the importance of determining the appropriate time of day for
behavioral and welfare assessment.
Acknowledgements
This work was sponsored by grants from the Federal Veterinary Office (Bern, Switzerland),
and UBS foundations. The authors would like to thank Robin Schneider, Hugo Battaglia and
the staff of the Central Biological Laboratory for support in housing mice. We thank Professor
Kurt Bürki for generously providing research facilities and resources.
Chapter 6: General Discussion
100
General Discussion
Anesthetic procedures are fairly common in laboratory mice. Although hampered by some
problems, such as narrow safety margins and the inability to respond to the needs of
individual animals [68], injection anesthesia (administered intraperitoneally or
subcutaneously) remains the major means of narcosis in laboratory rodents [5]. However,
anesthetic procedures can negatively influence not only animal well-being but also the quality
of the data generated.
In order to fulfill ever-growing legislative demands aimed at reducing distress to a minimum,
novel inhalation and balanced anesthetic protocols need to be developed and evaluated for
use in laboratory mice. To achieve these goals, data on physiological and behavioral
parameters in unrestrained, unstressed animals in their ‘home’ environment need to be
gathered.
Radiotelemetry is a powerful alternative to conventional methods of measurement of
physiological parameters in biomedical research. Radiotelemetry represents the only
technique currently available for data collection from unrestrained, freely moving mice [58].
By using this method, it is now possible to gather data continuously and/or for longer periods
of time from animals residing in their own familiar environment, thus minimizing the stress to
the animals and consequent experimental artifacts [148].
However, data collection by radiotelemetry in mice requires preliminary surgical implantation
of the telemetry transmitter. High levels of morbidity and mortality resulting from the adverse
effects of surgery, inadequate anesthesia or post-operative care have been described as the
major factors hampering the use of radiotelemetry in mice [148, 149]. For that reason we
suggest that experimenters holding basic or even advanced micro-surgical skills perform
initial trials in fresh mouse cadavers to establish these procedures and to become familiar
with the specifics of this kind of surgery. Although acknowledging the difficulties in providing
aseptic conditions in small rodent surgery (e.g., the cooling effect of extensive hair clipping
and disinfection, the impracticality of bandages to protect wounds), we argue that sterility
should be maintained during surgery to keep the bacterial burden and the risk of infection
low. In addition, antibiotic prophylaxis should also be administered during implantation.
Chapter 6: General Discussion
101
Together with opioid - NSAIDs modular analgesic treatment, fluid and energy supportive
treatment should be applied twice daily for at least a few days. Furthermore, all injections
should be given body-warm (37°C) and animals should be placed on heat support during the
recovery period [59]. Although previously not much discussed in the literature, we argue that
fluid and energy supportive therapy together with external heat provision might just give mice
the edge in surviving and recovering from a high impact surgery, especial when fragile
transgenic animals are used [59]. Moreover, a clearly defined monitoring plan plays a crucial
role in ensuring a satisfactory outcome of the experiment and helps to avoid unnecessary
suffering. Animals exhibiting unsatisfactory recovery or prolonged convalescence should be
released from the experiment and sacrificed before reaching a moribund stage. For this
purpose, a data sheet (Appendix 1) facilitating the systematic monitoring of critical symptoms
and providing advice on humane endpoints has been established and is recommended for
use after surgical implantation of radiotelemetry transmitters in laboratory mice.
In order to obtain reliable, reproducible and artifact-free data with telemetric measurements,
it is crucial to exclude environmental influences. In the second chapter of this work we draw
particular attention to the importance of standardized conditions, especially during telemetric
recordings. We recommended that the room in which experiments are performed is isolated
from electronic and acoustic noise. In addition, no disturbances, such as visitors or unrelated
experimental procedures, should be allowed when conducting measurements.
After establishing implantation and telemetry-system-use protocols in our laboratory, we
further aimed to test volatile and balanced anesthetics regimes in an approach and duration
that closely mirrors laboratory routine (e.g. spontaneous respiration, surgical depth, 50
minutes duration), as described in chapters 3 and 4 of this work.
Isoflurane and sevoflurane are currently the most commonly used volatile anesthetics in
human medicine but, despite the fact that modern volatile anesthetics have been shown to
induce and maintain anesthesia of superior quality compared to injection mono-anesthetics
[95], their use in animals is still not widely accepted in laboratory routine [5]. Moreover,
despite several publications describing the efficacy of isoflurane [77], the literature on the use
of sevoflurane or balanced protocols for laboratory mice remains quite scarce.
As MAC values represent a measure of anesthetic potency and are used not only for dose
estimation during application but also as a unit of comparison between substances, their
Chapter 6: General Discussion
102
determination represents a cornerstone of our work. In chapter 3, we showed that isoflurane
and sevoflurane quickly induced loss of the writhing reflex, and during maintenance with 1.5
MAC provided safe anesthesia with stable cardio-vascular parameters (heart rate, ECG) for
up to 50 minutes at a depth that was sufficient to abolish purposeful movement after noxious
stimulation. However, marked respiratory depression, hypercapnia and subsequent
respiratory acidosis were induced by both volatile anesthetics tested in our study—a major
but not unexpected side-effect. Both isoflurane and sevoflurane are known to reduce
respiratory drive and the ventilatory responsiveness to CO2 challenge [150]. In humans the
reduction of tidal volume (Vt) is partially compensated by an increase in respiratory
frequency [150]. However, in mice volatile anesthetics at clinically relevant concentrations
(1.5 MAC) markedly reduce respiratory rate with a minimal increase in tidal volume, and so
induce an overall decrease in respiratory minute volume [103]. After termination of
anesthesia, all animals recovered quickly but displayed a marked increase in heart rate and
core body temperature during the first 12h after anesthesia. Locomotor activity (ACT as
measured by the telemetry system) was increased only marginally and thus could not explain
the changes in heart rate.
In order to further refine anesthetic regimens in mice, and especially to address the problems
of induction-related stress, respiratory depression and a possible lack of analgesia, a
sedative/analgesic component (either fentanyl-midazolam or S(+)-ketamine) was included in
the previously established inhalation anesthesia protocol. The combination of fentanyl and
midazolam is known to produce deep sedation in humans [32, 151]. Further, both
substances are known to reduce the MAC of volatile narcotics [127, 152-154]. Midazolam
has even been shown to produce anti-nociceptive effects in mice [155] and to influence the
affective component of pain in healthy human volunteers [156] when administered
systemically. On the other hand, ketamine differs from most other drugs used for anesthesia
because it induces ‘dissociative anesthesia’ (a catalepsy-like state) and has a significant
analgesic effect. These properties are possibly due to the action on opioid and NMDA
receptor complex, among other factors [157, 158]. Its wide use in laboratory animal
anesthesia can be also attributed to the fact that its application routes include i.v. as well as
i.m., i.p. and p.o. [3]. In the study described in chapter 4, sevoflurane was chosen as the
volatile component as it is considered less aversive to mice than isoflurane [3].
Subcutaneous injection of fentanyl–midazolam prior to sevoflurane inhalation anesthesia
induced a gas-saving effect, whereas ketamine displayed no such effect. Application of
Chapter 6: General Discussion
103
fentanyl-midazolam had the advantage of inducing immediate sedation and prevented
adverse reactions as well as extensive movement at the time of induction with sevoflurane.
The markedly lower heart rate during the induction phase of the fentanyl–midazolam–
sevoflurane protocol suggests a transient bradycardia due to fentanyl action. It also indicates
a potential benefit of sedation through stress reduction during the initial phase of anesthesia.
Upon evaluation of ECG recordings, cardiac arrhythmias were readily observed with the
S(+)-ketamine-sevoflurane protocol. As S(+)-ketamine is known not to affect cardio-vascular
parameters negatively in humans [151, 159, 160], the underlying cause of the observed
arrhythmias might be explained by the action of ketamine on myocardial ionic currents, which
exert different effects in different species [161]. As no blood-pressure measurements were
performed we cannot conclude if, and to what extent, these events influenced blood pressure
or cardiac output. Further, despite being known not to influence central respiratory drive [34,
151, 159], we observed irregular breathing patterns (episodes of tachypnea followed by
apnea) in all mice during S(+)-ketamine-sevoflurane anesthesia. This side-effect has been
demonstrated to occur if ketamine is highly dosed [162]. We cannot exclude the possibility of
over-dosing in this study. In this regard, we would like to point out that a major problem in our
study was that dose recommendations for ketamine vary greatly [68, 163] and, to our
knowledge, no clear dose recommendation for S(+)-ketamine for mice is available.
