This first edition of the WHO guidelines on tularaemia is the 

result of an international collaboration, initiated at a WHO meeting 

in Bath, UK in 2003. The target audience includes clinicians, 

laboratory personnel, public health workers, veterinarians, and 

any other person with an interest in zoonoses. 

Tularaemia is a bacterial zoonotic disease of the northern 

hemisphere. The bacterium (Francisella tularensis) is highly 

virulent for humans and a range of animals such as rodents, hares 

and rabbits. Humans can infect themselves by direct contact with 

infected animals, by arthropod bites, by ingestion of contaminated 

water or food, or by inhalation of infective aerosols. There is 

no human-to-human transmission. In addition to its natural 

occurrence, F. tularensis evokes great concern as a potential 

bioterrorism agent. F. tularensis subspecies tularensis is one of 

the most infectious pathogens known in human medicine.

In order to avoid laboratory-associated infection, safety measures 

are needed and consequently, clinical laboratories do not 

generally accept specimens for culture. However, since clinical 

management of cases depends on early recognition, there is an 

urgent need for diagnostic services. 

The book provides background information on the disease, 

describes the current best practices for its diagnosis and 

treatment in humans, suggests measures to be taken in case of 

epidemics and provides guidance on how to handle F. tularensis

in the laboratory. 

isBn 978 92 4 154737 6
WHO Guidelines on
 

Tularaemia
 

EpidEmic and pandEmic 

alErt and rEsponsE 
WHO Library Cataloguing-in-Publication Data 

WHO Guidelines on Tularaemia. 

1.Francisella tularensis – classification. 2.Tularemia – epidemiology. 3.Tularemia 

– transmission. 4.Tularemia – drug therapy. I.World Health Organization.
 

ISBN 978 92 4 154737 6 (NLM classification: WC 380)
 

© World Health Organization 2007 WHO/CDS/EPR/2007.7 

All rights reserved. Publications of the World Health Organization can be obtained from WHO Press, World Health 

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e-mail: permissions@who.int). 

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Errors and omissions excepted, the names of proprietary products are distinguished by initial capital letters. 

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event shall the World Health Organization be liable for damages arising from its use. 

Printed in France 
 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Acknowledgments

Contents 

Acknowledgements	 v
 

Abbreviations	 vii
 

1.	 Introduction	 1
 

2.	 The	infectious	agent	 3
 

2.1 Taxonomy 3
 

2.2 Species and subspecies that differ in virulence and geographical range 4 


2.3 Seasonality, age and gender distribution 4 


3.	 Epidemiology	 5
 

3.1 Geographical distribution of tularaemia cases 5 


3.2 Vectors 5
 

3.3 Disease and pathology in animals 8 


4.	 Clinical	expression	in	humans	 11
 

4.1 Source of infection and clinical outcome according to F. tularensis subspecies 11
 

4.2 Signs and symptoms 12
 

4.3 Differential diagnoses 17
 

5.	 Treatment		 21
 

5.1 Recommendations for treatment and prophylaxis 21
 

5.2 Antibiotic agents 22
 

5.3 Antibiotic resistance of F. tularensis 25
 

6.	 Laboratory	diagnostics	and	discrimination	of	subspecies	and	strains	 27
 

6.1 Case definitions 27
 

6.2 Diagnostic specimens 27
 

6.3 Bacteriology 29
 

6.4 Diagnostic tests 30
 

6.5 Supplemental characterization 31
 

6.6 Biosafety considerations 33
 

7.	 Surveillance	and	outbreak	management	 35
 

7.1 Surveillance 35
 

7.2 Surveillance in humans 36
 

7.3 Surveillance in animals 36
 

7.4 Outbreak management (humans) 37
 

8.	 Considerations	for	handling F. tularensis	 41
 

8.1 Safety measures in the laboratory 41
 

8.2 Vaccines and vaccination 43
 

9.	 References	 47
 

Annexes	 	 59
 

A.	Diagnostic	protocols	 61
 

A.1 Gram stain 61
 

A.2 Culture on agar 62
 

A.3 Antigen detection 64
 

A.4 Molecular detection 70
 

A.5 Serology 73
 

A.6 References 

iii 
 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

wHo guidelines on tulArAemiA 

B.	 Tests	for	supplemental	characterization		 81 

B.1 Antimicrobial susceptibility 81 

B.2 Molecular biotyping 82 

B.3 References 87 

C.	Protocols	for	preparation	of	selected F. tularensis culture	media	 89 

C.1 Cysteine heart agar base with 9% chocolatized sheep blood (CHAB) 89 

C.2 GCII agar with 1% haemoglobin and 1% IsoVitaleX 90 

D.	Reagent	list	 93	 

E.	 Transport	of	specimens	and	cultures	of	F. tularensis		 95 

E.1 Shipping F. tularensis 95 

E.2 Training 95 

E.3 References 96 

F.	 Checklist	for	outbreak	investigation	 99 

F.1 Preparations 99 

F.2 Outbreak investigation in the field 100 

F.3 Example for a tularaemia outbreak, Kosovo (Serbia) 101 

G.	Protocols	for	trapping	and	sampling	small	mammals		 105 

G.1 Trapping of small mammals 105 

G.2 Protocols for collecting diagnostic samples from small mammals 105 

G.3 References 107 

Index	 	 	 109 

List	of	figures 

3.1 Geographical distribution of tularaemia 6 

4.1 Primary ulcer in a case of tularaemia 16 

4.2 Lymph node enlargement 16 

4.3 Hilar enlargement 16 

4.4 Extended right-sided consolidation 16 

5.1 Distribution of MIC values for six quinolones 26 

A.1 F. tularensis on CHAB agar after 48 h incubation 63 

A.2 Isolate of F. tularensis stained by the direct fluorescence method 65 

A.3 Slide agglutination for F. tularensis 67 

E.1 Flowchart for the classification of infectious substances and patient specimens 96 

List	of	tables 

3.1 Animals found to be infected with F. tularensis 10 

4.1 Clinical forms of tularaemia 13 

4.2 Differential diagnosis by route of acquisition 18 

5.1 Minimum inhibitory concentrations (MICs) 22 

6.1 Discriminating characteristics of Francisella species and subspecies 33 

6.2 Polymerase chain reaction (PCR) methods to discriminate F. tularensis subspecies 33 

C.1 Chemically-defined enrichment supplement for F. tularensis 91 

D.1 Diagnostic reagents for F. tularensis 93 

iv 
Acknowledgments

Acknowledgements
 

Chief	Editor Arne Tärnvik, Umeå University, Umeå, Sweden 

Secretariat Regula Leuenberger, World Health Organization, Geneva, 

Switzerland 

Chairs Roland Grunow, Robert Koch Institute, Berlin, Germany 

Jeannine Petersen, Centers for Disease Control and Prevention, Fort 

Collins, CO, USA 

Anders Sjöstedt, Umeå University, Umeå, Sweden 

Richard W. Titball, Defence Science and Technology Laboratory, 

Porton Down, Salisbury, Wiltshire SP4 0JQ, United Kingdom 

Other	contributors Pedro Anda, Instituto de Salud Carlos III, Madrid, Spain 

Tina Broman, Swedish Defence Research Agency, Umeå, Sweden 

May Chu, World Health Organization, Geneva, Switzerland 

Karen Elkins, Food and Drug Administration, Rockville Pike, MD, 

USA 

Mats Forsman, Swedish Defence Research Agency, Umeå, Sweden 

Anders Johansson, Umeå University, Umeå, Sweden 

Michael Kosoy, Centers for Disease Control and Prevention, Fort 

Collins, CO, USA 

Gérard Krause, Robert Koch Institute, Berlin, Germany 

Francis E. Nano, University of Victoria, Victoria, BC, Canada 

Andrew Pearson, Health Protection Agency, London, United 

Kingdom 

Note: All the contributors listed above are members of the informal 

WHO working group on tularaemia, established at the WHO 

meeting Improving public health preparedness for and response to the 

threat of epidemics: tularaemia network, Bath, United Kingdom, 14–15 

September 2003 (document: WHO/CDS/CSR/LYO/2004.6).1 

The authors wish to express their gratitude for the intellectual contributions and expert advice 

provided throughout the process of compiling these guidelines. Special support was given 

at all stages by May Chu, World Health Organization. At various stages, the following colleagues were involved: Alim Aikimbayev, Almaty, Kazakhstan; Karen Elkins, Rockville Pike, 

MD, USA; Art Friedlander, Frederick, MD, USA; Tatiana A. Gremyakova, Moscow, Russian 

Federation; Alec Macela, Hradec Kralove, Czech Republic; Paul Mead, Fort Collins, CO, USA; 

Irina Meschcheryakova, Moscow, Russian Federation; Torsten Mörner, Uppsala, Sweden; 

Marty Schreifer, Fort Collins, CO, USA; Martin Wale, Nottingham, United Kingdom. 

1	 All external experts/contributors have signed declarations of interest in accordance with WHO policy 

and no conflict of interests were declared. The declarations of interest are available on request. 

v 
wHo guidelines on tulArAemiA 

Expert review of the document was performed by Alim Aikimbayev, Pedro Anda, Erik 

Bäck, Tina Broman, Mats Forsman, Tatiana A. Gremyakova, Zdenek Hubálek, Richard F. 

Jacobs, Anders Johansson, Aynur Karadenizli, Gérard Krause, Michael Kosoy, Alec Macela, 

Gunnar Sandström, and Sam R. Telford. 

The authors wish to acknowledge the following colleagues at the Department of Epidemic 

and Pandemic Alert and Response, World Health Organization, Geneva, Switzerland, for 

their constructive collaboration on these guidelines: Ottorino Cosivi, Philippe Dubois, Bradford Kay, and Stefano Lazzari. 

Excellent secretarial work was performed for the initial period by Williamina Wilson and 

Eglé Lorenzin, both at the World Health Organization, Geneva, Switzerland. 

WHO gratefully acknowledges the financial support of the Centers for Disease Control 

and Prevention, USA, the Humanitarian Aid Agency of the Federal Department of Foreign 

Affairs, Switzerland, and the Department of International Cooperation, Ministry of Foreign 

Affairs, Italy. 

We deeply regret the untimely death of Dr Regula Leuenberger. She was instrumental 

in ensuring the successful completion of this project and her dedication and commitment to her work will be greatly missed by her colleagues. 

vi 
Acknowledgments

Abbreviations
 

BCYE	 buffered charcoal yeast extract 

BSA	 bovine serum albumin 

CA	 enriched chocolate agar 

CFU	 colony-forming unit 

DNA	 deoxyribonucleic acid 

ELISA	 enzyme-linked immunosorbent assay 

FITC	 fluorescein isothiocyanate 

IHR	 International Health Regulations 

LD50	 The amount of a material, given all at once, which causes the death of 50% of a 

group of test animals 

LPS	 lipopolysaccharide 

LVS	 live vaccine strain 

MIC	 minimum inhibitory concentration 

MBC	 minimum bactericidal concentration 

OD	 optical density 

OIE	 World Organisation for Animal Health 

PBS	 phosphate-buffered saline 

PCR	 polymerase chain reaction 

rpm	 revolutions per minute 

RT	 room temperature 

SBA	 sheep blood agar 

TGBA	 thioglycollate-glucose-blood agar 

Type A	 F. tularensis subspecies tularensis (Jellison type A) 

Type B	 F. tularensis subspecies holarctica (Jellison type B) 

WHO	 World Health Organization 

vii 

wHo guidelines on tulArAemiA

1.	 Introduction 

Arne tärnvik 

Tularaemia is a bacterial zoonotic disease of the northern hemisphere. The bacterium (Francisella tularensis) is highly virulent for humans and a range of animals such as rodents, hares 

and rabbits. It may cause epidemics and epizootics. F. tularensis is transmitted to humans (i) 

by arthropod bites, (ii) by direct contact with infected animals, infectious animal tissues or 

fluids, (iii) by ingestion of contaminated water or food, or (iv) by inhalation of infective aerosols. There is no human-to-human transmission. 

Tularaemia is reported from most countries in the northern hemisphere, although its 

occurrence varies widely from one region to another. In some countries, endemic regions 

with frequent outbreaks are close to regions that are completely free of tularaemia. There is 

also a wide variation with time. In an endemic area, tularaemia may occur annually within a 

5-year period, but may also be absent for more than a decade. The reasons for this temporal 

variation in the occurrence of outbreaks are not well understood. When, after a long lapse, the 

first case of a new outbreak appears, the disease may be more or less forgotten and is therefore 

not easily diagnosed. 

F. tularensis subspecies tularensis (type A) is one of the most infectious pathogens known 

in human medicine. The infective dose in humans is extremely low: 10 bacteria when injected 

subcutaneously and 25 when given as an aerosol (McCrumb, 1961; Saslaw et al., 1961a; Saslaw 

et al., 1961b). For example, on Martha’s Vineyard in the United States of America, two adolescents contracted respiratory tularaemia after mowing a grassed area (Feldman et al., 2001). It 

is believed that an aerosol of F. tularensis subspecies tularensis was generated after the carcass 

of a rabbit which had died of tularaemia was accidentally shredded by the lawnmower. 

The risk posed by tularaemia can be properly managed, provided the public health system is well prepared. In order to avoid laboratory-associated infection, safety measures are 

needed and consequently clinical laboratories do not generally accept specimens for culture. 

However, since clinical management of cases depends on early recognition, there is an urgent 

need for diagnostic services. In addition to its natural occurrence, F. tularensis causes great 

concern as a potential bioterrorism agent. 

The present guidelines on tularaemia (i) provide background information on the disease, 

(ii) describe the current best practices for its diagnosis and treatment in humans, (iii) suggest measures to be taken in case of epidemics and (iv) provide guidance on how to handle 

F. tularensis in the laboratory. The target groups for these guidelines include clinicians, laboratory personnel, public health workers, veterinarians, and any other person with an interest 

in zoonoses. 

The guidelines are the result of an international collaboration, initiated at a WHO meeting 

in Bath, United Kingdom of Great Britain and Northern Ireland in 2003, continued in Umeå, 

Sweden, in 2004 and finalized in Geneva, Switzerland, in 2005. 

Each chapter of the guidelines was developed by a selected group of scientists with extensive developmental work and general experience relevant to the field covered. Chairs of each 

group met regularly throughout the process to ensure consistency and for critical review. The 

document was also subject to internal external peer reviews. 

As with many other areas, understanding of the nature of tularaemia and its causative 

agent is evolving rapidly across all aspects discussed in the guidelines, including taxonomy, 

epidemiology, epizootiology, detection, diagnostics, therapy and prophylaxis. It is envisaged 

therefore that modifications to these guidelines will become necessary every three years. 

1
 

 

 

 

2. tHe inFectious Agent

2.	 The	infectious	agent 

tina Broman, mats Forsman, Jeannine Petersen, Anders sjöstedt 

Historically, tularaemia was a public health problem in the former Soviet Union and in the 

USA. For example, in the USA some 1400 cases were reported from 1990 to 2000, as compared 

to more than 14 000 cases from 1920 to 1945 (Hayes et al., 2002). In the former Soviet Union, 

extensive outbreaks occurred during and after the Second World War. During the winter of 

1941–42, 67 000 cases were reported from the region surrounding Rostov-on-Don (Jusatz, 

1952). There are no data available that explain the decline in cases since the 1950s. However, 

it is assumed that this may be due to less-frequent exposure of humans to rodents, rabbits and 

hares which in turn may be related to a decrease in the number of hunters and a decrease in 

the percentage of the population living in rural settings. In the Russian Federation, vaccination programmes may have contributed to the decline. Experiences from the former Soviet 

Union during the Second World War and from Bosnia and Herzegovina, and United Nations 

Administered Province of Kosovo (Serbia) during the recent civil wars suggest that tularaemia can increase significantly during and after warlike conditions, or after natural disasters 

that disrupt the normal hygiene and sanitary conditions of a society (Jusatz, 1952; Reintjes et 

al., 2002). There are indications that the epidemiology of the disease may be changing. In Canada, for example, contact with rabbits was the most common source of infection before the 1950s, 

while contact with muskrats appears to be of greater importance today (Martin et al., 1982). 

