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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

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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


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


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


Ethics statement ............................................................................................................88

Animals ..........................................................................................................................88

Experimental groups ......................................................................................................89

Experimental treatments and data recording ..................................................................89

Behavioral 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


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


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


°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


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


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


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.


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


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


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


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


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].


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].


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].


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.


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/

http://www.jove.com/video/3260/


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-


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.


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.


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


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.


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].


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.


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


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


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]


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

*

*

*

*


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).


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.


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


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


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,


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.


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


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


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


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


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


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.


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


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 postanesthetic days in both anesthetics. The alterations in body weight,

food and water intake were not statistically significant.


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


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


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]


HC

O3 [m

mo

l/L

iter]

0

5

10

15

20

25

30


Isoflurane

Sevoflurane

Normal


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

]


day 2day 1 day 3 day 4

Isoflurane

Sevoflurane

-100

-50

0

50

100

150

200 locomotor activity

**

*

*

anaesthesia


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


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


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


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


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)


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


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


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.


66


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


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


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


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


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.


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].


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.


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


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.


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


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)


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.


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


KS S-ketamine-sevoflurane

S sevoflurane

normal


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


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



pH

0

10

20

30

40

50

60

70

80

90

100



pO

2(m

m H

g)

110



S sevoflurane


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

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rt r

ate

(beats

per

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ute

)

c

ore

bo

dy t

em

pe

ratu

re

(°C

)



S sevoflurane

*

*

*

*

*

*

*

*180


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


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


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


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


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


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


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.


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’


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.


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


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,


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 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


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).


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

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(n

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., 1

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)

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- 4 - 2 0 2 4

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Function 1 (p ≤ 0.0001, 89%)

- 4 - 2 02

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C D

Surgery without analgesiaSurgery with analgesiaAnesthesiaControl

Function 1 (p= 0.0012, 78%)

total 18 hours 0 – 6 hours

6 – 12 hours 12 – 18hours


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


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).


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

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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

*

*

*

*

*

*

**

*

*

**

**

**

** *

**

*

*

**

*

* **

*


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.


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


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.


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.


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


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


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


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


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


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


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.

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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.

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