In all protocols tested in chapter 4 (sevoflurane mono-anesthesia, fentanyl-midazolam-
sevoflurane and S(+)-ketamine-sevoflurane), the respiratory rate dropped far below baseline
values in resting mice. Hence acidosis, hypoxia, and hypercapnia were again the most
prominent side effects. Although respiratory rate during fentanyl-midazolam-sevoflurane
anesthesia was constantly higher than in other protocols, it improved blood-gas analysis
results only marginally. The reduction in sevoflurane MAC was obviously not sufficient to
overcome the respiratory depressant effects of fentanyl and midazolam [164-166]. Although
known to produce additive respiratory depression [151], we suggest that their positive
sedative and analgesic properties outweigh their effects on respiration in the approach used
in this study.
Arguably, the choice of opioid component could influence respiratory drive greatly during
anesthesia. Alfentanyl and remifentanyl, although commonly used in similar protocols in
human medicine [29], might produce some of the same respiratory effects as fentanyl (µ-
agonists), but would need constant re-injection as they are short acting [29]. Buprenorphine
Chapter 6: General Discussion
104
could be a possible agent to test in this regard. As well as for its slow action and dissociation
from the µ-opioid receptor (providing longer analgesia), it is known to have a leveling-off
effect on respiratory depression, in contrast to fentanyl, which reduces respiratory drive
dose-dependently to the point of apnea in rats and humans [167]. Partial agonism of
buprenorphine at the µ-opioid receptor is generally held responsible for the ceiling
phenomenon [167-169]. As partial agonism indicates a partial (respiratory depressant) effect
despite full µ-receptor occupancy we suggest there is a need for a study to test the safety
and efficacy of buprenorphine for ‘balanced anesthesia’ in laboratory mice.
Although in many aspects superior to the other protocols, fentanyl-midazolam-sevoflurane
anesthesia also induced a heart rate and core body temperature increase for several hours
following termination of anesthesia. Again, these effects could not be explained simply by the
increase in locomotor activity.
Therefore we aimed to shed some light on this unexplained cardio-vascular activity in the
study presented in chapter 5. It is mainly focused on the impact of anesthesia on the natural
species-specific behaviors of mice. Therefore, behavioral alterations during the 18 hours
following anesthesia and minor surgery with or without post-operative pain treatment were
analyzed. In contrast to most physiological and clinical parameters, behavior can be
observed easily in a non-invasive manner and can provide a sensitive correlate to the
internal state of an animal. The results showed that minor surgery with or without pain
treatment induced alterations in spontaneous home-cage behaviors such as self-grooming,
resting and locomotion. A short inhalation anesthesia also induced similar changes, albeit to
a lesser extent. Marked increases in self-grooming and a distorted pattern of resting behavior
that were observed after anesthesia, correlate with the increase in heart rate and core body
temperature demonstrated in previous studies, and could be the underlying cause of heart
rate and body temperature changes. It could be argued that these results indicate a post-
anesthesia stress-coping reaction invoking excessive self-grooming and mild restlessness. In
turn, this might cause the observed increase in heart rate. Further, it is widely accepted that
even slight hypothermia during anesthesia is often followed by post-operative shivering,
which increases heart rate [170].
Remarkable findings were made when the observation period of 18 hours was divided into
three consecutive 6-hour time segments. The segments were aligned with the light cycle in
the animal room. With this sub-analysis, circadian-dependent behavioral display patterns
Chapter 6: General Discussion
105
became apparent. Our study demonstrated that although unchanged for the first 12h, during
post-treatment hours 12–18 (first 6 hours of the dark phase) duration of locomotion behavior
in the anesthesia group was decreased by 35% compared to that of the control group
(untreated animals). Thus, the bulk of locomotion-decreasing effects is actually displayed at
12-18 hours post-treatment, i.e. during the active phase of the circadian rhythm. In contrast,
self-grooming was not affected by the light-dark cycle in the animal room. Self-grooming in
rodents was characterized as displacement or adjunctive behavior [171]. Stressors like
novelty and handling can elicit grooming behavior in rats [172, 173] and has been known to
occur after surgery in mice [37, 174]. While no correlation with indicators of anxiety were
found, it seems that grooming coincides better with the period after arousal and rather
reflects the process of habituation to a stressful situation [171]. Moreover, central
dopaminergic activation, most likely via the D1 receptor, was reported to induce intense
grooming behavior [175-178]. Recently, it has been shown that a D1 receptor-mediated
arousal mechanism likely plays an important role in emergence from general anesthesia
[179].
While it is known that volatile anesthetics can impair cognitive function in laboratory mice
[180], syndromes like post-operative delirium (POD) or post-operative nausea and vomiting
(PONV), which are well described in humans [181, 182], have never been discussed in mice.
Although rodents cannot vomit [183], nausea-indicative behavior (pica) has been described
for rats [184]. As none of the tested anesthetic protocols caused reduced food- or water-
intake or significantly decreased body weight, we assume that such syndromes, if present,
are of mild/moderate grade. The possibility of POD or PONV has not been discussed in the
literature to date, so the question of their existence in laboratory mice remains unanswered.
However, looking into this matter in detail might make a substantial contribution to animal
welfare in the future. In our opinion, these findings taken together suggest that post-
anesthesia and post-surgical recovery periods represent a stressful time for mice. Although
pain-free, after only being anesthetized, animals still feel aroused, possibly even drowsy and
ill. In order to cope with this newly emerging situation, mice will engage in excessive
adjunctive behaviors (like self-grooming) which in turn interrupts their normal resting pattern
and may elevate heart rate.
The demands on newly developed anesthetic approaches in mice for routine laboratory use
are numerous. The small size of mice, together with their inaccessible arteries and veins, a
Chapter 6: General Discussion
106
high body surface / body mass ratio and very limited monitoring possibilities make this small
mammal an anesthetic challenge. As the vast majority of users have only a basic knowledge
of anesthesia, the user-friendliness of such protocols plays a crucial role. Besides providing
sufficient depth of anesthesia, the protocol must not be too elaborate or work intensive, or
demand a high level of skill due to the ever-present possibility of misuse. Despite the
advantages of inhalation anesthesia, such as better control over the depth of anesthesia and
minimal biotransformation of the agents used, injectable anesthetic protocols have
traditionally been preferred in mice, possibly because minimal equipment and training are
required and initial costs are lower [5, 163]. However, several injectable protocols in routine
use (e.g. ketamine-xylazine) have been shown not to produce a plateau of anesthesia
sufficient for surgery in all cases [68, 163]. Apart from this, most injectable protocols are
known to markedly reduce heart rate, blood pressure and body temperature and require
prolonged recovery (immobilization) periods [68, 93, 163]. Moreover, although it is known
that age, sex, strain and even housing conditions can greatly influence susceptibility to
injectable narcotics [73, 185-187], large variations in recommended doses are described in
the literature [3, 68, 163], making the choice of an appropriate injectable protocol even more
difficult. Altogether, these variables and circumstances lead to a narrow safety margin
regarding death rate and surgical tolerance with anesthesia protocols in mice that rely on
injectable agents only. In comparison, age-, sex- and strain-dependent differences in
requirements for volatile anesthetics (MAC) are modest [88] and are not perceived as
hampering since agents are administered in a controllable way, resulting in a broad safety
margin. Moreover, inhalation anesthesia in mice provides high flexibility regarding the
duration and depth of anesthesia and thus could be applied to different needs ranging from
short immobilization to surgical interventions. Further, as demonstrated in this work and in
the literature, volatile narcotics can provide safe surgical anesthesia by eliciting fewer
depressant effects on the cardio-vascular system [74, 93]. Interestingly, rather than features
of the anesthetic or reported side effects, it is often difficulties in operating and handling the
equipment and the initial costs of acquisition that are stated as reasons for the use of
injectable- over inhalation-based anesthetic protocols [188, 189]. This situation further
underlines the need to develop and refine simple and easy-to-use inhalation and inhalation-
based anesthetic regimes for laboratory mice.
To summarize this work, we suggest that radio-telemetry is the best tool currently available
for gathering cardio-vascular data in stress-free animals. Implantation surgery, although quite
Chapter 6: General Discussion
107
invasive, has a high rate of success if surgical, anesthetic, analgesic and supportive
measures are applied correctly. Further, we believe that the results shown in the Chapters 3
and 4 clearly demonstrate that isoflurane and sevoflurane can produce a safe and stable
anesthesia of superior quality as compared to widely used injectable protocols.
Premedication with subcutaneous injection of fentanyl in combination with midazolam
improved standard sevoflurane mono-anesthesia of mice in our laboratory setting.