Francisella tularensis, the infectious agent of tularaemia, is a potential bioterrorist agent 

(Dennis et al., 2001). During the Cold War, the pathogen was possibly developed into biological warfare weapons in both the east and the west (references in World Health Organization, 2004a). The pathogen is attractive for these developments because of its very high 

infectivity and relative stability in aerosols which facilitates its dissemination. 

On solid culture media, bacterial colonies become visible within 2–3 days of incubation at 

37 °C and a preliminary confirmation can be easily and rapidly obtained by agglutination of 

the bacteria in immune serum. Due to scientific achievements of the past decade, molecular 

techniques are now available for rapid and final identification of F. tularensis, and for differentiation of subspecies. 

2.1	 Taxonomy 

F. tularensis is not closely phylogenetically related to any pathogen or commensal of the 

human flora. It belongs to a group of intracellular bacteria, which includes mycobacteria Listeria, Legionella, Brucella, Coxiella and Rickettsia. 

Francisella tularensis is one of two species in the genus Francisella, which is the only genus 

of the family Francisellaceae, a member of the gamma-subclass of Proteobacteria (Sjöstedt, 

2005). The two species, F. tularensis and F. philomiragia, show a 16S rRNA sequence similarity 

of ≥98.3% (Forsman, Sandström & Sjöstedt, 1994). The family is distinguished by a unique set 

of phenotypic characteristics including a coccoidal morphology, Gram-negativity, a capability 

to degrade only a limited number of carbohydrates resulting in acid but no gas production, a 

growth requirement for cysteine, and a unique fatty acid composition (Sjöstedt, 2005). 

Based on 16S rRNA gene sequence analysis, several tick endosymbionts are closely related 

to F. tularensis and have been preliminarily placed in the Francisella group (Sun et al., 2000; 

Scoles, 2004). Caedibacter taeniospiralis, a protozoan endosymbiont also shows 16S rRNA similarity to members of Francisellaceae (Beier et al., 2002). In addition, several potential new 

Francisella species, including several with high 16S rRNA similarity to F. tularensis, have been 

identified in soil samples in the Houston area of the USA (Barns et al., 2005). 

3
 
 

 

wHo guidelines on tulArAemiA 

2.2	Species	and	subspecies	which	differ	in	virulence		 

and	geographical	range 

With the recent reassignment of Francisella novicida as a subspecies of F. tularensis, there are 

currently four proposed subspecies, each of which displays several distinct biochemical, epidemiological, and virulence characteristics (Jellison, 1974; Sjöstedt, 2005). The 16S rRNA 

similarity is ≥99.8% among the subspecies (Forsman, Sandström & Sjöstedt, 1994). 

The highly virulent F. tularensis tularensis (type A) is found in the USA, whereas the less 

virulent F. tularensis holarctica (type B) is found throughout the northern hemisphere. Subspecies are further discriminated into isolates and strains. An isolate is a population of bacterial 

cells in pure culture derived from a single colony on a primary isolation plate. In the case of 

F. tularensis and other fastidious organisms forming small-size colonies this definition is often 

violated. Several colonies are unavoidably sampled on a primary isolation plate. A strain is an 

isolate or group of isolates displaying specific genetic or phenotypic characteristics that set it 

apart from other isolates of the same species. 

Strains of subspecies mediasiatica have been isolated only in Kazakhstan and Turkmenistan. Little work is known regarding these strains but experimental studies in rabbits have 

indicated a moderate virulence comparable to that of subspecies holarctica. 

Subspecies novicida shows a low virulence in experimental models and causes disease in 

immunocompromised individuals (Hollis et al., 1989) but it has not been isolated from any 

animals. However, isolates of subspecies novicida have been linked to waterborne transmission in Australia, Spain and the USA (Hollis et al., 1989; Whipp et al., 2003). The isolate from 

Australia is the only F. tularensis strain originating from the southern hemisphere to date. 

F. philomiragia is associated with salty water (Atlantic as well as Mediterranean) and is of 

low virulence, although it may cause disease in immunocompromised individuals. See section 4.1 for further information on the mode of transmission and clinical symptoms. 

2.3	Seasonality,	age	and	gender	distribution 

Although the age-related incidence rate of tularaemia is unknown, the disease is known to 

occur at all ages. Males have a higher incidence in all age categories. In Sweden, an overrepresentation among males has been attributed to their more frequent outdoor professional and 

leisure activities (Eliasson et al., 2002). In most countries where tularaemia is endemic, the 

disease is seasonal; its incidence seems to be highest during late spring, the summer months 

and early autumn (Olsufiev, 1977; Hayes et al., 2002; Tärnvik et al., 2004). Often, the number 

of cases shows wide variations from one year to another and this is probably related to climatic factors such as temperature and precipitation. However, there are virtually no data linking 

specific climatic conditions and outbreaks of tularaemia. This is an important area for future 

research that may yield important tools for predicting and possibly preventing outbreaks. 

4
 
3. ePidemiology

3.	 Epidemiology 

Anders sjöstedt 

There is no evidence for human-to-human transmission of tularaemia. F. tularensis has been 

isolated from a variety of animals which may function as vectors for transmission to humans 

(Table 3.1). Transmission of F. tularensis subspecies tularensis is associated with rabbits, ticks 

and sheep, whereas F. tularensis subspecies holarctica is often isolated in association with 

streams, ponds, lakes and rivers. Beavers and muskrats in North America and lemmings and 

beavers in Scandinavia might play a role in maintaining the water association of the bacterium (Parker et al., 1951; Hopla, 1974; Hörnfeldt, Löfgren & Carlsson, 1986; Mörner et al., 

1988a). There is also evidence that the bacterium can persist for months in watercourses, 

possibly in association with protozoa (Berdal et al., 1996). For further references see section 

7.4.1. It should be noted, however, that the epizootology of tularaemia is not yet fully understood (Hopla, 1974). For instance, the respective importance of various susceptible animals 

as reservoirs for F. tularensis is still poorly known. Similarly, the role of various arthropods in 

the transmission of F. tularensis among animals and between animals and humans is not yet 

well understood. 

3.1	 Geographical	distribution	of	tularaemia	cases 

Tularaemia has been reported in many countries of the northern hemisphere, but so far 

not from the southern hemisphere (Figure 3.1). An isolate of subspecies novicida has been 

found in Australia, but without the typical clinical presentation of tularaemia (Whipp et 

al., 2003). Endemic foci have existed for a long time in the Russian Federation, Kazakhstan 

and Turkmenistan (Olsufiev, 1977) as well as Finland and Sweden (Tärnvik, Priebe & Grunow, 2004). Annual cases are reported from most countries in eastern Europe whereas there 

are few reported cases in countries of continental western Europe. Outbreaks comprising 

hundreds of cases have occurred recently in Portugal, Spain, Sweden and UN Administered Province of Kosovo (Serbia) (de Cavalho et al., 2007; Pérez-Castrillòn et al., 2001; 

Eliasson et al., 2002; Reintjes et al., 2002). Cases are also reported in Japan and the northwestern and north-eastern regions of China. In the western hemisphere, cases regularly occur in the USA and Canada and a few cases have been reported from Mexico. In the 

USA, some 120 cases have been reported annually from 1990 to 2000 (Hayes et al., 2002). 

More than 50% of these cases occurred in Arkansas, Missouri, South Dakota and Oklahoma. Small outbreaks have been reported repeatedly from the island Martha’s Vineyard in 

Massachusetts (Feldman et al., 2001; Feldman et al., 2003). 

3.2	Vectors 

3.2.1	 Arthropods 

A wide range of arthropods has been identified in the transmission of tularaemia between 

mammalian hosts. In the USA, ticks are considered as the most important vectors east of the 

Rocky Mountains, where F. tularensis has been detected in at least 13 species representing 

four genera: Amblyomma, Dermacentor, Haemaphysalis, and Ixodes (Hopla, 1974). Flies (Chrysops spp.) are the most common vectors in Utah, Nevada, and California. Several outbreaks 

of human tularaemia in Utah have been linked to transmission by Chrysops discalis (Jellison, 

1950). In the former Soviet Union, the bacterium has been demonstrated to be transmitted by 

mosquitoes (Aedes spp., Culex spp., Anopheles spp.) and ticks (Ixodes spp.). However, in ticks 

5
 
wHo guidelines on tulArAemiA 

Figure	 3.1	 The geographical distribution of tularaemia is indicated in grey. Locally, the disease often has an 

uneven distribution with recurrent epidemic outbreaks in geographically restricted areas. The data were compiled from publications in the medical literature between 1952–2006. Credit: Anders Johansson, Umeå University, 

Umeå, Sweden and from a recent publication (de Carvalho et al., 2007) 

the prevalence appeared to be low; an investigation of 120 000 adult Ixodes ricinus in north

western areas of the former Soviet Union between 1960 and 1964 resulted in identification of 

0.01% positive ticks (Olsufiev & Dunayeva, 1970). In central Europe, the ticks Dermacentor 

reticulatus and Ixodes ricinus are important vectors. In areas of Austria and the Czech Republic, 

2% of D. reticulatus ticks contained F. tularensis (Hubalek et al., 1997). In Finland and Sweden, 

clinical experience and epidemiological data support the role of mosquitoes as vectors for 

transmission of F. tularensis to humans (Christenson, 1984; Eliasson et al., 2002). In Sweden, 

mosquitoes of the species A. cinereus have been demonstrated to be infected (Olin,1942). Data 

reported both from the former Soviet Union and the USA show that flies of the family Taba

nidae may serve as vectors for the disease (Krinsky, 1976). This family includes true horse

flies (Tabanus spp. and Chrysozona spp.) and deer flies (Chrysops spp.). These flies are found 

in nearly all parts of the world and females of all species are blood-sucking. F. tularensis has 

been detected in at least 20 flea species of eight genera (Amphipsylla, Cediopsylla, Ceratophyllus, 

Ctenophthalmus, Malaracus, Megabothris, Neopsylla, and Pulex) but their role in the spread of 

­

­

­

F. tularensis is unclear (Olsufiev & Dunayeva, 1970). 

Although the exact mechanism of transmission of tularaemia by flies and mosquitoes is 

still elusive, it should be noted that F. tularensis has never been demonstrated to reside in 

the salivary glands of any species. Maybe spread is simply mechanical, by contamination of 

mouthparts when a fly or a mosquito is biting an infected host. Alternatively, F. tularensis may 

be mechanically rubbed into the skin when the arthropod is swatted. Another source of infec

tion of flies or mosquitoes has been suggested to be contaminated water (Pavlovsky, 1966). 

3.2.2	 Mammals 

Tularaemia outbreaks have been described in hares (Mörner et al., 1988), prairie dogs (Petersen 

et al., 2004a), and mink (Henson, Gorham & Shen, 1978). It should be noted that although 

some studies suggest that mammals harbour and secrete the bacteria for longer periods of 

time (Bell, 1980), there is no evidence that they constitute a major natural reservoir for F. tu­

­

6
 
3. ePidemiology 

larensis. Nevertheless, F. tularensis is thought to be spread in the environment principally by 

various terrestrial and aquatic mammals such as ground squirrels, rabbits, hares, beavers, 

muskrats and, in particular, rodents such as meadow voles and water voles (Table 3.1). Systematic studies in the former Soviet Union revealed that among these animals, voles and 

mice were presumed to be the main source for the spread of F. tularensis to humans (Olsufiev, 

1977). In particular, the most commonly-identified vectors were the water vole (Arvicola terrestris), the common vole (Microtus arvalis), the red-backed vole (Clethrionomys spp.) and the 

domestic mouse (Mus musculus). The role of non-rodent species such as rabbits and hares in 

the spread of tularaemia to other mammals and humans is not well known but it has been 

suggested that they may infect hunters in central Europe (Pfahler-Jung, 1989). In certain parts 

of the USA, transmission via ticks from the rabbit is an important cycle for the spread of 

F. tularensis subspecies tularensis to other mammals and humans (Taylor et al., 1991). 

3.2.3	 Rodent	population	size	and	transmission	to	humans 

Outbreaks of the disease in humans often follow outbreaks of tularaemia in rodents. For example, outbreaks in humans in the Russian Federation were shown to be linked to epizootics of 

the disease in common voles (Mörner, 1992). In regions where tularaemia is endemic, antibodies to F. tularensis are frequently detected in the sera of trapped wild animals such as 

muskrats, beavers and common voles (Mörner, Sandström & Mattsson, 1988). 

Widespread epizootics of tularaemia have also been associated with a preceding increase 

in the density of the rodent population size. The results from a recently published investigation showed that the number of human cases of tularaemia in the Novosibirsk region of the 

Russian Federation from 1956–2000 was correlated to the density of the water rat population 

(Efimov, Galaktionov & Galaktionova, 2003). In Sweden, a strong correlation between peaks 

in vole and hare populations and outbreaks of tularaemia in humans has been reported during the 1960s and 1970s (Hörnfeldt, Löfgren & Carlsson, 1986). However, in recent years, no 

correlation with the size of the rodent populations was found in Sweden (Tärnvik, Sandström 

& Sjöstedt, 1996). 

Tularaemia transmission patterns may also change over time. In Canada, for example, contact with rabbits was the most common route of spread of infection before the 1950s while 

more recently, the water-living muskrat appears to be of greater importance (Martin et al., 

1982). 

3.2.4	 Reservoirs	in	the	environment 

Faeces from the field vole (Microtus agrestis) were demonstrated to be a source of human 

infection in the 1960s in Sweden (Dahlstrand, Ringertz & Zetterberg, 1971). Hay stored in 

barns was contaminated with the faeces and humans contracted tularaemia in the barns via 

inhalation. On Martha’s Vineyard in the USA, two fatal cases of tularaemia were associated 

with grass mowing and exposure by aerosol formation to the carcass of an infected rabbit 

(Feldman et al., 2001). 

In a few instances, it has been reported that F. tularensis has been isolated from mud (Parker et al., 1951). However, there are no recent reports on the isolation of the bacterium from 

mud or soil. It is unclear whether this means that the bacterium rarely exists in mud or soil 

or whether it is difficult to cultivate F. tularensis from such sample types. Nevertheless, 16S 

rRNA sequences highly similar to those previously reported for F. tularensis were recently 

found in soil samples in Houston, Texas, USA (Barns et al., 2005). Although no bacteria were 

cultivated, the gene sequences indicated that they were derived from bacteria that belong 

to the family Francisellaceae but were distinct from known species and formed several new 

clades potentially representing new species or genera. These findings will allow comparative 

analyses of samples from other geographical locations. 

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3.3	Disease	and	pathology	in	animals 

Signs and symptoms of tularaemia in wild animals are not well documented and are mostly 

based on postmortem examinations. The most common finding upon necropsy is an enlarged 

spleen and pinpoint white necrotic lesions in the spleen and liver (Bell, 1980; Hopla & Hopla, 

1994). The manifestations are likely to be dependent upon the susceptibility of the particular 

animal species to tularaemia. The best-documented clinical cases are in domestic cats and 

dogs, captive monkeys, prairie dogs and laboratory animals. 