Advantages included a short and quiet induction phase, physiological respiratory rate and
reduced post-anesthetic side-effects on heart rate possibly due to desirable sedative traits of
the fentanyl-midazolam combination. A gas-saving effect was evident in the fentanyl-
midazolam-sevoflurane treatment, corroborating the analgesic potential of the opioid
component (fentanyl) in this modular anesthesia approach. Changes in cardio-vascular
parameters observed after inhalation anesthesia could be explained by interruption of the
normal circadian rhythm and the effects upon duration of certain spontaneous, home-cage
based behaviors, suggesting activation of coping mechanisms during potentially stressful
post-anesthetic period. Further, we believe that results displayed, highlight the requirement
for a better understanding of species-specific circadian rhythms of behaviors as well as the
importance of determining the appropriate time of day for behavioral assessment of welfare
in laboratory mice.
References
1. European Commission 2010, Sixth Report on the Statistics on the Number of
Animals used for Experimental and other Scientific Purposes in the Member States of the
European Union, SEC(2010) 1107, Brussels
2. Bundesamt für Veterinärwesen 2013, Tierversuchsstatistik 2012, Bern
3. Flecnell, P 2009, ‘Laboratory Animal Anesthesia Third Edition’, Elsevier, London, pp.
300
Chapter 7: References
108
4. Institute for Laboratory Animal Research (ILAR) 2011, ‘Guide For The Care and Use
of Laboratory Animals Eight Edition’, National Academies Press, Washington DC
5. Stokes EL, Flecknell PA, Richardson CA 2009, ‘Reported analgesic and anaesthetic
administration to rodents undergoing experimental surgical procedures’, Laboratory Animals,
vol 43, no. 2, pp. 149-54.
6. Steffey, E 2007, ‘Inhalation Anesthetics’, in Tranquilli WJ, Thurmon JC, Grimm KA
(ed.),’Lumb & Jones Veterinary Anesthesia and Analgesia Fourth Edition’, Blackwell
Publishing, Iowa, pp. 355 - 85.
7. Collins JG, Kendig JJ, Mason P 1995, ‘Anesthetic actions within the spinal cord:
contributions to the state of general anesthesia’, Trends in Neurosciences, vol 18, no.12,
pp.549-53.
8. Rampil IJ, Mason P, Singh H 1993, ‘ Anesthetic potency (MAC) is independent of
forebrain structures in the rat’, Anesthesiology, vol 78, no. 4, pp. 707-12
9. Antognini JF, Schwartz K 1993, ‘Exaggerated anesthetic requirements in the
preferentially anesthetized brain’, Anesthesiology, vol 79, no. 6, pp.1244-9
10. Hamacher J, Arras M, Bootz F, Weiss M, Schramm R, Moehrlen U 2008, ‘Microscopic
wire guide-based orotracheal mouse intubation: description, evaluation and comparison with
transillumination’, Laboratory Animals, vol 42, no. 2, pp.222-30
11. Moehrlen U, Schwoebel F, Reichmann E, Stauffer U, Gitzelmann CA, Hamacher J
2005, ‘Early peritoneal macrophage function after laparoscopic surgery compared with
laparotomy in a mouse model’, Surgical Endoscopy, vol 19, no. 7, pp. 958-63
12. Su X, Looney M, Robriquet L, Fang X, Matthay MA 2004, ‘Direct visual instillation as
a method for efficient delivery of fluid into the distal airspaces of anesthetized mice’,
Experimental Lung Research, vol 30, no. 6, pp. 479-93
13. Singer T, Brand V, Moehrlen U, Fehrenbach H, Purkabiri K, Ott SR 2010, ‘Left-sided
mouse intubation: description and evaluation’, Experimental Lung Research, vol 36, no. 1,
pp. 25-30
Chapter 7: References
109
14. Holaday DA, Fiserova-Bergerova V, Latto IP, Zumbiel MA 1975, ‘Resistance of
isoflurane to biotransformation in man’, Anesthesiology, vol 43, no. 3, pp. 325-32
15. Roth D, Petersen-Felix S, Bak P, Arendt-Nielsen L, Fischer M, Bjerring P 1996,
‘Analgesic effect in humans of subanaesthetic isoflurane concentrations evaluated by evoked
potentials’, British Journal of Anaesthesia, vol 76, no. 1, pp.38-42
16. Kharasch ED, Karol MD, Lanni C, Sawchuk R 1995, ‘Clinical sevoflurane metabolism
and disposition. I. Sevoflurane and metabolite pharmacokinetics’, Anesthesiology, vol 82, no.
6, pp.1369-78
17. Yasuda N, Lockhart SH, Eger EI, 2nd, Weiskopf RB, Liu J, Laster M 1991,
‘Comparison of kinetics of sevoflurane and isoflurane in humans’, Anesthesia and Analgesia,
vol 72, no. 3, pp. 316-24
18. Merkel G, Eger EI 1963, ‘A comparative study of halothane and halopropane
anesthesia including method for determining equipotency’, Anesthesiology, vol 24, pp. 346-
57
19. M. Perouansky RP, H. Hemmings Jr 2009, ‘Ihaled anesthetics: Mechanisms of
Action’, in Miller RD (ed), Miller's Anesthesia Seveth Edition, Churchill Livingstone,
Philadephia, pp. 515-39
20. White PF, Johnston RR, Eger EI 1974, ‘Determination of anesthetic requirement in
rats’, Anesthesiology, vol 40, no. 1, pp. 52-7
21. Orliaguet G, Vivien B, Langeron O, Bouhemad B, Coriat P, Riou B 2001, ‘Minimum
alveolar concentration of volatile anesthetics in rats during postnatal maturation’,
Anesthesiology, vol 95, no. 3, pp. 734-9.
22. de Jong RH, Eger ED 1975, ‘MAC expanded: AD50 and AD95 values of common
inhalation anesthetics in man’, Anesthesiology, vol 42, no. 4, pp.384-9
23. Zbinden AM, Petersen-Felix S, Thomson DA 1994, ‘Anesthetic depth defined using
multiple noxious stimuli during isoflurane/oxygen anesthesia. II. Hemodynamic responses’,
Anesthesiology, vol 80, no. 2, pp. 261-7
Chapter 7: References
110
24. Zbinden AM, Maggiorini M, Petersen-Felix S, Lauber R, Thomson DA, Minder CE
1994, ‘Anesthetic depth defined using multiple noxious stimuli during isoflurane/oxygen
anesthesia. I. Motor reactions’, Anesthesiology, vol 80, no. 2, pp. 253-60
25. Fourcade HE, Stevens WC, Larson CP, Jr., Cromwell TH, Bahlman SH, Hickey RF
1971, ‘The ventilatory effects of Forane, a new inhaled anesthetic’, Anesthesiology, vol 35,
no 1, pp.26-31
26. Cesarovic N, Nicholls F, Rettich A, Kronen P, Hassig M, Jirkof P 2010, ‘Isoflurane and
sevoflurane provide equally effective anaesthesia in laboratory mice’, Laboratory Animals,
vol 44, no. 4, pp.329-36
27. Crile GW 1910, Phylogenetic Association in Relation to Certain Medical Problems,
Ether Day Address, The Barta Press, Boston
28. Lundy JS 1926, ‘Balanced anesthesia’, Minnesota Medical Journal, vol 9, pp. 399-
404
29. Fukuda K 2009, Opioids, in Miller RD (ed), Miller's Anesthesia Seveth Edition,
Churchill Livingstone, Philadephia, pp. 769-825
30. Pypendop BH, Ilkiw JE 2005, ‘The effects of intravenous lidocaine administration on
the minimum alveolar concentration of isoflurane in cats’, Anesthesia and Analgesia, vol 100,
no. 1, pp. 97-101
31. Ortega M, Cruz I 2011, ‘Evaluation of a constant rate infusion of lidocaine for
balanced anesthesia in dogs undergoing surgery’, The Canadian Veterinary Journal, vol 52,
no. 8, pp. 856-60
32. Ulmer BJ, Hansen JJ, Overley CA, Symms MR, Chadalawada V, Liangpunsakul S
2003, ‘Propofol versus midazolam/fentanyl for outpatient colonoscopy: administration by
nurses supervised by endoscopists’, Clinical Gastroenterology and Hepatology, vol 1, no. 6,
pp. 425-32
33. Ryder S, Way WL, Trevor AJ 1978, ‘Comparative pharmacology of the optical
isomers of ketamine in mice’, European Journal of Pharmacology, vol 49, no. 1, pp. 15-23
Chapter 7: References
111
34. Lin H-C 2007, Dissociative Anesthetics, In William J. Tranquilli JCT, Kurt A. Grimm
(ed), ’Lumb & Jones Veterinary Anesthesia and Analgesia Fourth Edition’, Blackwell
Publishing, Iowa, pp. 301-355
35. Corssen G, Miyasaka M, Domino EF 1968, ‘Changing concepts in pain control during
surgery: dissociative anesthesia with CI-581. A progress report’, Anesthesia and Analgesia,
vol 47, no. 6, pp. 746-59
36. European Council 2010, ‘Directvive 2010/63/EU on the protection of animals used for
scientific purposes’, Official Journal of the European Union, Bruxelles
37. Leach MC, Klaus K, Miller AL, Scotto di Perrotolo M, Sotocinal SG, Flecknell PA
2012, ‘The assessment of post-vasectomy pain in mice using behaviour and the Mouse
Grimace Scale’, Public Library of Science One (Plos One), vol 7, no. 4, e35656
38. Committee on Recognition and Alleviation of Pain in Laboratory Animals, National
Research Council (CoRaAoPiL) 2009, ‘Recognition and Alleviation of Pain in Laboratory
Animals’, The National Academies Press, Washington DC
39. Arras M 2007, ‘Improvement of pain therapy in laboratory mice’, Alternativen zu
Tierexperimenten, vol 24, spec. no: 6-8
40. Richardson CA, Flecknell PA 2005, ‘Anaesthesia and post-operative analgesia
following experimental surgery in laboratory rodents: are we making progress?’, Alternatives
to Laboratory Animals, vol 33, no. 2, pp. 119-27
41. Lamont LA 2007, Opioids, Nonsteroidal Anti-inflammatories and Analgesic Adjuvants,
In William J. Tranquilli JCT, Kurt A. Grimm (ed), ’Lumb & Jones Veterinary Anesthesia and
Analgesia Fourth Edition’, Blackwell Publishing, Iowa, pp. 241 - 70.
42. Khwaja FS, Quann EJ, Pattabiraman N, Wynne S, Djakiew D 2008, ‘Carprofen
induction of p75NTR-dependent apoptosis via the p38 mitogen-activated protein kinase
pathway in prostate cancer cells’, Molecular Cancer Therapeutics, vol 7, no. 11, pp.3539-45
Chapter 7: References
112
43. Hayes KE, Raucci JA, Jr., Gades NM, Toth LA 2000, ‘An evaluation of analgesic
regimens for abdominal surgery in mice’, Contemporary Topics in Laboratory Animal
Science, vol 39, no. 6, pp.18-23
44. Bender HM 1998, ‘Pica behavior associated with buprenorphine administration in the
rat’, Laboratory Animal Science, vol 48, no. 1,pp. 5
45. Aung HH, Mehendale SR, Xie JT, Moss J, Yuan CS 2004, ‘Methylnaltrexone prevents
morphine-induced kaolin intake in the rat’, Life Sciences, vol 74, no. 22, pp. 2685-91
46. Dinda A, Gitman M, Singhal PC 2005, ‘Immunomodulatory effect of morphine:
therapeutic implications’, Expert Opinion on Drug Safety, vol 4, no. 4, pp. 669-75
47. Flores CM, Hernandez MC, Hargreaves KM, Bayer BM 1990, ‘Restraint stress-
induced elevations in plasma corticosterone and beta-endorphin are not accompanied by
alterations in immune function’, Journal of Neuroimmunology, vol 28, no. 3, pp. 219-25
48. Watanabe Y, Gould E, McEwen BS 1992, ‘Stress induces atrophy of apical dendrites
of hippocampal CA3 pyramidal neurons’, Brain Research, vol 588, no. 2, pp. 341-5
49. Li S, Fan YX, Wang W, Tang YY 2012, ‘Effects of acute restraint stress on different
components of memory as assessed by object-recognition and object-location tasks in mice’,
Behavioural Brain Research, vol 227, no. 1, pp.199-207
50. Howland JG, Cazakoff BN 2005, ‘Effects of acute stress and GluN2B-containing
NMDA receptor antagonism on object and object-place recognition memory’, Neurobiology of
Learning and Memory, vol 93, no. 2, pp. 261-7
51. Brockway BP, Mills PA, Azar SH 1991, ‘A new method for continuous chronic
measurement and recording of blood pressure, heart rate and activity in the rat via radio-
telemetry’, Clinical and Experimental Hypertension Part A, Theory and Practice vol 13, no. 5,
pp. 885-95
52. Riley JL 1970, ‘Radio telemetry system for transmitting deep body temperatures’, The
Cornell Veterinarian, vol 60, no. 2, pp. 265-73
Chapter 7: References
113
53. Carson VG, Kado RT, Wenzel BM 1972, ‘A telemeter for monitoring the
electrocardiograms of freely moving mice’, Physiolology and Behavior, vol 8, no. 3, pp. 561-3
54. Kramer K, van Acker SA, Voss HP, Grimbergen JA, van der Vijgh WJ, Bast A 1993,
‘Use of telemetry to record electrocardiogram and heart rate in freely moving mice’, Journal
of Pharmacological and Toxicological Methods, vol 30, no. 4, pp. 209-215
55. Data Sciences International LTD, revised 2012, implantable-telemetry devices and
specification overview 2013, viewed 11.11.2013, URL:
http://www.datasci.com/products/implantable-telemetry/specification-overview.
56. Weiergraber M, Henry M, Hescheler J, Smyth N, Schneider T 2005,
‘Electrocorticographic and deep intracerebral EEG recording in mice using a telemetry
system’, Brain Research Protocols, vol 14, no. 3, pp. 154-64
57. Gross V, Luft FC 2003, ‘Exercising restraint in measuring blood pressure in conscious
mice’, Hypertension, vol 41, no. 4, pp. 879-81
58. Kramer K, Kinter LB 2003, ‘Evaluation and applications of radiotelemetry in small
laboratory animals’, Physiological Genomics, vol 13, no. 3, pp. 197-205
59. Schuler B, Rettich A, Vogel J, Gassmann M, Arras M 2009, ‘Optimized surgical
techniques and postoperative care improve survival rates and permit accurate telemetric
recording in exercising mice’, BMC Veterinary Research, vol 28, no. 5, viewed 11.11.2013, <
http://www.biomedcentral.com/1746-6148/5/28 >
60. Pritchett-Corning KR, Luo Y, Mulder GB, White WJ 2011, ‘Principles of rodent surgery
for the new surgeon’, Journal of Visualized Experiments, vol 47, viewed 11.11.2013, <
http://www.jove.com/video/2586/principles-of-rodent-surgery-for-the-new-surgeon?ID=2586 >
61. Rettich A, Kasermann HP, Pelczar P, Burki K, Arras M 2006, ‘The physiological and
behavioral impact of sensory contact among unfamiliar adult mice in the laboratory’, Journal
of Applied Animal Welfare Science, vol 9, no. 4, pp. 277-88
62. Spani D, Arras M, Konig B, Rulicke T 2003, ‘Higher heart rate of laboratory mice
housed individually vs in pairs’, Laboratory Animals, vol 37, no. 1, pp. 54-62
Chapter 7: References
114
63. Arras M, Rettich A, Cinelli P, Kasermann HP, Burki K 2007, ‘Assessment of post-
laparotomy pain in laboratory mice by telemetric recording of heart rate and heart rate
variability’, BMC Veterinary Research, vol. 16, no. 3, viewed 11.11.2013, <
http://www.biomedcentral.com/1746-6148/3/16 >
64. Zeller A, Arras M, Jurd R, Rudolph U 2007, ‘Mapping the contribution of beta3-
containing GABAA receptors to volatile and intravenous general anesthetic actions’, BMC
Pharmacology, vol 7, no. 2, viewed 11.11.2013, <
http://www.biomedcentral.com/1471-2210/7/2 >
65. Zeller A, Arras M, Jurd R, Rudolph U 2007, ‘Identification of a molecular target
mediating the general anesthetic actions of pentobarbital’, Molecular Pharmacology, vol 71,
no. 3, pp. 852-9
66. Hacker SO, White CE, Black IH 2005, ‘A comparison of target-controlled infusion
versus volatile inhalant anesthesia for heart rate, respiratory rate, and recovery time in a rat
model’, Contemporary Topics in Laboratory Animal Science, vol 44, no. 5, pp. 7-12
67. Alves HC, Valentim AM, Olsson IA, Antunes LM 2007, ‘Intraperitoneal propofol and
propofol fentanyl, sufentanil and remifentanil combinations for mouse anaesthesia’,
Laboratory Animals, vol 41, no. 3, pp. 329-36
68. Arras M, Autenried P, Rettich A, Spaeni D, Rulicke T 2001, ‘Optimization of
intraperitoneal injection anesthesia in mice: drugs, dosages, adverse effects, and anesthesia
depth’, Comparative Medicine, vol 51, no. 5, pp. 443-56
69. Thal SC, Plesnila N 2007, ‘Non-invasive intraoperative monitoring of blood pressure
and arterial pCO2 during surgical anesthesia in mice’, Journal of Neuroscience Methods, vol
159, no. 2, pp. 261-7
70. Cruz JI, Loste JM, Burzaco OH 1998, ‘Observations on the use of
medetomidine/ketamine and its reversal with atipamezole for chemical restraint in the
mouse’, Laboratory Animals, vol 32, no. 1,pp. 18-22
Chapter 7: References
115
71. Homanics GE, Quinlan JJ, Firestone LL 1999, ‘Pharmacologic and behavioral
responses of inbred C57BL/6J and strain 129/SvJ mouse lines’, Pharmacology, Biochemistry
and Behavior, vol 63, no. 1, pp. 21-6
72. Lovell DP 1986, ‘Variation in pentobarbitone sleeping time in mice. 2. Variables
affecting test results’, Laboratory Animals, vol 20, no. 2, pp. 91-6
73. Lovell DP 1986, ‘Variation in pentobarbitone sleeping time in mice. 1. Strain and sex
differences’, Laboratory Animals, vol 20, no. 2, pp. 85-90
74. Zuurbier CJ, Emons VM, Ince C 2002, ‘Hemodynamics of anesthetized ventilated
mouse models: aspects of anesthetics, fluid support, and strain’, American Journal of Heart
and Circulatory Physiology, vol 282, no. 6, pp. 2099-105
75. Woodward WR, Choi D, Grose J, Malmin B, Hurst S, Pang J 2007, ‘Isoflurane is an
effective alternative to ketamine/xylazine/acepromazine as an anesthetic agent for the
mouse electroretinogram’, Documenta Ophtalmologica – Advences in Ophtalmology, vol
115, no. 3, pp. 187-201
76. Heavner JE 2001, ‘Anesthesia update: agents, definitions, and strategies’,
Comparative Medicine, vol 51, no. 6, pp. 500-3
77. Szczesny G, Veihelmann A, Massberg S, Nolte D, Messmer K 2004, ‘Long-term