3.3.1	 Case	descriptions	in	naturally-infected	animals 

Tularaemia has been diagnosed in two cats that were examined because of pyrexia and lethargy; both had a history of exposure to wild rabbits (Woods et al., 1998). One cat was vomiting 

and the other was anorexic. Physical examination revealed dehydration, lymphadenopathy 

and hepatomegaly. Acute F. tularensis infection in three domestic cats was associated with 

marked signs of depression, oral/lingual ulceration and regional or generalized lymphadenomegaly (Baldwin et al., 1991). 

One case of tularaemia in a dog was characterized by acute onset of anorexia, pyrexia, 

lymphadenopathy and tonsillitis (Gustafson & DeBowes, 1996). The disease was self-limiting 

and clinical signs resolved within five days with only supportive therapy. A second case demonstrated acute onset of lethargy, anorexia and weakness (Meinkoth, Morton & Meinkoth, 

2004). The dog had eaten an adult rabbit 36 h earlier. 

In a Canadian zoological garden, three black and red tamarins (Sanguinus nigricollis) and 

one talopoin (Cercopithecus talapoin) died of tularaemia. They collapsed and died quickly 

after showing few symptoms. A second talapoin developed abscesses in the tongue and submandibular area and recovered after treatment (Nayer, Crawshaw & Neufeld, 1979). A golden-headed lion tamarin (Leontopithecus chrysomelas) demonstrated profuse salivation, ocular 

and nasal discharge and severe tongue ulcerations (Hoelzle et al., 2004). 

Prairie dogs affected during an outbreak showed rapid deterioration of their status and 

within two days developed dehydration, ataxia, and severe diarrhoea (La Regina, Lonigro & 

Wallace, 1986). In an outbreak of tularaemia among captive prairie dogs, unusual lethargy, 

severe dehydration, and grossly enlarged cervical lymph nodes were characteristic clinical 

features (Petersen et al., 2004a). 

3.3.2	 Course	of	disease	in	laboratory	animals 

A number of laboratory animals, such as guinea-pigs, rabbits, rats, mice and monkeys, have 

been used in studies of the course of experimental tularaemia. Experimental work showed 

guinea-pigs to be highly susceptible to subcutaneous, intraperitoneal, and intranasal challenge with strain SCHU S4 of subspecies tularensis (Downs et al., 1947). Rats appear to be 

much more resistant to F. tularensis than other commonly used experimental animals. The 

subcutaneous and intradermal LD50 of strain SCHU S4 for rats was found to be at least 

10 000-fold higher than for mice, rabbits or guinea-pigs (Downs et al., 1947). After an intraperitoneal challenge with SCHU S4, rats developed pyogranulomatous, necrotic hepatitis 

within 24 h (Moe et al., 1975). Lesions containing mostly macrophages developed within the 

red pulp in the spleen. 

Rabbits 

Extensive experimental work on tularaemia has been performed in rabbits. Aerogenic exposure 

resulted in necrosis of the bronchi by day 2 (Baskerville & Hambleton, 1976). Alveoli were 

distended by neutrophils and necrotic alveolar epithelial cells and the cervical and bronchial 

lymph nodes were enlarged. Necrotic foci were found in the liver and the white pulp of the 

spleen. By day 3, necrotic foci had increased considerably in size in lung, spleen and liver, and 

cervical and bronchial lymph nodes were necrotic. 

8
 
3. ePidemiology 

Mice 

Mice have been widely utilized in the experimental model of tularaemia. Generally, few differences in susceptibility among various inbred strains to challenge with the live vaccine 

strain (LVS) or strain SCHU S4 have been detected (Anthony, Skamene & Kongshavn, 

1988; Shen, Chen & Conlan, 2004). Studies before 1970 mostly used outbred mice whereas 

a number of studies employing inbred mice have been published after 2002. All available 

mouse strains appear to be highly susceptible to subcutaneous or aerosol infection with low 

doses of subspecies tularensis or holarctica and death occurs within one week at most (Shen, 

Chen & Conlan, 2004). Conlan and colleagues (2003) found that histopathological changes 

in the lung were inconspicuous and only small areas of necrosis were visible. Changes in the 

spleen were more marked and, by day 5, showed almost complete loss of follicle structures 

and abundant necrotic areas. Liver changes were observed within one day of infection and 

were characterized by inflammatory necrotic foci. These increased by day 4 with vast areas 

of necrosis but very little inflammatory response. Many of these changes were similar for 

subspecies holarctica and subspecies tularensis (Conlan et al., 2003). Overall, it appeared that 

the histological findings did not distinguish an infection caused by subspecies holarctica from 

that caused by subspecies tularensis. 

Monkeys 

Historically, monkeys have been used to study both the pathogenesis of tularaemia and to 

evaluate vaccines. Grivet monkeys administered strain SCHU S4 intranasally became moribund from day 4 (Baskerville, Hambleton & Dowsett, 1978). Most of the pulmonary infection 

was confined to the upper respiratory tract as evidenced by copious mucopurulent oculonasal discharge. On histological examination, all monkeys developed extensive necrotizing 

rhinitis and pharyngitis. The lower airways only became infected on day 6 after administration. Concomitantly, regional lymph nodes, spleen and liver became heavily infected. In the 

liver, numerous macroscopic surface lesions were present by day 4. These consisted mostly 

of necrotic hepatocytes and Kupffer cells. By day 7, granulomas were beginning to develop at 

foci of liver infection. In the spleen extensive necrosis of the white and red pulp occurred. 

Two studies examined tularaemia in Rhesus macaques challenged by an aerosol of a strain 

of subspecies holarctica (Schricker et al., 1972; Hall, Kovatch & Schricker, 1973). Mortality 

(3–18%) correlated to the number of bacteria inoculated (103–106 CFU). Infectious foci in the 

lungs showed mixed infiltrates of polymorphs, lymphocytes and histiocytes and progressed 

within 6 to 21 days to focal liquefactive necrosis followed by granuloma development. Hepatic 

changes consisted of circumscribed areas of mononuclear infiltrates, accumulation of mixed 

inflammatory cells in the sinusoids and enlargement of Kupffer cells. Splenomegaly was 

observed during the symptomatic period. Congestion of the red pulp and neutrophil accumulation in the sinuses were noted. Thus, tularaemia in monkeys has more similarities to 

tularaemia in humans than in rodents. See also section 8.2. 

3.3.3	 Serological	evidence	of	tularaemia	in	wild	animals 

Seroconversion has been documented in free-living animals, primarily in carnivores such 

as kit foxes and coyotes (McCue & O’Farrell, 1988; Gese et al., 1997). Other animals which 

may survive infection and develop immunity also may demonstrate seroconversion (class 

II in Olsufiev & Dunayeva, 1970; Table 3.1). For example, 21% of the investigated sera from 

Scandinavian beaver showed increased antibody titres to F. tularensis (Mörner, Sandström 

& Mattsson, 1988). It is likely that seroconversion is indicative of infection with subspecies 

holarctica since subspecies tularensis in most animal species causes a uniformly fatal disease. 

9
 
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wHo guidelines on tulArAemiA 

Animals found to be infected with Table	3.1 

F. tularensis: their sensitivity to tularaemia and epizootiological significance (modified from Olsufiev and 

Animal	species	 Dunayeva, 1970) 

Geographical	distribution	 Potentiala	 References 

Muskrat (RODENTS
 

Ondatra zibethica) North America, Europe and northern Asia Class 1 Dunaeva & Emel’Ianova, 1950; Parker et al., 1951;
 

Young et al., 1969 

Common vole (Microtus arvalis) Central Europe Class 1 Nekrasov et al., 1985 Water vole (Arvicola terrestris) Northern, central and eastern Europe Class 1 Tropin, 1969; Bell & Stewart, 1975; Nekrasov et al., 

1985 

Red-backed vole (Clethrionomys glareolus) Central Europe Gurycova et al., 2001 Domestic mouse (Mus musculus) Worldwide Nekrasov et al., 1985 Mice (Mus; Micromys; some Apodemus spp.) Worldwide Class 1 Gurycova, 2001 et al.; Nekrasov et al., 1985 Lemmings (Lemmus spp.) Alaska; northern Scandinavia Class 1 Algazin & Bogdanov, 1978; Berdal et al., 2000 Rats (Rattus spp.) Worldwide Class 2 Feldman et al., 2001; Nekrasov et al., 1985 Dormouse (Dryomys nitedula) Central, southern and eastern Europe Class 2 Friedl, Heinzer & Fankhauser, 2005 True beaver (Castor fiber) Europe; northern Asia Class 3 Mörner, Sandström & Mattsson, 1988 American beaver (Castor canadensis) North America Class 2 Bell et al., 1962; Parker et al., 1951 Field mouse (Apodemus agrarius) Eastern Europe and Asia Class 2 Gurycova et al., 2001; Nekrasov et al., 1985 Hamsters (Cricetulus; Cricetus; Central Asia Class 1 MMWR, 2005; Baitanaev & Aikimbaev, 1977 

Mesocricetus; Phodopus spp.) 

Cottontail rabbits (

LAGOMORPHS 

Sylvilagus spp.) North America Class 1 Woolf, Shoemaker & Cooper, 1993 

Hares (Lepus spp.) Worldwide Class 1 Cerny, 2001 

Jack-rabbits, black-tailed hare, white-tailed hare California, USA Class 1 Simons, Stevens & Reeves, 1953 

Red-toothed shrews (

INSECTIVORES 

Sorex spp.) North America; Europe and northern Asia. Class 1 Gurycova et al., 2001; Khlyap, 1989 

Hedgehog (Erinaceus spp.) Europe; Asia Class 2 Farhang-Azad, Mescerjakova & Neronov, 1973 Steppe hedgehogs (Hemiechinus spp.) Mediterranean; Cyprus; Asia Minor – south-east Asia. Class 2 Difference vs Erinaceus Water shrews (Neomys spp.) Europe; Asia Class 2 Mikhailovskii, Litvinova & Sadovnikova, 1981 Moles (Talpa spp.) Europe; Japan Class 1 Fujita, Takada & Ohara, 1985 

a 

after inoculation of 10Potential sensitivity of animal species to tularaemia classified as: class 1: acute disease after inoculation of 1–10 bacteria; rapid multiplication in blood and tissues; class 2: death 

8–109 bacteria; survival may occur at lower doses and then provide immunity; in the article by Olsufiev (Olsufiev and Dunayeva, 1970), it was suggested that 

a class 3 also exists, comprising genera resistant to F. tularensis; Canis, Felis, Meles, Nyctereutes, Vulpes, Mustela. Since there exists no experimental data on these genera, they have not been included in the Table. 
4. clinicAl exPression in HumAns

 

 

 

 

 

 

 

 

 

4.	 Clinical	expression	in	humans 

Pedro Anda, Andrew Pearson, Arne tärnvik 

Tularaemia is transmitted to humans by arthropod bites (ticks, flies, mosquitoes), by direct 

contact with infected animals, by contact with tissues or fluids from infectious animals, by 

ingestion of contaminated water or food, or by inhalation of infective aerosols. There is no 

human-to-human transmission. Early diagnosis and treatment may be difficult in regions 

where tularaemia occurs rarely because the clinical expression of the disease is similar to a 

wide variety of acute infectious diseases. After acquisition, the bacteria replicate in lymphoid 

tissues of the host. In the acute phase of disease, the bacteria multiply rapidly. The severity 

of the disease depends on a capability of the patient to mobilize an immunological response. 

This response depends on cell-mediated immunity rather than on antibodies. Antibodies are 

good indicators of exposure to the bacterium but do not play a crucial role in protection. See 

also section 8.2. The mean incubation period of tularaemia is 3–5 days, but may range from 

1–21 days. The onset of disease is abrupt, including rapid development of fever with chills, 

fatigue, general body aches and headache. This is the same for type A and type B tularaemia. 

4.1	 Source	of	infection	and	clinical	outcome	according	to		 

F. tularensis	subspecies 

4.1.1	 Subspecies	tularensis	(type	A) 

F. tularensis subspecies tularensis occurs in the USA. It is one of the most infectious pathogens known in human medicine. The bacterium is transmitted among animals and from 

animals to humans by ticks and occasionally deerfly bites, or by aerosols. Human-to-human 

transmission by arthropods has never been described. Humans may also become infected by 

handling infectious animals. 

Subspecies tularensis can cause serious clinical manifestations and significant mortality if 

untreated (Dienst, 1963). It may occasionally lead to rhabdomyolysis and septic shock (Kaiser 

et al., 1985; Provenza, Klotz & Penn, 1986; Klotz, Penn & Provenza, 1987). Before the advent 

of antibiotics, the fatality rate of type A tularaemia was 5–15% and in its most severe forms 

as high as 30–60%. Currently, it is less than 2% (Dennis et al., 2001). High fever is accompanied by progressive weakness, malaise, anorexia and loss of weight. Respiratory symptoms, 

including a dry cough, a sore throat and substernal pain may occur whether or not the disease 

is acquired by inhalation. Gastrointestinal symptoms such as nausea, vomiting and diarrhoea 

are more likely to occur in type A than in type B tularaemia. 

4.1.2	 Subspecies	holarctica	(type	B) 

Type B tularaemia occurs in Eurasia and in North America. In contrast to type A, type B 

is mainly associated with streams, ponds, lakes, rivers, and semi-aquatic animals such as 

muskrats and beavers. Therefore, some authors consider type B tularaemia to be a waterborne 

disease. In favour of this, subspecies holarctica has recently been shown to survive and replicate in protozoa (Abd et al., 2003). Nevertheless, subspecies holarctica has been found also 

in hares and other animals. The bacterium is transmitted to humans by direct contact with 

infectious animals, arthropod bites, aerosols, or ingestion of contaminated food or water. 

Type B tularaemia is much less severe than type A tularaemia and fatal cases are rare. In type 

B tularaemia, fever predominates and is accompanied by focal symptoms and generally milder 

symptoms than those of type A tularaemia. However, it is frequently associated with suppurative complications (see section 4.2.9) and requires a considerable period of convalescence. 

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Worldwide, the prevalence of disease caused by type B is much higher than that caused by 

type A. An increase in prevalence of type B tularaemia has been observed during times of 

war. During the Second World War, at least 100 000 cases occurred each year. More recently, 

war-associated outbreaks occurred in UN Administered Province of Kosovo (Serbia) in 2000 

and 2003, each comprising more than 300 cases (Reintjes et al., 2002). In contrast to the large 

Second World War outbreaks in the former Soviet Union, the UN Administered Province of 

Kosovo (Serbia) outbreaks occurred in areas that were not recognized as tularaemia-endemic. A possible intentional release was therefore suspected; however this was not indicated by 

a recently developed model to evaluate such suspicions based on scores for a variety of social, 

political, and clinical criteria (Grunow & Finke, 2002). 

4.1.3	 Subspecies	mediasiatica 

F. tularensis subspecies mediasiatica is only rarely reported as a cause of human disease. With 

regard to virulence, it relates more to subspecies holarctica than to subspecies tularensis (Olsufjev & Meshcheryakova, 1982). 

4.1.4	 Subspecies	novicida 

F. tularensis subspecies novicida is of low virulence. It occasionally causes disease in 

immunocompromised individuals. Subspecies novicida was first isolated in 1951 from water 

samples in Utah, USA, and has subsequently been reported as a cause of tularaemia in at least 

four patients, all from the USA (Hollis et al., 1989; Clarridge et al., 1996). Three patients all 

had underlying immunocompromising conditions. They presented with high fever, and blood 

cultures yielded growth of subspecies novicida. A fourth patient slowly developed an enlarged 

granulomatous neck gland, from which the bacterium was isolated. Subspecies novicida has 

also been isolated from patients with various clinical manifestations in Canada, Australia and 

Spain (Bernard et al., 1994; Escudero et al., 2003; Whipp et al., 2003) which indicates a broad 

range of clinical manifestations and a wide geographical distribution for this subspecies. 