anaesthesia using inhalatory isoflurane in different strains of mice-the haemodynamic
effects’, Laboratory Animals, vol 38, no. 1, pp. 64-9
78. Diven K 2003, ‘Inhalation anesthetics in rodents’, Laboratory Animals, vol 32, no. 3,
pp. 44-7
79. Dubin S 1999, ‘Use of a biocompatible adhesive paste to improve the performance of
rodent anesthesia face masks’, Laboratory Animals, vol 28, no. 8, pp. 50-1
80. Horne D, Ogden B, Houts J, Hall A 1998, ‘A nonrebreathing anesthetic delivery
system for mice’, Laboratory Animals, vol 27, no. 7, pp. 32-4
81. Klein RC 2008, ‘Fire safety recommendations for administration of isoflurane
anesthesia in oxygen’, Laboratory Animals, vol 37, no. 5, pp. 223-4
Chapter 7: References
116
82. Smith JC, Bolon B 2002, ‘Atmospheric waste isoflurane concentrations using
conventional equipment and rat anesthesia protocols’, Contemporary Topics in Laboratory
Animal Science, vol 41, no. 2,pp. 10-7
83. Smith JC, Bolon B 2003, ‘Comparison of three commercially available activated
charcoal canisters for passive scavenging of waste isoflurane during conventional rodent
anesthesia’, Comparative Medicine, vol 42, no. 2, pp.10-5
84. Smith JC, Bolon B 2006, ‘Isoflurane leakage from non-rebreathing rodent
anaesthesia circuits: comparison of emissions from conventional and modified ports’,
Laboratory Animals, vol 40, no. 2, pp. 200-9
85. Fleisher LA, Beckman JA, Brown KA, Calkins H, Chaikof E, Fleischmann KE 2007,
‘Guidelines on perioperative cardiovascular evaluation and care for noncardiac surgery: a
report of the American College of Cardiology/American Heart Association’, Task Force on
Practice Guidelines (Writing Committee to Revise the 2002 Guidelines on Perioperative
Cardiovascular Evaluation for Noncardiac Surgery): developed in collaboration with the
American Society of Echocardiography, American Society of Nuclear Cardiology, Heart
Rhythm Society, Society of Cardiovascular Anesthesiologists, Society for Cardiovascular
Angiography and Interventions, Society for Vascular Medicine and Biology, and Society for
Vascular Surgery, Circulation, vol 116, no. 17, pp. 418-99
86. Nicklas W, Baneux P, Boot R, Decelle T, Deeny AA, Fumanelli M 2002,
‘Recommendations for the health monitoring of rodent and rabbit colonies in breeding and
experimental units’, Laboratory Animals, vol 36, no. 1, pp. 20-42
87. Joo DT, Gong D, Sonner JM, Jia Z, MacDonald JF, Eger EI 2001, ‚Blockade of AMPA
receptors and volatile anesthetics: reduced anesthetic requirements in GluR2 null mutant
mice for loss of the righting reflex and antinociception but not minimum alveolar
concentration’, Anesthesiology, vol 94, no. 3, pp. 478-88
88. Sonner JM, Gong D, Eger EI 2000, ‘Naturally occurring variability in anesthetic
potency among inbred mouse strains’, Anesthesia and Analgesia, vol 91, no. 3, pp. 720-6
Chapter 7: References
117
89. Sonner JM, Gong D, Li J, Eger EI, Laster MJ 1999, ‘Mouse strain modestly influences
minimum alveolar anesthetic concentration and convulsivity of inhaled compounds’,
Anesthesia and Analgesia, vol 89, no. 4, pp. 1030-4
90. Ichinose F, Mi WD, Miyazaki M, Onouchi T, Goto T, Morita S 1998, ‘Lack of
correlation between the reduction of sevoflurane MAC and the cerebellar cyclic GMP
concentrations in mice treated with 7-nitroindazole’, Anesthesiology, vol 89, no. 1, pp. 143-8
91. Mogil JS, Smith SB, O'Reilly MK, Plourde G 2005, ‘Influence of nociception and
stress-induced antinociception on genetic variation in isoflurane anesthetic potency among
mouse strains’, Anesthesiology, vol 103, no. 4, pp. 751-8
92. Doherty TJ, Geiser DR, Frazier DL 1997, ‘Comparison of halothane minimum alveolar
concentration and minimum effective concentration in ponies’, Journal of Veterinary
Pharmacology and Therapeutics, vol 20, no. 5, pp. 408-10
93. Janssen BJ, De Celle T, Debets JJ, Brouns AE, Callahan MF, Smith TL 2004, ‘Effects
of anesthetics on systemic hemodynamics in mice’, American Journal of Heart and
Circulatory Physiology, vol 287, no. 4, pp. 1618-24
94. Loepke AW, McCann JC, Kurth CD, McAuliffe JJ 2006, ‘The physiologic effects of
isoflurane anesthesia in neonatal mice’, Anesthesia and Analgesia, vol 102, no. 1, pp. 75-80
95. Matsuda Y, Ohsaka K, Yamamoto H, Natsume K, Hirabayashi S, Kounoike M 2007,
‘Comparison of newly developed inhalation anesthesia system and intraperitoneal
anesthesia on the hemodynamic state in mice’, Biological and Pharmaceutical Bulletin, vol
30, no. 9, pp. 1716-20
96. Henke J, Strack T, Erhardt W 2004, ‘Klinischer Vergleich einer Iso- und
Sevofluranmononarkose beim Gerbil (Meriones unguiculatus)’, Berliner und Munchener
tierarztliche Wochenschrift, vol 117, no. 7-8, pp. 296-303
97. Sjoblom M, Nylander O 2007, ‘Isoflurane-induced acidosis depresses basal and
PGE(2)-stimulated duodenal bicarbonate secretion in mice’, American Journal of
Gastrointestinal and Liver Physiology, vol 292, no. 3, pp. 899-904
Chapter 7: References
118
98. Wiersema AM, Dirksen R, Oyen WJ, Van der Vliet JA 1997, ‘A method for long
duration anaesthesia for a new hindlimb ischaemia-reperfusion model in mice’, Laboratory
Animals, vol 31, no. 2, pp. 151-6
99. Dahan A, Teppema LJ 2003, ‘Influence of anaesthesia and analgesia on the control
of breathing’, British Journal of Anaesthesia, vol 91, no. 1, pp. 40-9
100. Baumans V 2005, ‘Science-based assessment of animal welfare: laboratory animals’,
Revue Scientifique et Technique, vol 24, no. 2, pp. 503-13
101. Stasiak KL, Maul D, French E, Hellyer PW, VandeWoude S 2003, ‘Species-specific
assessment of pain in laboratory animals’, Contemporary Topics in Laboratory Animal
Science, vol 42, no. 4, pp. 13-20
102. Flores JE, McFarland LM, Vanderbilt A, Ogasawara AK, Williams SP 2008, ‘The
effects of anesthetic agent and carrier gas on blood glucose and tissue uptake in mice
undergoing dynamic FDG-PET imaging: sevoflurane and isoflurane compared in air and in
oxygen’, Molecular Imaging and Biology, vol 10, no. 4, pp. 192-200
103. Groeben H, Meier S, Tankersley CG, Mitzner W, Brown RH 2003, ‘Heritable
differences in respiratory drive and breathing pattern in mice during anaesthesia and
emergence’, British Journal of Anaesthesia, vol 91, no. 4, pp. 541-5
104. Groeben H, Meier S, Tankersley CG, Mitzner W, Brown RH 2004, ‘Influence of
volatile anaesthetics on hypercapnoeic ventilatory responses in mice with blunted respiratory
drive’, British Journal of Anaesthesia, vol 92, no. 5, pp. 697-703
105. Cornett PM, Matta JA, Ahern GP 2008, ‘General anesthetics sensitize the capsaicin
receptor transient receptor potential V1’, Molecular Pharmacology, vol 74, no. 5, pp. 1261-8
106. Eilers H 2008, ‘Anesthetic activation of nociceptors: adding insult to injury?’,
Molecular Interventions, vol 8, no. 5, pp. 226-9
107. Enderle AK, Levionnois OL, Kuhn M, Schatzmann U 2008, ‘Clinical evaluation of
ketamine and lidocaine intravenous infusions to reduce isoflurane requirements in horses
Chapter 7: References
119
under general anaesthesia’, Veterinary Anaesthesia and Analgesia, vol 35, no. 4, pp. 297-
305
108. Hendrickx JF, Eger EI, Sonner JM, Shafer SL 2008, ‘Is synergy the rule? A review of
anesthetic interactions producing hypnosis and immobility’, Anesthesia and Analgesia, vol
107, no. 2, pp. 494-506
109. Muir WW, Wiese AJ, March PA 2003, ‘Effects of morphine, lidocaine, ketamine, and
morphine-lidocaine-ketamine drug combination on minimum alveolar concentration in dogs
anesthetized with isoflurane’, American Journal of Veterinary Research, vol 64, no. 9, pp.
1155-60
110. Tonner PH 2005, ‘Balanced anaesthesia today’, Best Practice and Research, vol 19,
no. 3, pp. 475-84
111. Filibeck U, Castellano C 1980, ‘Strain dependent effects of ketamine on locomotor
activity and antinociception in mice’, Pharmacology, biochemistry and Behavior,.vol 13, no. 3,
pp. 443-7
112. Nishizawa N, Nakao S, Nagata A, Hirose T, Masuzawa M, Shingu K 2000, ‘The
effect of ketamine isomers on both mice behavioral responses and c-Fos expression in the
posterior cingulate and retrosplenial cortices’, Brain Research, vol 857, no. 1-2, pp. 188-92
113. Sarton E, Teppema LJ, Olievier C, Nieuwenhuijs D, Matthes HW, Kieffer BL 2001,
‚The involvement of the mu-opioid receptor in ketamine-induced respiratory depression and
antinociception’, Anesthesia and Analgesia, vol 93, no. 6, pp. 1495-500
114. Kip A.L. 2007, ‘Anticholinergics and sedatives’, In William J. Tranquilli JCT, Kurt A.
Grimm (ed), ’Lumb & Jones Veterinary Anesthesia and Analgesia Fourth Edition’, Blackwell
Publishing, Iowa, pp 203–39
115. Boehm CA, Carney EL, Tallarida RJ, Wilson RP 2010, ‘Midazolam enhances the
analgesic properties of dexmedetomidine in the rat’, Veterinary Anaesthesia and Analgesia,
vol 37, no. 6, pp. 550-6
Chapter 7: References
120
116. Theil DR, Stanley TE, 3rd, White WD, Goodman DK, Glass PS, Bai SA 1993,
‘Midazolam and fentanyl continuous infusion anesthesia for cardiac surgery: a comparison of
computer-assisted versus manual infusion systems’, Journal of Cardiothoracic and Vascular
Anesthesia, vol 7, no. 3, pp. 300-6
117. Bhargava HN 1981, ‘Antagonism of ketamine-induced anesthesia and hypothermia
by thyrotropin releasing hormone and cyclo(His-Pro)’, Neuropharmacology, vol 20, no. 7, pp.
699-702
118. European Union Council 1986, ‘Directive on the approximation of laws, regulations
and administrative provisions of the Member States regarding the protection of animals used
for experimental and other scientific purposes’, 86/609/EEC, Official Journal of European
Communities, Bruxelles
119. Chae YJ, Zhang J, Au P, Sabbadini M, Xie GX, Yost CS 2010, ‘Discrete change in
volatile anesthetic sensitivity in mice with inactivated tandem pore potassium ion channel
TRESK’, Anesthesiology, vol 113, no. 6, pp. 1326-37
120. Eckel B, Richtsfeld M, Starker L, Blobner M 2010, ‘Transgenic Alzheimer mice have a
larger minimum alveolar anesthetic concentration of isoflurane than their nontransgenic
littermates’, Anesthesia and Analgesia, vol 110, no. 2, pp. 438-41
121. Sonner JM 2002, ‘Issues in the design and interpretation of minimum alveolar
anesthetic concentration (MAC) studies’, Anesthesia and Analgesia, vol 95, no. 3, pp. 609-
14
122. Arras M, Rettich A, Seifert B, Kasermann HP, Rulicke T 2007, ‘Should laboratory
mice be anaesthetized for tail biopsy?’, Laboratory Animals, vol 41, no. 1, pp. 30-45
123. Jurd R, Arras M, Lambert S, Drexler B, Siegwart R, Crestani F 2003, ‘General
anesthetic actions in vivo strongly attenuated by a point mutation in the GABA(A) receptor
beta3 subunit’, Federation of American Societies for Experimental Biology Journal, vol 17,
no. 2, pp. 250-2
Chapter 7: References
121
124. Liao M, Laster MJ, Eger EI, Tang M, Sonner JM 2006, ‘Naloxone does not increase
the minimum alveolar anesthetic concentration of sevoflurane in mice’, Anesthesia and
Analgesia, vol 102, no. 5, pp. 1452-5
125. Doherty TJ, Frazier DL 1998, ‘Effect of intravenous lidocaine on halothane minimum
alveolar concentration in ponies’, Equine Veterinary Journal, vol 30, no. 4, pp. 300-3
126. Valverde A, Doherty TJ, Hernandez J, Davies W 2004, ‘Effect of lidocaine on the
minimum alveolar concentration of isoflurane in dogs’, Veterinary Anaesthesia and
Analgesia, vol 31, no. 4, pp. 264-71
127. Inagaki Y, Sumikawa K, Yoshiya I 1993, ‘Anesthetic interaction between midazolam
and halothane in humans’, Anesthesia and Analgesia, vol 76, no. 3, pp. 613-7
128. Melvin MA, Johnson BH, Quasha AL, Eger EI 1982, ‘Induction of anesthesia with
midazolam decreases halothane MAC in humans’, Anesthesiology, vol 57, no. 3, pp. 238-41
129. Sebel PS, Glass PS, Fletcher JE, Murphy MR, Gallagher C, Quill T 1992, ‘Reduction
of the MAC of desflurane with fentanyl’, Anesthesiology, vol 76, no. 1, pp. 52-9
130. Solano AM, Pypendop BH, Boscan PL, Ilkiw JE 2006, ‘Effect of intravenous
administration of ketamine on the minimum alveolar concentration of isoflurane in
anesthetized dogs’, American Journal of Veterinary Research, vol 67, no. 1, pp. 21-5
131. de Jong RH, Eger EI 1975, ‘MAC expanded: AD50 and AD95 values of common
inhalation anesthetics in man’, Anesthesiology, vol 42, no. 4, pp. 384-9
132. Ulugol A, Dost T, Dokmeci D, Akpolat M, Karadag CH, Dokmeci I 2000, ‘Involvement
of NMDA receptors and nitric oxide in the thermoregulatory effect of morphine in mice’,
Journal of Neural Transmitters, vol 107, no. 5, pp. 515-21
133. Lee EJ, Woodske ME, Zou B, O'Donnell CP 2009, ‘Dynamic arterial blood gas
analysis in conscious, unrestrained C57BL/6J mice during exposure to intermittent hypoxia’,
Journal of Applied Physiology, vol 107, no. 1, pp. 290-4
Chapter 7: References
122
134. Lam AM, Clement JL, Knill RL 1980, ‘Surgical stimulation does not enhance
ventilatory chemoreflexes during enflurane anaesthesia in man’, Canadian Anaesthetists'
Society Journal, vol 27, no. 1, pp. 22-8
135. Sutherland RW, Drummond GB 1996, ‘Effects of surgical skin incision on respiration
in patients anaesthetized with enflurane’, British Journal of Anaesthesia, vol 76, no. 6, pp.