4.1.5	 Francisella philomiragia 

F. philomiragia is of low virulence and occasionally causes disease in immunocompromised 

individuals. In such individuals, F. philomiragia may cause pneumonia or fever with or without 

organ manifestations, such as meningitis or peritonitis. Epidemiological data suggest that the 

source of infection with F. philomiragia is associated with salt water. Five out of the 16 cases 

reported to date (Wenger et al., 1989; Sicherer et al., 1997; Friis-Møller et al., 2004) had a 

history of sea-water exposure in relation to near-drowning and were treated with high-dose 

corticosteroids. Seven patients had chronic granulomatous disease, a condition in which the 

phagocytic host defence is severely compromised. 

4.2	Signs	and	symptoms 

The clinical manifestations of tularaemia depend also on the route of acquisition (Table 4.1). 

When the bacteria are acquired through skin or oral mucous membranes, the result is usually 

a conspicuous enlarged tender node. When bacteria are inhaled, the infection will result in 

deep lymph-node enlargement. 

4.2.1	 Ulceroglandular	and	glandular	forms 

In terms of numbers of epidemics, ulceroglandular and glandular tularaemia are by far the 

most frequent forms of the disease. In European countries, they may comprise more than 

95% of the outbreaks. These forms are acquired by vector-borne transmission, direct contact 

with an infected animal, or indirect contact such as from tools used while dressing animals. 

Although acquisition by direct contact may occur in the absence of visible skin lesions, skin 

injuries will facilitate infection (Anda et al., 2001). Moreover, tularaemia associated with ani12
 
 

4. clinicAl exPression in HumAns 

Table	4.1 

Clinical forms of tularaemia 

Form	 Route	of	acquisition 

Ulceroglandular or glandular	 Vector-borne and direct contact (touching infected animals or material 

contaminated with F. tularensis) 

Oculoglandular	 Touching the eye with contaminated fingers or possibly from infective dust 

Oropharyngeal	 Ingesting contaminated food or water 

Respiratory	 Inhaling contaminated dust or laboratory-acquired infection 

Typhoidal	 Unknown (probably oral or respiratory) 

mal bites is occasionally reported (Arav-Boger, 2000; Friedl, Heinzer & Fankhauser, 2005). 

In ulceroglandular tularaemia, a primary ulcer develops at the site of bacterial exposure. 

The primary tularaemia ulcer is most often innocent (Figure 4.1) and may not even be noticed 

until the patient is medically examined. In subjects showing a number of mosquito bites, it 

may be difficult to decide which bite was infected. At about the time of onset of fever, a small 

papule appears and within a few days, it develops into a pustule surrounded by a zone of 

inflammation. Although the ulcer is usually solitary, several papules and pustules may occasionally be found. The ulcer soon heals, leaving a red thin area of 1 cm2 eventually developing 

into a scar which resembles that of the bacilli Calmette-Guérin (BCG) vaccination. The term 

glandular tularaemia is used only to indicate that in some cases the primary ulcer is nondetectable. 

Within a few days of onset of fever, the patient will perceive a regional lymph node 

enlargement. The lymph node soon becomes tender and palpable and often even visible. The 

overlying skin may be reddened and oedematous. Besides fever and unspecific symptoms, 

the lymph node enlargement is a major cause of concern and will often be the reason why the 

patient seeks medical attention (Figure 4.2 ). Provided appropriate therapy is started within a 

week of onset of disease (i.e. fever), the swelling of the lymph node resolves without further 

complications. However, when there is a treatment delay of more than 2 weeks, the risk of 

lymph node suppuration is as high as 30–40% (Kavanaugh, 1935; Stuart & Pullen, 1945). In 

an outbreak of tularaemia in Spain, where the ulceroglandular form was associated with a 

mean treatment delay of 43.4 ± 29.1 days, the presence of a constitutional syndrome without 

rigors was observed in most patients (Pérez-Castrillòn et al., 2001). In these cases, the lymph 

node remained enlarged and became soft upon palpation indicating an insidious melting 

process. Incision within the first weeks of infection should be avoided because of the risk of 

local spread of the infection (Kavanaugh, 1935). 

4.2.2	 Oculoglandular	tularaemia 

By touching the eye with a contaminated finger or possibly by exposure to F. tularensiscontaining dust, the oculoglandular form of tularaemia may be acquired (Kavanaugh, 1935; 

Foshay, 1940; Guerrant et al., 1976; Steinemann et al., 1999). This form comprises less than 

1% of all human cases of tularaemia, although higher figures (4.2%) have been described 

in more recent outbreaks (Pérez-Castrillòn et al., 2001). Together with fever and unspecific symptoms, the patients present with a unilateral conjunctivitis, usually expressed as an 

intense red conjunctiva with granulomatous lesions on the palpebral conjunctiva, swelling 

of the eyelids, excessive lacrimation, photophobia, and mucopurulent discharge (Chappell, 

Brainard & Shock, 1981). A large, tender preauricular lymph node will develop, sometimes 

changing the contour of the cheek. 

4.2.3	 Oropharyngeal	tularaemia 

Through ingestion of contaminated water or food, the oropharyngeal form may be contracted. A study in Finland between 1967 and 1983 included about 1100 cases of tularaemia 

of which 127 presented with disease localized in the head and neck region. Of these 127 cas13
 
wHo guidelines on tulArAemiA 

es, 32 were compatible with the oropharyngeal form (Luotonen et al., 1986). In Turkey, the 

oropharyngeal form is common; in a literature survey of Turkish outbreaks between 1936 

and 2004, 387 of 507 cases (77%) were referred to as the oropharyngeal form (Karadenizli 

et al., 2005). The patient presents with an ulcerative-exudative stomatitis and pharyngitis, 

with or without tonsillar involvement (Foshay, 1940; Hughes & Etteldorf, 1957; Luotonen 

et al., 1986). Physical examination shows redness and pustular changes in the mouth and 

pharyngeal mucous membranes. The disease is associated with an excessive regional neck 

lymphadenitis, although remarkably the lymph node enlargement is most often unilateral. 

These signs are easily misinterpreted as being caused by streptococci. In that case, patients 

will receive penicillin which does not affect F. tularensis. For instance, in a Turkish outbreak of 

oropharyngeal tularaemia, diagnosis was delayed and suppurating lymph nodes developed 

in 40% of the cases (Helvaci et al., 2000). 

Before a case of tularaemia can be diagnosed as oropharyngeal tularaemia, ulceroglandular 

tularaemia needs to be excluded. This is not always easy, because patients with tularaemia 

acquired by mosquito or tick bites in the head and neck region may show neck lymph node 

enlargement in the absence of a primary skin ulcer. An epidemiological investigation of the 

cases’ food and water sources may allude to oropharyngeal tularaemia. 

4.2.4	 Respiratory	tularaemia 

The respiratory form of tularaemia is contracted by inhalation of aerosolized F. tularensis. 

Most frequently, this occurs during farming activities. Outbreaks occur less frequently than 

those of ulceroglandular tularaemia, but may involve a high number of cases. The source of 

bacterial aerosols is believed to be carcasses of rodents or lagomorphs remaining in the fields 

after death from tularaemia. Respiratory tularaemia may present with symptoms of pneumonia, including cough, chest pain, and an increased respiratory rate. It may present with high 

fever and unspecific symptoms such as nausea and vomiting. Respiratory symptoms may be 

absent. Pneumonia may occur as a primary manifestation of the respiratory form but may 

also appear as a complication in any form of tularaemia, due to bacteraemic spread. In respiratory tularaemia, the course of disease differs markedly between type A and type B (Tärnvik 

& Berglund, 2003). 

In type A disease, pneumonia is a fulminant condition which, prior to the advent of antibiotic therapy, was associated with a fatality rate of 30–60%. The onset is sudden with chill, 

high fever, dyspnoea, dry or productive cough, pharyngitis, chest pain, headache, profuse 

sweating, drowsiness and general weakness. The condition is extremely severe and may simulate symptoms and signs of typhoid fever, including mental deterioration and a pulse-temperature dissociation (Stuart & Pullen, 1945; Dienst, 1963; Avery & Barnett, 1967; Evans et 

al., 1985; Gill & Cunha, 1997). 

Volunteers exposed to aerosols of virulent F. tularensis type A developed acute illness 3–5 

days later, including systemic symptoms. Less than half of them had radiological evidence of 

pneumonia (McCrumb, Snyder & Woodward, 1957; Saslaw et al., 1961b). Obviously, in these 

studies, the development of pneumonic changes may have been suppressed by an early application of antibiotic treatment. In a majority of patients contracting natural respiratory type A 

tularaemia, radiographic changes are demonstrable. The changes are highly variable. In the 

early phase, discrete peribronchial infiltrates may be discerned, followed by bronchopneumonial infiltration in one or more lobes. The X-ray appearance may simulate a wide variety 

of conditions, including pneumococcal pneumonia, tuberculosis, lymphoma, or carcinoma of 

the lungs. An intentional aerosol release of F. tularensis type A would be expected to result in 

clinical manifestations similar to those recognized in natural respiratory tularaemia. 

Respiratory type B tularaemia has been described in large outbreaks in Sweden and Finland: during the outbreak in Sweden caused by contaminated hay, only 7% of the 140 patients 

had symptoms of pneumonia (Dahlstrand, Ringertz & Zetterberg, 1971). In contrast, in an 

outbreak among farmers in Finland, 80% of the 53 cases showed radiological pulmonary 

14
 
4. clinicAl exPression in HumAns 

changes (Syrjälä et al., 1985a). Hilar adenopathy was the most common finding and occurred 

in 13 of 38 cases, followed by pneumonic infiltration (5 of 38 cases) and pleural effusion (1 of 

38 cases). See Figure 4.3 for respiratory type B tularaemia with hilar lymphadenopathy and 

Figure 4.4 for respiratory type B tularaemia with extended pulmonary consolidation. 

4.2.5	 Typhoidal	tularaemia 

Historically, the typhoidal form was defined as tularaemia devoid of skin or mucous membrane lesion and/or a remarkable lymph node enlargement (Francis & Callender, 1927). This 

definition dates back to the time when modes of transmission of F. tularensis were barely 

known. The classification of clinical symptoms and signs is now based on a combination 

of clinical expression and route of acquisition of the infection. A continued use of the term 

“typhoidal” seems unjustified and confusing (Avery & Barnett, 1967; Syrjälä et al., 1985a). 

Only when no route of infection can be established may the term still be acceptable. 

4.2.6	 Tularaemia	in	childhood 

Although conforming to general features of the disease, there are some differences in clinical 

symptoms and signs between children and adults. In Arkansas, USA, where type A tularaemia 

is prevalent, lymphadenopathy has been reported to occur more frequently in children than 

in adults, with a more frequent cervical localization and a tendency for delayed suppuration 

(Jacobs & Narain, 1983; Jacobs, Condrey & Yamauchi, 1985). Also pulmonary manifestations 

have been observed more frequently (Jacobs, Condrey & Yamauchi, 1985). A relative increase 

in degree of severity, reported in children in Scandinavia (Uhari, Syrjälä & Salminen, 1990), 

probably relates to an increased delay of diagnosis of tularaemia in childhood. 

4.2.7	 Tularaemia	in	immunosuppressed	patients 

Tularaemia occurs mainly among previously healthy subjects. The experience of tularaemia 

in immunosuppressed patients is limited. Since cell-mediated immunity is mandatory to host 

resistance against tularaemia, a fulminant course of disease may be expected in patients with 

T-cell deficiency, such as lymphoma, HIV disease and in those undergoing corticosteroid or 

cytostatic treatment. 

4.2.8	 Extraordinary	clinical	expression	of	tularaemia 

F. tularensis has been isolated occasionally from patients hospitalized because of febrile disease 

of unknown origin. A majority of these patients were compromised by neutrophil deficiency 

(Maranan et al., 1997; Sarria et al., 2003), transplantation-associated immunosuppressive 

therapy (Naughton et al., 1999; Khory et al., 2005), stomach cancer (Han, Ho & Safdar, 2004), 

HIV disease (Gries & Fairchok, 1996) or the presence of a prosthetic medical device (Pittman, 

Williams & Friedman, 1996; Cooper et al., 1999). Although these subjects lived in areas where 

F. tularensis is endemic, illness was not preceded by obvious exposure events and the route 

of transmission of F. tularensis remained unknown. Thus, F. tularensis has to be kept in mind 

when fastidious Gram-negative coccobacilli are isolated from blood or other deep sites. 

4.2.9	 Complications 

Tularaemia is associated with a variety of complications. In ulceroglandular type A tularaemia, 

pneumonia may present with parenchymal infiltrates and often also with pleural effusion. In 

an overview from the USA, ulceroglandular type A tularaemia was associated with pneumonia in 30% of the cases (Evans et al., 1985). Only occasionally, hilar lymphadenitis was 

found. In North America, where type A tularaemia is prevalent, fulminant manifestations are 

reported, including severe septicaemia (Provenza, Klotz & Penn, 1986), meningitis (Rodgers 

et al., 1998), endocarditis (Tancik & Dillaha, 2000), hepatic failure (Gourdeau et al., 1983; 

Evans et al., 1985; Ortego et al., 1986) and renal failure (Tilley, Garman & Stone, 1983; Penn 

15
 
wHo guidelines on tulArAemiA 

(B) 10 weeks after onset of disease and successful 

treatment with oral doxycycline (Tärnvik & Berglund, 

2003, reproduced with permission from the 

European Respiratory Journal). 

Figure	4.1 Primary ulcer in a human case 

of tularaemia (reproduced with the courtesy of 

Department of of Infectious Diseases, Umeå 

University, Sweden). 

Figure	4.3 Hilar enlargement in a 24-year-old 

farmer with fever but with no lower respiratory tract 

symptoms. Tularaemia was serologically verified 

and radiography was performed (A) 13 days, and 

Figure	4.2	Lymph node enlargement in a human 

case of glandular tularaemia (reproduced with the 

courtesy of Department of of Infectious Diseases, 

Umeå University, Sweden). 

Figure	4.4 Extended right-sided pulmonary 

consolidation in a 16-year-old male with fever and 

a productive cough. Tularaemia was serologically 

confirmed and radiography was performed (A) 11 

days, and (B) 2 months after the onset of disease 

and successful treatment with oral ciprofloxacin. 

(Tärnvik & Berglund, 2003, reproduced with 

permission from the European Respiratory Journal). 

16
 
17
 

4. clinicAl exPression in HumAns 

usually extremely 

occur early in the 

ating from confu­

& Kinasewitz, 1987). Septicaemia associated with type A tularaemia is 

severe and potentially fatal. High fever, abdominal pain and diarrhoea may 

course of disease and may be followed by changes in mental state deterior

sion to coma. A disseminated intravascular coagulation syndrome or an acute respiratory 

disease syndrome may occur and the patient may succumb to septic shock. 

In type B disease, meningitis (Hill et al., 1990) and septicaemia (Tärnvik et al., 1989; Hoel 

et al., 1991) are occasionally reported. All patients included in these reports recovered. As 

a complication of type B tularaemia, pneumonia seems to occur only rarely. In a Swedish 

outbreak in 1981, mainly of ulceroglandular disease, pneumonia was diagnosed in 1 of 400 

cases (Christenson, 1984). However, a delay in the diagnosis can increase the incidence of 

such complications. In a Spanish outbreak, a considerable delay in diagnosis occurred (mean 

47.5 pm 31 days) and 5 of the 142 patients developed pneumonia (Pérez-Castrillòn et al., 

2001). 