777-9
136. Matsumiya LC, Sorge RE, Sotocinal SG, Tabaka JM, Wieskopf JS, Zaloum A 2012,
‘Using the Mouse Grimace Scale to reevaluate the efficacy of postoperative analgesics in
laboratory mice’, Journal of American Associacion for Laboratory Animal Sciences, vol 51,
no. 1, pp. 42-9
137. Jirkof P, Fleischmann T, Cesarovic N, Rettich A, Vogel J, Arras M 2013, ‘Assessment
of postsurgical distress and pain in laboratory mice by nest complexity scoring’, Laboratory
Animals, vol 47, no. 3, pp. 153-61
138. Valentim AM, Alves HC, Olsson IA, Antunes LM 2008, ‘The effects of depth of
isoflurane anesthesia on the performance of mice in a simple spatial learning task’, Journal
of American Associacion for Laboratory Animal Sciences, vol 47, no. 3, pp. 16-9
139. Petrenko AB, Kohno T, Wu J, Sakimura K, Baba H 2008, ‘Spontaneous hyperactivity
in mutant mice lacking the NMDA receptor GluRepsilon1 subunit is aggravated during
exposure to 0.1 MAC sevoflurane and is preserved after emergence from sevoflurane
anaesthesia’, European Journal of Anaesthesiology, vol 25, no. 12, pp. 953-60
140. Mena MA, Perucho J, Rubio I, de Yebenes JG 2010, ‘Studies in animal models of the
effects of anesthetics on behavior, biochemistry, and neuronal cell death’, Journal of
Alzheimer’s Disease, vol 22, Suppl 3, pp. 43-8
141. Roughan JV, Flecknell PA 2000, ‘Effects of surgery and analgesic administration on
spontaneous behaviour in singly housed rats’, Research in Veterinary Science, vol 69, no. 3,
pp. 283-8
142. Van Oortmerssen GA 1970, ‘Biological significance, genetics and evolutionary origin
of variability in behaviour of inbred strains of mice’, Behaviour, vol 38, no. 1, pp. 1-92.
Chapter 7: References
123
143. Roughan JV, Wright-Williams SL, Flecknell PA 2009, ‘Automated analysis of
postoperative behaviour: assessment of HomeCageScan as a novel method to rapidly
identify pain and analgesic effects in mice’, Laboratory Animals, vol 43, no. 1, pp. 17-26
144. Jirkof P, Cesarovic N, Rettich A, Nicholls F, Seifert B, Arras M 2010, ‘Burrowing
behavior as an indicator of post-laparotomy pain in mice’, Frontiers in Behavioral
Neuroscience, vol 4, doi: 10.3389/fnbeh.2010.00165.eCollection 2010.
145. Wright-Williams SL, Courade JP, Richardson CA, Roughan JV, Flecknell PA 2007,
‘Effects of vasectomy surgery and meloxicam treatment on faecal corticosterone levels and
behaviour in two strains of laboratory mouse’, Pain, vol 130, no. 1-2, pp. 108-18
146. Oishi K, Ohkura N, Sei H, Matsuda J, Ishida N 2007, ‘CLOCK regulates the circadian
rhythm of kaolin-induced writhing behavior in mice’, Neuroreport, vol 18, no. 18, pp. 1925-8
147. Chassard D, Bruguerolle B 2004, ‘Chronobiology and anesthesia’, Anesthesiology,
vol 100, no. 2, pp. 413-27
148. Butz GM, Davisson RL 2001, ‘Long-term telemetric measurement of cardiovascular
parameters in awake mice: a physiological genomics tool’, Physiological Genomics, vol 5,
no. 2, pp. 89-97
149. Mills PA, Huetteman DA, Brockway BP, Zwiers LM, Gelsema AJ, Schwartz RS 2000,
‘A new method for measurement of blood pressure, heart rate, and activity in the mouse by
radiotelemetry’, Journal of Applied Physiology, vol 88, no. 5, pp. 1537-44
150. Farber N, Warltier D 2009, ‘Pulmonary Pharmacology’, in Miller RD (ed), Miller's
Anesthesia Seventh Edition, Churchill Livingstone, Philadephia, pp. 561 - 90
151. Reves JG, Glass P, Lubarsky D, McEvoy M, Martinez-Ruiz R 2009, ‘Intravenous
Anesthetics’, in Miller RD (ed), Miller's Anesthesia Seventh Edition, Churchill Livingstone,
Philadephia, pp. 719 - 60.
152. Daniel M, Weiskopf RB, Noorani M, Eger El 1998, ‘Fentanyl augments the blockade
of the sympathetic response to incision (MAC-BAR) produced by desflurane and isoflurane:
Chapter 7: References
124
desflurane and isoflurane MAC-BAR without and with fentanyl’, Anesthesiology, vol 88, no. 1,
pp. 43-9
153. Criado AB, Gomez e Segura IA 2003, ‘Reduction of isoflurane MAC by fentanyl or
remifentanil in rats’, Veterinary Anaesthesia and Analgesia, vol 30, no. 4, pp. 250-6
154. Seddighi R, Egger CM, Rohrbach BW, Cox SK, Doherty TJ 2011, ‘The effect of
midazolam on the end-tidal concentration of isoflurane necessary to prevent movement in
dogs’, Veterinary Anaesthesia and Analgesia, vol 38, no. 3, pp. 195-202
155. Chiba S, Nishiyama T, Yoshikawa M, Yamada Y 2009, ‘The antinociceptive effects of
midazolam on three different types of nociception in mice’, Journal of Pharmacological
Sciences, vol 109, no. 1, pp. 71-7
156. Coulthard P, Rood JP 1992, ‘An investigation of the effect of midazolam on the pain
experience’, The British Journal of Oral and Maxillofacial Surgery, vol 30, no. 4, pp. 248-51
157. White PF, Way WL, Trevor AJ 1982, ‘Ketamine--its pharmacology and therapeutic
uses’, Anesthesiology, vol 56, no. 2, pp. 119-36
158. Irifune M, Shimizu T, Nomoto M, Fukuda T 1992, ‘Ketamine-induced anesthesia
involves the N-methyl-D-aspartate receptor-channel complex in mice’, Brain Research, vol
596, no. 1-2, pp. 1-9
159. Soliman MG, Brindle GF, Kuster G 1975, ‘Response to hypercapnia under ketamine
anaesthesia’, Canadian Anaesthetists' Society Journal, vol 22, no. 4, pp. 486-94
160. Doenicke A, Angster R, Mayer M, Adams HA, Grillenberger G, Nebauer AE 1992,
‘Die Wirkung von S-(+)-Ketamin auf Katecholamine und Cortisol im Serum. Vergleich zu
Ketamin-Razemat’, Der Anaesthesist, vol 41, no. 10, pp. 597-603
161. Endou M, Hattori Y, Nakaya H, Gotoh Y, Kanno M 1992, ‘Electrophysiologic
mechanisms responsible for inotropic responses to ketamine in guinea pig and rat
myocardium’, Anesthesiology, vol 76, no. 3, pp. 409-18
162. Child KJ, Davis B, Dodds MG, Twissell DJ 1972, ‘Anaesthetic, cardiovascular and
respiratory effects of a new steroidal agent CT 1341: a comparison with other intravenous
Chapter 7: References
125
anaesthetic drugs in the unrestrained cat’, British Journal of Pharmacology, vol 46, no. 2, pp.
189-200
163. Buitrago S, Martin TE, Tetens-Woodring J, Belicha-Villanueva A, Wilding GE 2008,
‘Safety and efficacy of various combinations of injectable anesthetics in BALB/c mice’,
Journal of American Associacion for Laboratory Animal Sciences, vol 47, no.1, pp. 11-7
164. Zhang Z, Xu F, Zhang C, Liang X 2007, ‘Activation of opioid mu receptors in caudal
medullary raphe region inhibits the ventilatory response to hypercapnia in anesthetized rats’,
Anesthesiology, vol 107, no. 2, pp. 288-97
165. McCrimmon DR, Alheid GF 2003, ‘On the opiate trail of respiratory depression’,
American Journal of Regulatory, Integrative and Comparative Physiology, vol 285, no. 6, pp.