In both type A and type B tularaemia, immune-mediated skin reactions occur frequently, 

including erythema nodosum and erythema multiforme (Perman & Maclachlan, 1931; Syrjälä, Karvonen & Salminen, 1984; Evans et al., 1985; Greco et al., 1987, Akdis et al., 1993). 

4.2.10	 Blood	chemistry 

Routine blood chemical analyses show sparse abnormalities. In type A tularaemia, the white 6 3 blood cell count may be normal or raised up to 20 x 10 /mm and the differential count typically shows a relative increase of lymphocytes. Liver enzymes may show slightly increased 

values (Evans et al., 1985). 

A study of type B tularaemia in Finland comprised laboratory data on 52 cases with pulmonary changes and 42 cases with the ulceroglandular and glandular forms (Syrjälä, 1986). The 

mean leukocyte count was 8.3 x 10 /mm 6 3 and the differential count unremarkable. The mean 

C-reactive protein value peaked at 53 mg/l, which is remarkably low for an invasive disease. 

The sedimentation rate, however, remained increased at 30–50 mm/h for the first month after 

onset of disease. 

4.3	Differential	diagnoses 

Tularaemia may be mistaken for a range of other diseases (Table 4.2). Among these are a wide 

variety of conditions presenting with fever and lymph node enlargement. When the epidemiology is suggestive, tularaemia should be considered in any case of fever of unknown origin. 
 

 

 

 

 

 

 

 

 

 

 

 

wHo guidelines on tulArAemiA 

Table	4.2 

Differential diagnosis by route of acquisition 

Disease	 Route	 	 Difference	from	tularaemia		 

	 Ulc1	 Res2 

Staphylococcal x Staphylococcal disease such as furunculosis is less often associated 

with general symptoms 

Streptococcal x Streptococcal disease is usually associated with intense local inflammation, including erysipelas or more skin lesions or less extensive 

impetiginous lesions, whereas the tularaemia skin lesion is remarkably mild and limited in size. 

Anthrax x x Cutaneous anthrax is associated with a skin blister which undergoes 

necrosis. Although similar to the tularaemia ulcer in that the anthrax 

lesion is typically painless, it is assocated with extensive tissue damage. Respiratory anthrax develops more rapidly than tularaemia into 

a toxic fatal state, and this occurs irrespective of antibiotic therapy. In 

anthrax the radiological examination shows a mediastinal widening 

due to oedema and bleeding which may not be easily distinguished 

from mediastinal lymphadenitis such as present in tularaemia. 

HIV x Highly variable clinical expression, including fever and lymph node 

enlargement; antibody and/or antigen assays are confirmative. 

CMV disease x Febrile disease with lymph node enlargement; antibody and/or antigen assays are confirmative. 

EBV disease x Febrile disease with lymph node enlargement; antibody assays are 

confirmative. 

Pasteurellosis x Pasteurellosis usually results from cat and dog bites or scratches 

and is associated with intense local inflammation, including erythema 

and oedema. The local inflammation is more pronounced and the 

lymph node enlargement less prominent than in tularaemia. Pasteurella 

multocida is easily isolated from wound specimens. 

Mycobacteriosis x Mycobacterial disease may present with the same kind of local lymphadenopathy as tularaemia, although at a much slower rate of development. Moreover a cytological or histological specimen may show 

indistinguishable patterns in mycobacterial disease and tularaemia, 

including lymphocytes, epitheloidal cells and multinuclear giant cells. 

Mycobacterial disease is confirmed by demonstration of acid-fast 

bacilli; culture of the bacteria and PCR. 

Rickettsiosis x The spotted fever group of rickettsiosis, including Rocky Mountain 

fever in North America and fièvre boutonneuse in the Mediterranean 

region and Africa, present with fever and skin manifestations and are 

spread by ticks. Rocky Mountain typically involves an exanthema and 

the fièvre boutonneuse the development of eschar. Rickettsiosis is 

usually confirmed by demonstration of serum antibodies. 

Toxoplasmosis x Toxoplasmosis is a febrile zoonosis which may show lymph node 

enlargement and fever and which is usually confirmed by demonstration of serum antibodies. 

Cat-scratch disease x Cat-scratch disease is caused by Bartonela henselae and acquired 

by contact with cats; most often involving scratches. The primary 

skin lesion and lymph node enlargement are similar to tularaemia, but 

fever and other systemic symptoms are less pronounced. Laboratory 

confirmation is not easy although culture from blood may be successful. 

Plague x x Bubonic plague develops within 2–6 days of exposure such as by a 

flea bite. Buboes, which are enlarged lymph nodes, become extremely swollen and tender. The course of disease is usually more rapid and 

general symptoms more fulminant than those of ulceroglandular tularaemia. Septic shock may ensue, resembling shock associated with 

Gram-negative septicaemia. Plague pneumonia is rapidly fulminant, 

producing copious watery or purulent sputum production, haemoptysis, respiratory insufficiency and shock. It is almost invariably fatal. 

Plague is suspected by epidemiological circumstances and demonstrated by isolation of the causative agent. 

18
 
 

 

 

 

 

 

 

4. clinicAl exPression in HumAns 

Table 4.2 (contd) 

Disease	 

	 

Route	 

Ulc1	 

	 

Res2 

Difference	from	tularaemia		 

Lymphoma x Lymphoma is an obvious differential diagnosis of concern both in 

glandular and respiratory tularaemia. Hilar lymphadenopathy in tularaemia may be radiologically indistinguishable from lymphoma (see 

Figure 4.2). 

Brucellosis x Brucellosis is a febrile zoonosis which may show few specific signs 

and symptoms. It may be confirmed by blood culture and also by 

serology. 

Hantavirus disease x Hantavirus pulmonary syndrome is a severe febrile noncardiogenic 

pulmonary oedema which is spread by aerosols from rodents and 

occurs in the Americas. Hantavirus disease is confirmed by serology. 

Leptospirosis x Leptospirosis is a febrile zoonosis which is predominantly spread by 

rodents. It is usually confirmed by serology. 

Influenza x Influenza is rapidly diagnosed by direct immunofluorescent assay of 

respiratory secretions. PCR is a more sensitive method performed to 

an increasing extent in microbiological laboratories. 

Atypical pneumonia x Pneumonia caused by Mycoplasma pneumoniae, Chlamydia pneumoniae or Legionella pneumophila may resemble respiratory tularaemia. Epidemiological circumstances including spread among 

humans (Mycoplasma, Chlamydia) and association with irrigation and 

air-conditioning systems and showers (Legionella) may be suggestive. Usually serology will be confirmative. 

1 Ulceroglandular; 2 Respiratory. 

19
 

5. treAtment

5.	 Treatment 

Pedro Anda, Andrew Pearson, Arne tärnvik 

Despite improvements in therapy over the past decades, type A tularaemia is still associated 

with fatal outcome. Type B tularaemia, although much less virulent than type A, is frequently 

associated with suppurative complications and a considerable period of convalescence. In 

patients with prolonged fever of unknown origin, an empiric trial of medication with betalactam antibiotics is often initiated. However, these will not affect F. tularensis and will thus 

have no effect on the course of disease. Currently, no vaccine against tularaemia is available (see section 8.2). Early recognition and appropriate antibiotic treatment of the disease is 

essential. The following recommendations are derived from Dennis et al., 2001. 

5.1	 Recommendations	for	treatment	and	prophylaxis 

5.1.1	 Adults 

In severe tularaemia which requires hospitalization, parenteral administration of an aminoglycoside is the first choice for treatment. Gentamicin is preferred at 5 mg/kg daily, divided 

into two doses and monitored by assay of serum concentrations of the drug. If streptomycin is 

available, it is an alternative given by intramuscular injection 2 g daily, divided in two doses, 

for 10 days. In severe cases the treatment period will depend on clinical response, and may 

comprise more than 10 days. In less severe cases or in a mass casualty setting, oral ciprofloxacin or doxycycline is preferred. Ciprofloxacin 800–1000 mg daily, divided into two doses, 

may be given intravenously or by oral administration. The treatment period should be 10–14 

days. An alternative is doxycycline, 200 mg daily, divided in two oral doses and, due to the 

bacteriostatic nature of the drug, given for at least 15 days. 

5.1.2	 Children 

In cases which require hospitalization, parenteral administration of an aminoglycoside is 

preferred. The drug of choice is gentamicin, 5–6 mg/kg divided into two or three doses and 

monitored by assay of serum concentrations of the drug. If available, streptomycin 15 mg/kg 

twice daily (up to 2 g daily) is an alternative. In milder cases, particularly in areas endemic 

for the less virulent type B tularaemia, ciprofloxacin 15 mg/kg twice daily (up to 1 g daily), is 

a feasible alternative. Irrespective of the antibiotic chosen, treatment should be continued for 

at least 10 days. 

5.1.3	 Pregnant	women 

This is a situation where potential side-effects have to be weighed against the benefits of 

treatment of a severe infection. Even though gentamicin and ciprofloxacin are not approved 

for administration in pregnancy by the Food and Drug Administration of the USA, their use 

in pregnancy in tularaemia has been recommended by a working group on civilian biodefence (Dennis et al., 2001). Ciprofloxacin is an option and a brief course of gentamicin treatment is an alternative. Doses are the same as for non-pregnant subjects and the treatment 

period should be individualized. 

5.1.4	 Immunocompromised	patients 

In severe bacterial diseases, bactericidal agents are usually preferable for treatment in 

immunodeficiency. In tularaemia, however, there is little experience in this category of 

patients. An aminoglycoside should be the first-line drug, and ciprofloxacin an alternative. 

21
 
wHo guidelines on tulArAemiA 

22
 

Doses are the same as for non-compromised subjects and a treatment period of 14 days may 

be required. In case of the need for prolonged treatment, an initial aminoglycoside administration may be followed by a period of oral or parenteral ciprofloxacin treatment. 

5.1.5	 Post-exposure	prophylaxis 

Three situations are to be considered: 

1. Accidental exposure of laboratory personnel: antibiotic treatment should be initiated within 

24 h and a treatment period of 14 days is recommended with either ciprofloxacin 1000 mg 

daily divided in two doses, or oral doxycycline 200 mg daily, divided in two doses. 

2. Exposure most likely did not occur (in the laboratory): an increased vigilance may be sufficient, including daily measurement of body temperature for 14 days and a readiness to treat 

if symptoms appear. 

3. Incidental spread of F. tularensis by aerosol: potentially exposed persons should be instructed to be alert to the development of fever within 14 days of exposure, and treatment 

initiated if necessary according to schedule above. 

5.2	Antibiotic	agents 

Table 5.1 gives the minimal inhibitory concentrations (MICs) for various antibiotic agents 

against F. tularensis. 

Table	5.1 

Minimum inhibitory concentrations (MIC) (in mg/l) of various antimicrobial agents for 

F. tularensis in three studies 

Including 13 type A and 2 Type B isolates; the reference is: Baker, Hollis & Thornsberry, 1985. b Including 20 type B isolates; the references is: Scheel et al., 1993. 

Agent	 Bakera	 Scheelb	 Johanssonc 

	 	 	 Type	A	 Type	B 

Streptomycin 

Gentamicin 

≤0.5–4 

0.25–2 

2–16 

2–4 

0.064–2 

0.032–0.25 

0.032–1 

0.016–0.125 

Tetracycline 

Doxycycline 

Chloramphenicol 

Rifampin 

Ciprofloxacin 

≤0.25–2 

– 

≤0.25–4 

≤0.03–1 

– 

– 

<0.25–2 

1–8 

– 

0.031–0.125 

– 

0.125–2 

0.5–1 

0.25–2 

0.016–0.064 

– 

0.125–2 

0.25–1 

0.125–1 

0.016–0.064 

a 

c Including 8 type A and 16 Type B isolates; the reference is Johansson et al., 2002. 

5.2.1	 Aminoglycosides 

Streptomycin became established early as the drug of choice for treatment of tularaemia 

(Foshay, 1946). However, it is no longer widely used in human medicine because of its potential 

to cause vestibular toxicity and a frequent appearance of hypersensitivity reactions among the 

personnel involved in its administration. Therefore, streptomycin is no longer easily available and has largely been replaced by other aminoglycosides. In tularaemia, gentamicin is 

currently the preferred alternative. Nevertheless, streptomycin is still the drug of choice for 

tularaemia meningitis. 

Streptomycin 

Streptomycin is bactericidal in vitro and highly efficacious against F. tularensis. Out of 224 

streptomycin-treated cases documented in the literature, 217 were cured (Enderlin et al., 

1994). No relapses occurred and failures were restricted to severe cases, most frequently com­
5. treAtment 

plicated by renal dysfunction. In a study cited by Enderlin et al. (1994), of 141 patients treated 

with streptomycin, a mortality of 3% was reported, compared to 33% before the advent of 

antibiotic treatment. 

Gentamicin 

Although in a murine model, streptomycin was found to be more effective than gentamicin 

(Mason et al., 1980), most experience in human tularaemia suggests that gentamicin is a useful alternative. Gentamicin is the preferred aminoglycoside for parenteral treatment of severe 

cases which require hospitalization. 

A concern raised with gentamicin is the relatively poor penetration of the drug into cells 

under in vitro conditions. Nonetheless, gentamicin is internalized by pinocytosis although 

slowly, and susceptibility assays by in vitro cell systems show that gentamicin is capable of 

killing intracellular F. tularensis (Maurin, Mersali & Raoult, 2000). 

In a review of the literature, among 36 patients treated with gentamicin, 31 were cured 

(Enderlin et al., 1994). Of two relapses, one patient had been treated for only 6 days; the other 

was subjected to treatment after a delay of 43 days. This patient responded to a 9-day course, 

but developed tularaemia meningitis after withdrawal of treatment. 

5.2.2	 Chloramphenicol 

Chloramphenicol is bacteriostatic. It is now seldom used because it is associated with relapse 

and with rare but severe side-effects. The only advantage of chloramphenicol is a relatively 

high penetration into the cerebrospinal fluid which may be of value in treatment of tularaemia 

meningitis. 

5.2.3	 Tetracyclines 

Tetracyclines were widely used to treat tularaemia. Their disadvantage is their bacteriostatic 

nature and thus the risk of relapses (Sawyer et al., 1966; Enderlin et al., 1994), but tetracyclines are still valuable alternatives for oral treatment of tularaemia. For pharmacokinetic 

reasons tetracycline, which was used in the 1960s, has now been replaced by doxycycline. 

Tetracycline 

In tularaemia, cell-mediated immunity, and CD4 and CD8 T cells in particular, are mandatory 

to contain the infection. The peripheral blood T-cell response to F. tularensis usually becomes 

demonstrable in vitro 12–14 days after the onset of disease. Before that time, tetracycline can 

only be expected to inhibit the bacteria. Consequently, the bacteria will remain alive until 

bactericidal host mechanisms develop and become able to cope with the infection (Syrjälä et 

al., 1984; Tärnvik, 1979). To minimize the risk of relapse in case of treatment with bacteriostatic agents such as tetracycline, the treatment period needs to be long enough to allow the 

cell-mediated immune response to develop. 

A relationship between relapse and bacteriostatic action has been confirmed experimentally. When soldiers in the USA were voluntarily infected by an aerosol containing 25 000 

SCHU-S4 (type A) organisms (Sawyer et al., 1966), they developed tularaemia within 2–7 

days of exposure. Treatment with streptomycin, 2 g daily, divided into two doses, was started 

on the day of onset of fever and given for 6 days. Rapid cure occurred. When tetracycline was 

given, 0.5 g four times daily for 6 days, defervescence occurred as rapidly as with streptomycin but was followed by relapse within a week of withdrawal of treatment. In treatment trials, 

a daily dose of 2 g of tetracycline for 10 days or 1 g for 15 days was insufficient to prevent 

relapses. Only by increasing the dose to 2 g daily for 15 days, was the disease suppressed 

completely. These experimental data have been corroborated by experience from treatment 

of natural tularaemia. Among 50 cases reviewed, tetracycline treatment resulted in relapse in 

six cases (Enderlin et al., 1994). In at least three of these cases, the treatment period was less 

than 7 days. 

23
 
wHo guidelines on tulArAemiA 

Tetracycline has been tested as a prophylactic agent against tularaemia in volunteers. When 

initiated 24 h after exposure, oral tetracycline at a daily dose of 2 g for 14 days or 1 g for 28 days 

was sufficient to prevent disease, whereas 1 g daily for 14 days was not (Sawyer et al., 1966). 

Doxycycline 

A daily dose of 200 mg doxycycline is believed to correspond to 2 g of tetracycline. Based on 

experimental and clinical data on older formulations of tetracycline and by taking data on 

the immune response into consideration, 200 mg of doxycycline daily, divided into two oral 

doses, for at least 15 days is recommended in adults. On such a regimen, a mean serum concentration of 4 mg/l will be reached (Welling et al., 1977). This dose is expected to result in 

optimal antibacterial activity, since an MIC of 0.25–2 mg/l is reported by various laboratories 

(Table 5.1). A dose of 100 mg daily for 3 weeks might be an alternative, but is less convincingly supported by data from the literature. The side-effects of doxycycline are mild, mostly 

limited to gastrointestinal effects and mitigated by taking the drug with food. Unfortunately, 

doxycycline and other tetracyclines are not recommended for use in children under the age of 

8 years, due to possible adverse effects on developing teeth. 

5.2.4	 Quinolones 

More recently, quinolones have been introduced as new options for oral treatment of tularaemia. 

Most data are so far restricted to ciprofloxacin and to clinical use in type B tularaemia. Agar 

dilution assay and E-tests have shown MIC values for quinolones in F. tularensis in the range 

of 0.02–0.1 mg/l (Table 5.1) and broth dilution tests a MIC value of 0.25 mg/l and a minimum 

bactericidal concentration (MBC) also of 0.25 mg/l (Maurin, Mersali & Raoult, 2000), verifying the bactericidal activity of the agent. In Scandinavia, a first clinical report included four 

patients treated with oral ciprofloxacin 750 mg twice daily and one patient treated with norfloxacin 400 mg twice daily; all recovered within a few days without relapse (Syrjälä, Schildt 

& Raisainen, 1991). In further studies, 12 children were successfully treated (Johansson et al., 

2000a) and 41 of 43 adults (24 with ulceroglandular, 3 with pulmonary, and 14 with typhoidal tularaemia) showed excellent responses to oral ciprofloxacin given for 10 days (Johansson et al., 2001a). In a tularaemia epidemic in north-western Spain, comprising 142 patients, 

the efficacy of ciprofloxacin was reported to be higher than that of streptomycin or doxycycline and moreover, ciprofloxacin treatment was associated with fewer side effects (PérezCastrillòn et al., 2001). In another report from Spain, relapse was recorded in seven of 14 

patients treated with ciprofloxacin (Chocarro, Gonzalez & Garcia, 2000), although in that 

group of patients, treatment failure may have been due to a considerable delay from onset of 

disease to start of treatment. 

Apart from ciprofloxacin, only sporadic cases have been described using quinolones for 

tularaemia treatment. Two subjects were reported to be successfully treated with levofloxacin; 

both were acutely ill patients and neither relapsed within 12 months of follow-up (Limaye & 

Hooper, 1999). 

Altogether, quinolones offer new options for treatment of tularaemia. As far as type B tularaemia is concerned, they seem to be useful for children. Although in early studies, quinolones were shown to cause arthropathy in immature animals, the risk is currently considered to 

be low in humans and the drug to be safe for children. Obviously, the lack of oral alternatives 

has hampered a rational handling of tularaemia in childhood. A review of 67 children suffering from tularaemia showed a mean duration of symptoms as long as 26 days (range, 8–92 

days), probably related at least in part to the fact that 20 patients received drugs known to be 

ineffective against F. tularensis (Uhari, Syrjälä & Salminen, 1990). 

The therapeutic use of quinolones has so far been largely restricted to type B tularaemia. 

In vitro, type A (F. tularensis subspecies tularensis) also seems to be susceptible to quinolones. 

Isolates from various areas in the USA were found to have MIC values of ciprofloxacin as low 

as 0.016–0.064 mg/l (Johansson et al., 2002) (Figure 5.1). Moreover, these type A isolates and 

24
 
5. treAtment 

24 type B isolates all showed MIC values < 0.125 mg/l for six different quinolones. It needs to 

be emphasized, however, that the efficacy of ciprofloxacin for treatment of type A tularaemia 

has not been proven. On the contrary, there is experience of treatment failure in children in 

the USA (Jacobs RF, personal communication). It should be recalled also that ciprofloxacin 

and other quinolones are not approved for use in tularaemia by the Food and Drug Administration of the USA. 

5.2.5	 Other	antibiotics 

Beta-lactams such as penicillin are ineffective against F. tularensis. Although ceftriaxone has 

been found to be active in vitro, several cases of therapeutic failure following the use of this 

drug have been experienced (Cross & Jacobs, 1993; Enderlin et al., 1994). Neither is erythromycin a reliable drug for tularaemia, despite a susceptibility of type A organisms. Rifampicin 

is active in vitro but is not recommended for clinical use, due to a potential for induction of 

resistance (Bhatnagar et al., 1994; Johansson et al., 2000a). Co-trimoxazole and clindamycin 

are ineffective. 

5.3	 Antibiotic	resistance	of	F. tularensis 

No natural resistance in F. tularensis to antibiotics used for clinical therapy has been demonstrated. This is true for aminoglycosides, tetracyclines, chloramphenicol, and quinolones. 

Erythromycin resistance, however, is prevalent in Europe but not in North America. Although 

erythromycin is not included among agents used for treatment of tularaemia, erythromycin 

resistance may be used as an epidemiological marker. 

For experimental purposes, streptomycin- and tetracycline-resistant strains of F. tularensis 

have been developed and most probably, quinolone resistance is easily introduced as well. Yet, 

the risk for development of antibiotic resistance of importance in clinical practice is low. The 

reason is that tularaemia in humans is the end-stage of the infection. Most evidence indicates 

that F. tularensis is not transferred from one family member to another or from patients to 

hospital staff. Neither are there any data to suggest that tularaemia might be spread among 

humans by mosquitoes. 

Due to the potential use of F. tularensis for bioterrorism, antibiotic resistance remains of 

concern. Consequently, methods for rapid determination of the susceptibility of F. tularensis 

to various antibiotics, including aminoglycosides, tetracyclines, chloramphenicol, quinolones, 

and rifampicin are needed. 

25
 
wHo guidelines on tulArAemiA 

Figure	5.1 Distribution of MIC values for six quinolones. Open bars represent isolates of F. tularensis subspecies 

tularensis, and grey bars represent F. tularensis subspecies holarctica. (Johansson et al., 2002, reproduced with 

permission from the Scandinavian Journal of Infectious Diseases). 

Ciprofloxacin Gatifloxacin 

16 16 

14 14 

12 12 

4 4 

2 2 

0 0 

0,008 0,016 0,032 0,064 0,125 0,008 0,016 0,032 0,064 0,125 

MIC (mg/L) MIC (mg/L) 

Levofloxacin Grepafloxacin 

16 16 

14 14 

12 12 

No of isolates No of isolates No of isolates 10 

8 

6 

No of isolates No of isolates No of isolates 10 

8 

6 

10 10 

8 8 

6 6 

4 4 

2 2 

0 0 

0,008 0,016 0,032 0,064 0,125 0,008 0,016 0,032 0,064 0,125 

MIC (mg/L) MIC (mg/L) 

Moxifloxacin Sparfloxacin 

16 16 

14 14 

12 12 

10 10 

8 8 

6 6 

4 4 

2 2 

0 0 

0,008 0,016 0,032 0,064 0,125 0,008 0,016 0,032 0,064 0,125 

MIC (mg/L) MIC (mg/L) 

26 
6. lABorAtory diAgnostics And discriminAtion oF suBsPecies And strAins

6.	 Laboratory	diagnostics	and	 

	 discrimination	of	subspecies	and	strains 

Anders Johansson, Jeannine Petersen, Anders sjöstedt 

6.1	 Case	definitions 

The case definitions that describe the criteria for diagnosis of tularaemia are: suspect, presumptive and confirmed. The criteria for each are: 

Suspect. An exposure history consistent with risks known to be associated with tularaemia 

together with clinical symptoms compatible with tularaemia. 

Presumptive. Suggestive clinical symptoms and a clinical sample that tests positive for tularaemia by antigen or DNA detection. A single positive serum is also considered presumptive. 

Confirmed. Recovery of an isolate and identification of the culture as F. tularensis by antigen 

or DNA detection. Alternatively, paired serum specimens with a fourfold difference in titre 

(tube or microagglutination assay) or significantly (ELISA), with at least one serum positive, 

are also considered confirmatory. 

6.2	Diagnostic	specimens	 

6.2.1	 Human 

In principle, specimens for recovery of live bacteria should be collected before antibiotics are 

administered. In cases with preceding or ongoing therapy, culture may still be useful, particularly when beta-lactam antibiotics or other agents inactive against F. tularensis were used. 

The choice of specimen for diagnostic testing is dependent on the form of clinical illness; 

ulceroglandular, glandular, oculoglandular, oropharyngeal, respiratory, or typhoidal. The following specimens are acceptable for the various forms of illness as specified: 

Blood. Whole blood (for all clinical forms of illness). 

Serum. Serum is preferred; plasma and whole blood dired on paper filter may be an acceptable 

alternative. A first specimen should be collected as early in the course of infection as possible, 

followed by a second specimen taken in the convalescent period (at least 14 days later and 

preferably 3–4 weeks after onset of symptoms). Serum is acceptable for all clinical forms of 

illness. 

Respiratory secretions. Pharyngeal swabs, bronchial/tracheal washes or aspirates, sputum, 

transthoracic lung aspirates, or pleural fluid collection (for respiratory, typhoidal, oropharyngeal forms of illness). 

Swabs. Swabs of visible lesions or affected areas should be collected (for ulceroglandular and 

oculoglandular forms of illness). 

Aspirates. Aspirates from lymph nodes or lesions (for ulceroglandular, glandular, and oropharyngeal forms of illness). 

Tissue biopsies. Tissue samples from lymph nodes (for ulceroglandular, glandular, and oropharyngeal forms of illness). Invasive sampling, such as incision of an affected lymph node, 

should be avoided during the acute stage of disease. Experience indicates that such intervention may further the spread of the infection. There is little experience on the value of biopsies 

for diagnostic purposes. 

Autopsy materials. Samples from visible abscesses and from lymph node, lung, liver, spleen, 

cerebrospinal fluid, and bone marrow. 

27
 
wHo guidelines on tulArAemiA 

6.2.2	 Mammals 

Serum. Serum is the preferred specimen but plasma and whole blood dried on filter paper may 

be acceptable alternatives. A second specimen should be taken at least 14 days, and preferably 

3–4 weeks after the onset of symptoms (serum is acceptable for all clinical forms of illness). 

Aspirate. Aspirates from lymph nodes or lesions (for ulceroglandular, glandular, and oropharyngeal forms of illness). 

Autopsy materials. Samples from visible abscesses and from lymph node, lung, liver, spleen, 

cerebrospinal fluid, and bone marrow are acceptable specimens. See section 7.4.4 for recommendations on the collection of samples from dead animals. 

6.2.3	 Arthropods 

Ticks, deer flies and mosquitoes have all been implicated in the transmission of tularaemia. 

For recovery of live organisms, vectors should be transported live or frozen at -80 °C. Mouse 

inoculation can be used to isolate pathogenic F. tularensis subspecies from infected ticks and 

deerflies (Klock, Olsen & Fukushima, 1973; Hubalek et al., 1996). Molecular assays have also 

been described for testing ticks (Goethert, Shani & Telford, 2004; Kugeler et al., 2005). Multiple targets should be assayed and amplification products sequenced. Dermacentor spp. have 

been shown to harbour Francisella-like endosymbionts that cross-react with molecular targets 

used for the detection of F. tularensis (Scoles, 2004; Kugeler et al., 2005). 

6.2.4	 Environmental	specimens 

Despite the fact that F. tularensis is often referred to as living for months in mud or contaminated soils, environmental samples such as water, soil and rodent faeces should only be collected in the context of an epidemiological investigation. Optimal methods for recovery and 

identification of F. tularensis from environmental samples do not yet exist. Prior to application 

of any methodology for the detection of F. tularensis in environmental samples, the sensitivity 

and specificity of the test should be determined on spiked samples. If samples are collected 

in the context of an epidemiological investigation, samples should be split into two fractions. 

One set of samples should be spiked with a known number of F. tularensis in order to test the 

purification protocol prior to diagnostic testing of unspiked samples. In addition, molecular 

assays must also be evaluated for cross-reactivity with Francisella-like organisms present in 

soil and water (Barns et al., 2005). 

6.2.5	 Collection	and	transport	of	human	specimens 

Every effort should be made to collect and preserve specimens so that viable bacteria can be 

recovered. Decontaminate the surface area prior to specimen collection since contamination 

of the sample with normal flora could interfere with interpretation of culture results. Ensure 

that adequate volumes (depending on type of sample) are collected to avoid false negatives as 

a result of insufficient sample volume. Specimens should be labelled clearly with the patient’s 

name, identification number, source, specific site, date, time of collection, and the initials of 

the collector. To minimize loss in viability, specimens should be delivered to the laboratory 

within 24 h, preferably within 2 h. In case of prolonged transport (i.e. > 24 h), survival of 

F. tularensis is uncertain. Provided the appropriate transport medium is utilized (contact with 

laboratory recommended), molecular diagnostic techniques can still be applied. 

Blood. Two or more venous blood samples should be obtained, preferably from separate sites 

and comprising in total 30–40 ml, into bottles of a conventional blood culture system, such as 

the BACTEC system. Transport directly to the laboratory at room temperature. Hold at room 

temperature until placed onto the blood culture instrument or incubator. Do not refrigerate, 

but keep at ambient temperature (10–37 °C). If whole blood is to be tested by PCR, it is recommended that heparin is not used as an anticoagulant as it may inhibit the PCR reaction. 

28
 
6. lABorAtory diAgnostics And discriminAtion oF suBsPecies And strAins 

Serum. Serum is obtained by drawing the blood into a serum separator tube containing no 

additives or anticoagulants, allowing it to clot, and centrifuging to separate the serum. If 

serum is required for testing, separation from blood should take place as soon as possible 

after collection, preferably within 24 h at ambient temperature. Centrifuge for 10 min at 3000 

revolutions per min and transfer the serum into a plastic transport tube. Sera may be stored 

at 2–8 °C for up to 10 days. If testing is delayed for a long period, serum samples may be frozen. If separation on site is not possible, or is inadvisable for safety reasons, the blood sample 

should be stored at 2–8 °C. Do not freeze. 