1274-5
166. Sunzel M, Paalzow L, Berggren L, Eriksson I 1988, ‘Respiratory and cardiovascular
effects in relation to plasma levels of midazolam and diazepam’, British Journal of Clinical
Pharmacology, vol 25, no. 5, pp. 561-9
167. Dahan A, Yassen A, Bijl H, Romberg R, Sarton E, Teppema L 2005, ‘Comparison of
the respiratory effects of intravenous buprenorphine and fentanyl in humans and rats’, British
Journal of Anaesthesia, vol 94, no. 6, pp. 825-34
168. Cowan A, Doxey JC, Harry EJ 1977, ‘The animal pharmacology of buprenorphine, an
oripavine analgesic agent’, British Journal of Pharmacology, vol 60, no. 6, pp. 547-54
169. Kishioka S, Paronis CA, Lewis JW, Woods JH 2000, ‘Buprenorphine and
methoclocinnamox: agonist and antagonist effects on respiratory function in rhesus
monkeys’, European Journal of Pharmacology, vol 391, no. 3, pp. 289-97
170. Dorre N 2009, ‘The Postanesthesia Care Unit’, in Miller RD (ed), Miller's Anesthesia
Seventh Edition, Churchill Livingstone, Philadephia, pp. 2707 - 28
171. Spruijt BM, van Hooff JA, Gispen WH. Ethology and neurobiology of grooming
behavior. Physiological reviews. 1992 Jul;72(3):825-52. PubMed PMID: 1320764.
Chapter 7: References
126
172. Bindra D, Spinner N 1958, ‘Response to different degrees of novelty: the incidence of
various activities’, Journal of the Experimental Analysis of Behavior, vol 1, no. 4, pp. 341-50
173. Colbern DL, Isaacson RL, Green EJ, Gispen WH 1978, ‘Repeated intraventricular
injections of ACTH 1-24: the effects of home or novel environments on excessive grooming’,
Behavioral Biology, vol 23, no. 3, pp. 381-7
174. Jirkof P, Cesarovic N, Rettich A, Fleischmann T, Arras M 2012, ‘Individual housing of
female mice: influence on postsurgical behaviour and recovery’, Laboratory Animals, vol 46,
no. 4, pp. 325-34
175. Imaizumi M, Sawano S, Takeda M, Fushiki T 2000, ‘Grooming behavior in mice
induced by stimuli of corn oil in oral cavity’, Physiology and Behaviour, vol 71, no. 3-4, pp.
409-14
176. Marin C, Engber TM, Chaudhuri P, Peppe A, Chase TN 1996, ‘Effects of kappa
receptor agonists on D1 and D2 dopamine agonist and antagonist-induced behaviors’,
Psychopharmacology, vol 123, no. 2, pp. 215-21
177. Stoessl AJ 1994, ‘Dopamine D1 receptor agonist-induced grooming is blocked by the
opioid receptor antagonist naloxone’, European Journal of Pharmacology, vol 259, no. 3, pp.
301-3
178. Toyoshi T, Ukai M, Kameyama T 1992, ‘Combination of a delta opioid receptor
agonist but not a mu opioid receptor agonist with the D1-selective dopamine receptor agonist
SKF 38393 markedly potentiates different behaviors in mice, European Journal of
Pharmacology, vol 213, no.1, pp. 25-30
179. Taylor NE, Chemali JJ, Brown EN, Solt K 2013, ‘Activation of D1 dopamine receptors
induces emergence from isoflurane general anesthesia’, Anesthesiology, vol 118, no.1, pp.
30-9
180. Su D, Zhao Y, Xu H, Wang B, Chen X, Chen J 2012, ‘Isoflurane exposure during mid-
adulthood attenuates age-related spatial memory impairment in APP/PS1 transgenic mice’,
Public Library of Science One (Plos One), vol 7, no. 11, e50172
Chapter 7: References
127
181. Christian A 2009, ‘Postoperative Nausea and Vomiting’, in Miller RD (ed), Miller's
Anesthesia seveth edition, Churchill Livingstone, Philadephia, pp. 2729 - 55
182. Watcha MF, White PF 1992, ‘Postoperative nausea and vomiting. Its etiology,
treatment and prevention’, Anesthesiology, vol 77, no. 1, pp. 162-84
183. Horn CC, Kimball BA, Wang H, Kaus J, Dienel S, Nagy A 2013, ‘Why can't rodents
vomit? A comparative behavioral, anatomical, and physiological study, Public Library of
Science One (Plos One), vol 8, no. 4, e60537
184. Sanger GJ, Holbrook JD, Andrews PL 2011, ‘The translational value of rodent
gastrointestinal functions: a cautionary tale’, Trends in Pharmacological Sciences, vol 32, no.
7, pp. 402-9
185. Zambricki EA, Dalecy LG 2004, ‘Rat sex differences in anesthesia’, Comparative
Medicine, vol 54, no. 1, pp. 49-53
186. Dairman W, Balazs T 1970, ‘Comparison of liver microsome enzyme systems and
barbiturate sleep times in rats caged individually or communally’, Biochemical Pharmacology,
vol 19, no. 3, pp. 951-5
187. Quinlan JJ, Homanics GE, Firestone LL 1998, ‘Anesthesia sensitivity in mice that lack
the beta3 subunit of the gamma-aminobutyric acid type A receptor’, Anesthesiology, vol 88,
no. 3, pp. 775-80
188. He S, Atkinson C, Qiao F, Chen X, Tomlinson S 2010, ‘Ketamine-xylazine-
acepromazine compared with isoflurane for anesthesia during liver transplantation in
rodents’, Journal of American Associacion for Laboratory Animal Sciences, vol 49, no. 1, pp.
45-51
189. Woodward WR, Choi D, Grose J, Malmin B, Hurst S, Pang J 2007, ‘Isoflurane is an
effective alternative to ketamine/xylazine/acepromazine as an anesthetic agent for the
mouse electroretinogram’, Documenta Ophthalmologica Advances in Ophthalmology, vol
115, no.3, pp. 187-201
Appendix 1
128
Appendix 1
General condition and health monitoring data sheet for ECG-transmitter implantation in mice
Humane endpoints
If in poor general condition, i.e. the animal is substantially apathetic (no movement after being touched/pushed) and its body surface feels cold
despite warming, the animal should be euthanatized immediately.
If on day 4 after transmitter implantation the animal shows clear signs of apathy, is extremely aggressive or does not show any food intake, the
animal should be euthanatized immediately.
On day 8 after transmitter implantation the animal has to display a clear increase in body weight in comparison to the preceding post-operative
days. Moreover, it has to consume at least 80% of its pre-operative daily food intake. If one of these conditions is not met, the animal should be
euthanatized immediately.
Appendix 1
129
Date
Time
of
day
Body weight
(g)
Food
(g)
Water bottle
(g)
Glucose 15%
Bottle
(g)
Abnormalities of
outer
appearance,
posture, and
spontaneous
behaviour
Analgesic
s.c
[Buprenorphin
0.1 mg/kg]
Fluid s.c.
[300 μL
glucose (5%)
and 300 μL
saline (0.9%)]
Warming
cabinet
temp (°C)
Sign.
Transmitter implantation (surgeon’s notes)
Curriculum vitae
130
Acknowledgments
I thank Prof. Dr. Wolfgang Langhans, who gave me the opportunity to study and complete my PhD work under his supervision on ETH Zürich.
Special thanks go to my direct supervisor PD Dr. Margarete Arras, who advised, challenged
and supported me throughout my PhD. She contributed substantially to my development not
only as a doctoral student but as a research veterinarian as well.
Then I would also like to thank Prof. Dr. Beatrice Beck Schimmer for being my co-examiner
and provided me with crucial scientific input.
Further, I would like to thank Prof. Dr. Gregor Zünd, Dr. Hugo Battalia, Mr. Robin Schneider
and other members of the Center for Clinical Research, University Hospital Zürich for
providing me and my team with resources and environment in which we could strive and
prosper.
Moreover, I thank all members of Department of Surgical Research and Central Biological
Laboratory, especially my team members Dr. Paulin Jirkof, Dr. Thea Fleischmann and Dipl.
Biol. Flora Nicholls for constant and unselfish support over the years. None of this would be
possible without you guys!
Finally, I would like to thank my family.
Mama, tata puno vam hvala za duboku veru u mene i bezuslovnu podrsku sve ove godine.
Paseta, hvala ti na razumevanju i strpljenju. Zajedno smo prosli ovaj ovaj ne bas laki put.
Hvala ti za nasu divnu decu.