Biopsy. Obtain a tissue specimen. If the lesion is large or there are multiple lesions, collect 

multiple specimens from representative sites. Submit tissue or scraping in a sterile container. For small tissue samples, add several drops of sterile normal saline to keep the tissue 

moist. Transport at room temperature for immediate processing. If processing of specimen is 

delayed, keep specimen chilled (2–8 °C). Invasive sampling, such as incision of an ascending 

lymph node, should be avoided during the acute stage of disease. Experience indicates that 

such intervention may further the spread of the infection. There is little experience on the 

value of biopsies for diagnosis. 

Swabs. Take samples of fluid with a sterile swab. Try to get as much fluid as possible onto the 

swab. If using a swab transport carrier, the swab should be reinserted into the transport package and the swab fabric moistened with the transport medium inside the packet. For transport of specimens for PCR analysis, a buffer solution containing a nuclease inhibitor should 

be used (Johansson et al., 2000b). For culture, a rayon-tipped plastic applicator and a tube 

containing Amies agar with charcoal (Copan Italia, Brescia, Italy) showed good preservation 

(Johansson et al., 2000b). Transport at 2–8 °C; room temperature is acceptable. If processing 

of the specimen is delayed, store it at 2–8 °C. 

Respiratory or sputum specimens.	 Collect specimens in screw-capped containers and transport them to the laboratory as quickly as possible to reduce overgrowth by commensal oral 

flora. For transit periods up to 24 h, transport bacterial specimens at ambient temperature. If 

processing of the specimen is delayed, keep it chilled (2–8 °C). 

Aspirate. Aspirate the fluid from the abscess with a sterile needle and syringe. Transfer the 

aspirate aseptically into a sterile tube. Transport the sample at room temperature for immediate processing. Keep the specimen chilled (2–8 °C) if processing is delayed. 

Autopsy specimens. Obtain a tissue specimen. If the lesion is large or there are multiple lesions, 

collect specimens from representative sites. Submit tissue or scraping in a sterile container. 

For small tissue samples, add several drops of sterile normal saline to keep the tissue moist. 

Transport at room temperature for immediate processing. If processing of specimen is delayed, 

keep the specimen chilled (2–8 °C). Formalin-fixed specimens can also be prepared and must 

be packaged separately from unpreserved autopsy specimens for bacterial isolation. 

Regulations for transport of diagnostic and infectious specimens 

Specimens to be shipped to diagnostic laboratories require special attention to safe packing of 

the material. See Annex E for transport guidelines. 

6.3	 Bacteriology 

F. tularensis is a tiny, faintly-staining, Gram-negative, non-motile, non-spore-forming bacterium (Chu & Weyant, 2003; Sjöstedt, 2005). Cells are pleomorphic, typically appearing as 

short rods or coccoid forms. A double-layered cell wall surrounded by an outer layer and 

electron-transparent capsule encases the bacterium. Large amounts of lipids are contained in 

both the cell wall and capsule. The fatty acid composition is unique, being high in saturated 

29
 
wHo guidelines on tulArAemiA 

30
 

even-chain acids (C , C , C ) and two long-chain hydroxyl acids (3-OH C , 3-OH 10:0 14:0 16:0 16:0

C18:0) (Jantzen, Berdal & Omland, 1979). The lipid A portion of the lipopolysaccharide (LPS) 

is modified by the addition of a phosphate-linked galactosamine in at least one strain of subspecies holarctica (Phillips et al., 2004). Electron micrographs have shown the presence of pili 

on the surface of F. tularensis (Gil, Benach & Thanassi, 2004). F. tularensis is aerobic and considered a fastidious organism due to its requirement of cysteine for enhanced growth (Chu & 

Weyant, 2003; Sjöstedt, 2005). The organism is relatively inert biochemically, with only a few 

sugars (glucose, maltose, sucrose and glycerol) utilized. F. tularensis does not reduce nitrate, 

is oxidase and urease negative and weakly catalase positive. Personnel handling diagnostic 

cultures of F. tularensis are at considerable risk of infection and need to practise appropriate 

precautionary measures (see sections 6.6 and 8.1). 

6.4	 Diagnostic	tests 

Diagnosis of tularaemia is based on recovery of an isolate, antigen or molecular detection, 

and serology. Protocols for these methods can be found in Annex A. Accurate identification 

of F. tularensis in diagnostic laboratories relies heavily on the use of specialized reagents or 

media. Commercial biochemical identification systems available in clinical diagnostic laboratories cannot be relied upon to accurately identify F. tularensis. If a diagnosis of tularaemia 

is suspected, the physician should notify the laboratory in order to increase the likelihood of 

detection. 

6.4.1	 Serology 

Serology is commonly used for confirmation of tularaemia. Antibody responses against 

F. tularensis are generally detectable in patients 10–20 days post-infection (Koskela & Salminen, 1985). Agglutination, either microagglutination or tube agglutination, is the standard 

serological test used for determining the presence of antibody to F. tularensis (Brown et al., 

1980). The protocol for microagglutination is given in section A.5.1. Antigens used to measure 

immune response include FopA, LPS, and an outer membrane carbohydrate-protein fraction 

(OMP) and whole killed cells. ELISA tests have also been utilized, with the caveat that IgM 

responses may be sustained for long periods and are thus not indicative of recent infection 

(Viljanen, Nurmi & Salminen, 1983; Bevanger, Maeland & Kvan, 1994). More recently an 

ELISA (directed against LPS) combined with Western blot (against antigen extracted from 

whole killed cells) showed very good sensitivity and specificity for diagnosis of infection 

(Schmitt et al., 2005). The ELISA protocol is in section A.5.2. 

6.4.2	 Culture 

Culture provides a conclusive diagnosis of infection and an invaluable resource for molecular 

epidemiology, subtyping and discovery of novel species and subspecies. Whenever possible, 

culture should be attempted. F. tularensis grows well on several types of cysteine/cystine-supplemented agar including enriched chocolate agar (CA), cystine heart agar with 9% chocolatized blood (CHAB), buffered charcoal yeast extract (BCYE), thioglycollate-glucoseblood agar 

(TGBA), and GC Agar II with 1% haemoglobin and 1% IsoVitaleX. For media formulations 

see section C.1. F. tularensis can be isolated from nutrient enriched specimens (tissues) on 

sheep blood agar (SBA), but cysteine-enriched media are strongly recommended for subculture as the organism will usually fail to thrive with continued passage on SBA. Additionally, 

growth on CHAB provides presumptive identification of F. tularensis as the organism shows 

characteristic growth on this medium (green, opalescent, raised, shiny colonies at 24–48 

h). For growth characteristics by subspecies see secion 6.5.2. Mouse inoculation can also be 

used for recovery of an isolate. Mice are very sensitive to infection by F. tularensis and typically become ill within 3–4 days of inoculation. Tissue samples (spleen and liver) taken from 

moribund mice can be cultured on agar media to recover live organisms (see protocol A.2.1). 
 

 

6. lABorAtory diAgnostics And discriminAtion oF suBsPecies And strAins 

Samples from ulcers frequently yield pure cultures of F. tularensis while other specimens may 

contain contaminant bacteria. CHAB medium supplemented with antibiotics (CHAB-A) significantly improves recovery rates from tissue sources contaminated or overgrown by other 

flora (Petersen et al., 2004b). In specimens where the presence of mixed flora is likely, the use 

of CHAB-A should be considered. For blood culture, the BACTEC system or an equivalent 

system is recommended. 

6.4.3	 Antigen	detection 

Antigen detection can be useful for either direct identification of F. tularensis in clinical specimens or for confirmatory identification of isolates recovered in culture. Direct fluorescent 

antibody staining, using a FITC-labelled rabbit antibody directed against whole killed F. 

tularensis cells, is a rapid assay for identification of F. tularensis in primary specimens or for 

confirmation of recovered isolates (Petersen et al., 2004a, see protocol A.3.1). Slide agglutination can be used for the rapid confirmation of recovered isolates (see protocol A.3.2). 

Immunohistochemical staining, using a monoclonal antibody directed against the LPS, can 

be used to visualize F. tularensis in formalin-fixed tissues (Zeidner et al., 2004). In addition, 

antigen detection can be performed by antigen capture ELISA (Grunow et al., 2000, see section A.3.3). 

6.4.4	 Molecular	detection 

A variety of PCR methods have been described for the molecular detection of F. tularensis. 

PCR can be a valuable diagnostic tool when organisms are non-cultivable or when culture 

is not recommended due to biosafety concerns. The majority of PCR tests for F. tularensis 

have been conventional PCR assays targeted at the genes fopA or tul4 encoding the outer 

membrane proteins (Fulop, Leslie & Titball, 1996; Sjöstedt et al., 1997; Higgins et al., 2000; 

Johansson et al., 2000b). The tul4 PCR assay has been validated with specimens from wounds 

from patients with ulceroglandular tularaemia (Johansson et al., 2000b; see protocol A.4.1). 

A multitarget real-time TaqMan PCR assay for F. tularensis has also been developed (Versage 

et al., 2003). This assay has increased specificity and rapidity over conventional PCR assays 

and can provide added sensitivity when testing specimens in which the number of organisms is expected to be quite low. The assay targets include the ISFtu2 element, 23kDa and tul4 

genes. This assay has been evaluated with animal tissues, and a limited number of human 

specimens, but further evaluation with human specimens is needed (Versage et al., 2003; 

Kugeler et al., 2006). 

In cases where diagnostic efforts are not directed specifically towards F. tularensis, 16S 

rDNA sequence identification can be particularly useful (Clarridge, 2004). Universal 16S 

rDNA primers can be used to accurately identify isolates recovered in culture. Francisellaspecific 16S rDNA primers should be used for contaminated samples (Forsman, Sandström & 

Sjöstedt, 1994; see protocol A.4.1). 

6.5	Supplemental	characterization 

Once an isolate has been confirmed as F. tularensis, a variety of supplemental tests can be 

used for additional characterization. Protocols for these methods can be found in Annex B. 

These tests can be extremely useful for determining the source of infection, understanding 

transmission cycles or identifying species or subspecies. 

6.5.1	 Antimicrobial	susceptibility 

Antimicrobial susceptibility testing is not routinely performed since naturally occurring 

resistance to the antibiotics used for clinical treatment of tularaemia has not been reported. 

Broth microdilution using cation-adjusted Mueller-Hinton broth (CAMHB) with 2% defined 

growth supplement can be used for susceptibility testing (Clinical and Laboratory Standards 

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wHo guidelines on tulArAemiA 

Institute, 2005). The E-test, a quantitative gradient diffusion test, has also been utilized to 

determine MICs of various antibiotics for F. tularensis (Ikäheimo et al., 2000; see protocol 

B.1.1). 

6.5.2	 Differentiation	of	species	and	subspecies 

Methods to discriminate subspecies of F. tularensis allow the distinction of isolates of subspecies tularensis from the other subspecies. This is crucial in the USA where both subspecies 

tularensis and subspecies holarctica occur concurrently. 

6.5.3	 Growth	characteristics 

F. tularensis subspecies tularensis, holarctica, and mediasiatica are slow-growing, fastidious 

organisms generally requiring sulfhydryl compounds for good growth. These three subspecies may be isolated from tissue on SBA. For subculture, cysteine/cystine-supplemented 

media are generally required as the organism usually fails to thrive with continued passage 

on SBA. F. tularensis subspecies novicida and F. philomiragia are both non-fastidious organisms 

and can be isolated on general microbiological solid media, including SBA. The presence of 

cysteine/cystine in the media can enhance the growth. Most isolates of F. philomiragia and 

F. tularensis subspecies novicida, but not other subspecies, can grow in nutrient broth containing 6% NaCl. 

6.5.4	 Biochemical	biotyping 

Differences in biochemical reactivity exist between the species and subspecies (Table 6.1) 

(Chu & Weyant, 2003; Sjöstedt, 2005). The oxidase test can be used to differentiate between 

F. tularensis and F. philomiragia. F. tularensis subspecies tularensis and holarctica differ in citrulline ureidase activity and glycerol fermentation and conventional biochemical assays can be 

utilized for biochemical differentiation based on glycerol fermentation or citrulline ureidase 

activity (Sandström et al., 1992). Alternatively, the automated system, BIOLOG, may also be 

used to detect glycerol fermentation (Petersen et al., 2004a). 

6.5.5	 Molecular	biotyping 

Molecular differences have been exploited to develop PCR subtyping assays. These PCR 

assays can provide F. tularensis subspecies identification based on size differences in amplified DNA products. Molecular subtyping assays are directed against an upstream region of 

an RNA helicase (the Ft-M19 assay), the presence or absence of the ISFtu2 insertion sequence 

element (ISFtu2 assay), the F. tularensis region of difference 1 (RD1 assay) and the pdpD gene 

(pdpD-2 assay) (Johansson et al., 2000c; Broekhuijsen et al., 2003; Johansson et al., 2004; 

Nano et al., 2004; Petersen et al., 2004a; Byström et al., 2005). Primer sequences and sizes of 

amplified gene products for the subspecies are given in Table 6.2 and protocol B.2. As many 

of these assays have only been validated with limited numbers of strains, it is recommended 

to verify the subspecies by an independent method. Real-time PCR assays have also been 

developed for classification of F. tularensis subspecies tularensis and F. tularensis subspecies 

holarctica (see protocol B.2.3). 

6.5.6	 Discrimination	of	strains 

Methods to discriminate strains of F. tularensis (i) provide the ability to trace the origin of 

isolates or the source of an outbreak and (ii) enable progress in epidemiological research. 

A multi-locus variable number tandem repeat assay (MLVA) for F. tularensis, based on 25 

different repeats in the genome, shows excellent discrimination of individual isolates and 

provides information regarding population structures and phylogenetic relationships within 

F. tularensis (Johansson et al., 2004). The method also allows accurate subspecies classification. At present, MLVA is the best method for obtaining high-resolution characterization of F. 

32
 
 

 

 

 

 

 

 

 

6. lABorAtory diAgnostics And discriminAtion oF suBsPecies And strAins 

Table	6.1 

Discriminating characteristics of Francisella species and subspecies 

Characteristic	 F. tularensis	subspecies	 F. philomiragia 

	 tularensis holarctica mediasiatica novicida 

Cysteine/cystine requirement + + + – – 


Carbohydrate fermentation:
 

Maltose + + – weak +
 

Sucrose – – – + +
 

D-glucose + + – + weak
 

Glycerol + – + weak –
 

Citrulline ureidase production + – + + nt
 

Oxidase productiona – – – – +
 

H2S production on TSI +
 

Cell size (μm) 0.2–0.7x0.2 0.2–0.7x0.2 0.2–0.7x0.2 0.7x1.7 0.7x1.7
 

a Tested with Kovacs reagent; +, positive; –, negative; nt, not tested; TSI, triple sugar iron agar. 

Table	6.2 

Polymerase chain reaction (PCR) methods to discriminate F. tularensis subspecies 

Assay	primer	sequence	(5’	to	3’)	 Product	size	(base	pairs) 

F. tularensis subspecies F. philomiragia 

tula holb medc novd 

Ft-M19	 F:aggcggagatctaggaaccttt 250 220 250 ±250 na 

R:agcccaagctgactaaaatcttt 

ISFtu2	 Tuf1705:gatagatacacgccttgctcaa; 390 1248 nt 390 na 

Tu B R431:acccagccaatgcctaaata 

pdpD-2	 F:tgggttattcaatggctcag 280 na nt 136e nt 

R:tcttgcacagctccaagagt 

RD1	 F:tttatataggtaaatgttttacctgtacca 1522 924 1453 3322e nt 

R:gccgagtttgatgctgaaaa 

a tularensis. b holarctica. c mediasiatica. d novicida. e Only one strain of F. tularensis subspecies novicida was tested. 

F, forward; R, reverse; nt, not tested; na, no amplification. 

tularensis. Analysis of 192 F. tularensis strains indicates that F. tularensis subspecies tularensis 

shows more genetic diversity than F. tularensis subspecies holarctica, suggesting the former 

subspecies to be older in evolutionary terms. Furthermore, two distinct clades, designated A.I 

and A.II, within subspecies tularensis have been identified (Johannson et al., 2004). Pulsedfield gel electrophoresis (PFGE) has also been used to define two subpopulations of type A 

(type A-east and type A-west) in the USA. A comparative PFGE and epidemiological analysis 

of human tularaemia in the USA has provided data suggesting that type A-west infections are 

clinically less severe than both type B and type A-east infections (Staples et al., 2006). 

6.6	 Biosafety	considerations 

Human error, poor laboratory techniques, and misuse of equipment cause the majority of 

laboratory-acquired infections. Safety measures in the laboratory are discussed in section 8.1. 

A compendium of technical methods to avoid or minimize such problems can be found in 

Part IV of the WHO Laboratory Biosafety Manual (World Health Organization, 2004b). 

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wHo guidelines on tulArAemiA 

6.6.1	 Diagnostic	laboratory-acquired	infections 

Personnel handling diagnostic cultures of F. tularensis are at considerable risk of infection and 

need to take precautionary measures. Due to the extremely low infectious dose, tularaemia 

has been one of the most commonly reported laboratory-associated bacterial infections (Pike, 

1976; Centers for Disease Control and Prevention, 2000; Shapiro & Schwartz, 2002). Since in 

most routine laboratories, cultures from clinical samples are handled at the bench, there is an 

obvious risk of exposure. It is therefore essential that laboratory personnel are informed of the 

possibility of tularaemia as a differential diagnosis when samples are submitted for diagnostic tests, to ensure that the appropriate biosafety guidelines are followed. F. tularensis may be 

present in lesion exudates, respiratory secretions, blood, tissues from infected animals, fluids 

from infected animals, and fluids from infected arthropods. Recognized laboratory hazards 

include contact of skin or mucous membranes with infectious materials, accidental inoculation, ingestion, and exposure to aerosols and infectious droplets. 

6.6.2	 Diagnostic	laboratory	requirements	(biosafety	level) 

The WHO Laboratory Biosafety Manual provides practical guidance on biosafety techniques 

for use in laboratories at all levels. For diagnostic laboratories, biosafety levels are also 

described in this Manual (World Health Organization, 2004b). For diagnostic laboratories 

biosafety level 2 practices, containment equipment, and facilities are recommended for activities involving clinical materials of human or animal origin suspected or known to contain 

F. tularensis. Biosafety level 3 and animal biosafety level 3 practices, containment equipment, 

and facilities are recommended, respectively, for all manipulations of suspect cultures, animal 

necropsies and for experimental animal studies. Preparatory work on cultures or contaminated materials for automated identification systems should be performed at biosafety level 3. 

6.6.3	 Decontamination	and	sterilization 

Information on decontamination and sterilization is available in chapter 8. 

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7. surveillAnce And outBreAk mAnAgement

 

 

 

 

 

 

 

 

7.	 Surveillance	and	outbreak	management 

mats Forsman, roland grunow, michael kosoy, gérard krause 

Tularaemia has been reported from most countries of the northern hemisphere. The 

epizootology of tularaemia is highly complex (Hopla, 1974). For instance, the distribution of 

cases in humans is patchy in the sense that, within a country or region, cases frequently occur 

at one location but are absent from surrounding areas. The reason for this distribution pattern is not yet fully understood. The differences in habitat and reservoir between subspecies 

tularensis and subspecies holarctica need to be considered in surveillance programmes and 

during outbreak investigations. Type A tularaemia (subspecies tularensis) occurs in the USA 

in relatively dry environments. It is highly virulent in humans but also in rabbits, sheep and 

other animals. Type B tularaemia (subspecies holarctica) occurs in Eurasia and in the USA. 

It is less virulent in humans and animals, although in humans it is often associated with 

severe complications. F. tularensis subspecies holarctica is mainly associated with streams, 

ponds, lakes, rivers, and semi-aquatic animals such as muskrats. F. tularensis is transmitted 

to humans by direct contact with infectious animals, arthropod bites, aerosols, or intake of 

contaminated food or water. Human-to-human transmission by aerosols or via arthropods 

has never been described. More information on the various vectors for transmission can be 

found in chapter 3. 

7.1	 Surveillance 

7.1.1	 Concept	of	surveillance 

Surveillance of disease is the continuing scrutiny of all aspects of occurrence and spread of a 

disease that are pertinent to effective control. Its main purpose is to detect changes in trend 

or distribution of diseases in order to initiate investigations or control measures. Surveillance 

generally uses methods distinguished by their practicability, uniformity, and frequently their 

rapidity, rather than by their complete accuracy. This includes the systematic collection and 

evaluation of morbidity and mortality reports, special reports of field investigations of epidemics and individual cases, isolation and identification of the infectious agent by laboratories, 

and information regarding immunity levels (including serological prevalence) in segments 

of the population. A report summarizing these data should be prepared and distributed to 

all persons cooperating in the surveillance and others with a need to know the results of the 

surveillance activities (Last, 1988). 

Surveillance needs to be based on clear case definitions (see section 6.1). As shown in analysing an outbreak of tularaemia in UN Administered Province of Kosovo (Serbia) in 1999– 

2000 (Reintjes et al., 2002), the interpretation of case counts may be complicated by unknown 

and/or rapidly changing population sizes. Case numbers need to be related to the population 

at risk of contracting the disease in order to compare disease occurrence over time or across 

geographical areas. Therefore, considerable effort needs to be made not only to collect reliable 

and complete data on cases but also to have recent population figures to hand. In outbreak 

situations, age-group and sex-specific incidence rates may provide hints to the source of infection. 

7.1.2	 Reasons	for	disease	surveillance 

A disease surveillance system, when adequately designed, allows not only the early detection 

of outbreaks and the identification of risk areas, but also assessment of the impact of actions 

targeted at reducing the size of the outbreak and at preventing further spread of the disease. 

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wHo guidelines on tulArAemiA 

Such documentation may do much to mitigate panic and to re-establish normality in a population affected by an outbreak (Dennis et al., 2001; World Health Organization, 2004a). In 

any disease surveillance system, awareness of the respective disease is crucial. This is more 

difficult for rare or sporadically-occurring diseases such as tularaemia (Dembek et al., 2003). 

A surveillance system may also provide scientific insight into the epidemiology of a disease, 

such as the distribution patterns of F. tularensis (Gurycova et al., 2001). 

7.2	 Surveillance	in	humans 

7.2.1	 Routine	surveillance	based	on	notifiable	disease	reports 

Mostly, public health surveillance is based on statutory reporting of notifiable diseases by 

physicians and/or laboratories to public health departments. This system allows the monitoring of large geographical areas at low cost and with relatively good geographical and 

socioeconomic representativeness. Reporting compliance among physicians and laboratories 

needs to be stimulated by regular information on the disease and easy reporting procedures. 

Reporting forms should be short, simple and unambiguous. A well-established notifiable disease reporting system may facilitate in-depth follow-up. More detailed and/or more reliable 

data may be collected by specially trained personnel. Such data may also include additional 

laboratory diagnoses. Tularaemia is a notifiable disease in many countries. 

7.2.2	 Sentinel	surveillance 

Sentinel surveillance relies on a set of reporting sources that agree to report all cases of one or 

more specific conditions. This is useful for diseases that are not notifiable and thus for which 

no routine surveillance system is in place. Sentinel surveillance may also be carried out in 

addition to routine surveillance in order to collect supplementary data. Due to a higher motivation and/or expertise of the reporting sources, sentinel surveillance is expected to provide 

qualitatively better data than routine surveillance based on notifiable-disease reports. However, due to a comparatively low number of reporting sources, the general population may not 

be well represented by the data collected. 

7.2.3	 International	surveillance	networks 

WHO has established an operational function (Global Outbreak Alert and Response 

Operations) to support the implementation of the international aspects of the International 

Health Regulations (IHR 2005; World Health Organization, 2006). This function relies on 

a systematic follow-up of information on suspected outbreaks and provision of support to 

outbreak response activities. This function includes: ”epidemic disease intelligence”, which 

is the pro-active collection of unverified information on possible outbreaks from all available 

sources (e.g. news wires, web sites, etc); ”outbreak verification”, which implies verifying the 

existence of an outbreak and is generally done through official counterparts in ministries of 

health or UN agencies; and ”outbreak response”, which implies providing technical assistance to contain the national and international public health consequences of outbreaks and 

is offered immediately by WHO, but conducted only upon request or acceptance of Member 

States. From 1 January 2001 to 15 November 2005, WHO monitored four events concerned 

with tularaemia. Two of these events involved infected pet animals and their shipment to other 

countries. The other two events involved higher than expected numbers of infections among 

humans in areas where tularaemia is known to occur naturally. For one of these events, WHO 

assisted in obtaining the needed diagnostic laboratory materials. 

7.3	 Surveillance	in	animals 

One of the main missions of the World Organisation for Animal Health (OIE), Paris, France is 

to report on the world animal health situation in all transparency. In order to fulfil its mandate 

in this respect, OIE manages the World Animal Health Information System, based on the 

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7. surveillAnce And outBreAk mAnAgement 

commitment of Member States to notify to OIE the main animal diseases, including zoonoses 

such as tularaemia (World Organisation for Animal Health, 2005a). Member States are committed to provide information (immediate notification and follow-up reports), six-monthly 

reports and annual information, as laid down in chapter 1.1.2 of the Terrestrial Animal Health 

Code entitled ”Notification and epidemiological information” (World Organisation for 

Animal Health, 2005b) which describes conditions and reasons for notification. 

The World Animal Health Information System has two components: first, the OIE Early 

Warning System which includes an alert procedure to warn the international community of 

exceptional epidemiological events in Member States. This alert system is aimed at the veterinary services of Member States and other stakeholders, enabling them to take any necessary 

protective measures as quickly as possible to prevent the introduction of pathogens originating from infected countries. Second, the OIE Monitoring System disseminates monthly and 

annual information provided by Member States on animal diseases and zoonoses (World 

Organisation for Animal Health, 2005a). In addition, the OIE publications Manual of Diagnostic Tests and Vaccines for Terrestrial Animals include standards for laboratory diagnostic tests 

for tularaemia and the production and control of biological products (principally vaccines) for 

veterinary use and contribute to the international harmonization of methods for the surveillance and control of this disease (World Organisation for Animal Health, 2005b). 

For zoonotic diseases such as tularaemia, surveillance in animals may prevent or minimize 

outbreaks in humans. Evidence in support of a tularaemia epizootic is a number of carcasses 

of water rats, mice, muskrats, hares, or rabbits, more than ”usual” (Mörner et al., 1988). Systematic surveys of natural foci of tularaemia would allow the early detection of an epizootic, 

but are highly demanding of labour and resources. Thus experience to date in efficacy of such 

approaches for tularaemia is very limited. Surveys in animals could be proposed in order to 

monitor changes in population structure and density of tularaemia susceptible lagomorphs 

and rodents. These surveys could be achieved by (i) systematic and directed investigation of 

susceptible mammals and arthropods in a region of interest; (ii) searching and testing carcasses and desiccated remnants (skin, bones) of dead animals; and (iii) examining water and mud 

samples collected close to places with dead animals or evident rodent activity. Contacting local 

mammalogists or parasitologists (academic, government) who have been engaged in research 

in the area may provide information about population trends or even archived blood or tissue 

samples for baseline determinations. In addition, these experts may provide invaluable logistical support and more “hands”. It could be of interest to investigate antibody prevalence in sera 

of carnivores if available, e.g. within the frame of other prevalence studies. Carnivores can also 

be indicators of F. tularensis prevalence as they are exposed by consuming infected moribund 

or dead animals. However, the utility of this approach needs to be investigated. 

7.4	 Outbreak	management	(humans) 

The main tasks of an outbreak investigation are to discover or confirm the causative agent of 

the outbreak and to limit the risk of exposure. In most cases, tularaemia outbreaks are selflimiting, but large outbreaks may occur under poor hygienic conditions. For instance, this was 

the case after the war in UN Administered Province of Kosovo (Serbia) in 1999. The situation 

was worsened by financial, social, and administrative constraints which made it difficult to 

prevent further spread of tularaemia (Reintjes et al., 2002). In this section, the requirements 

for an investigation of a tularaemia outbreak will be described. 

7.4.1	 Natural	outbreaks 

Epidemics in humans may occur after direct contact with infected animals (Greco & Ninu, 

1985; Andres-Puertas et al., 1999), vector transmission (Christenson, 1984; Markowitz et al., 

1985; Eliasson et al., 2002), oral intake of infected water (Greco et al., 1987; Rogutskii et al., 

1997; Anda et al., 2001; Pazdiora et al., 2002; Gurcan et al., 2004), or inhalation of bacteriacontaining aerosols or dust (Dahlstrand, Ringertz & Zetterberg, 1971; Syrjälä et al., 1985a; 

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Feldman et al., 2001). In some outbreaks, multiple modes of acquisition may be involved concomitantly (Lopez et al., 1982; Tikhenko et al., 2001). 

7.4.2	 Outbreaks	after	deliberate	release	of	Francisella tularensis 

Francisella tularensis has been considered as a possible biological warfare agent and has been 

stockpiled or otherwise weaponized by States forces (references in World Health Organization, 2004a). Since tularaemia occurs naturally, in most cases a deliberate attack would be 

rather difficult to differentiate from the natural occurrence of the disease (Grunow & Finke, 

2002). However, various “non-conclusive” criteria such as intelligence information may indicate indirectly the possible use of a biological warfare agent. The existence of a threat in terms 

of a political or terrorist environment or armed conflict, in connection with suspected or proven access to a biological agent and capabilities to deploy it as a weapon, could provide reasons 

to suspect an intentional release of Francisella. Further reasons to suspect a deliberate use of 

Francisella tularensis include: (i) the isolation and identification of genetically-manipulated 

Francisella or of genetic or biological variants uncommon in a given region (e.g. subspecies 

tularensis or mediasiatica as a causative agent in an outbreak in Europe would not be expected 

as a natural occurrence); (ii) an unusual antibiotic resistance in the pathogen; (iii) an unusual 

incidence of pneumonic or oropharyngeal manifestations which may be associated with a 

deliberate release of Francisella in aerosols or by contamination of food and drinking-water. 

The recent improvements in genetic identification of Francisella genomes and molecular typing may be helpful to differentiate between a deliberate release and natural occurrence of 

F. tularensis. See section 6.5. 

7.4.3	 Outbreak	investigation 

Situations in which an outbreak investigation may be helpful 

The possibilities to recognize and investigate a tularaemia outbreak depend on various factors such as the political and socioeconomic situation in the affected area and the availability 

of laboratory resources and specialists. In general, field investigations may be helpful in the 

following situations: 

— when the source of infection is unclear; 

— when the outbreak occurs as an emerging disease in non-endemic areas; 

— during the outbreak, when the effectiveness of measures targeted at the elimination of 

further sources of infection need to be verified or documented; 

— when there is a concern of a deliberate release of the pathogen; 

— when there are intentions and resources to collect more information about the epidemiology of tularaemia in the context of scientific research programmes. 

A checklist of practical advice on how to carry out an outbreak investigation can be found in 

Annex F. It should be emphasized that the best results and most benefit may be achieved when 

the investigation is initiated at the earliest possible stage of the outbreak. The most common 

approach is an epidemiological investigation, e.g. a case-control study, to identify risk factors. 

This was done during the outbreak in UN Administered Province of Kosovo (Serbia) in 1999 

(Reintjes et al., 2002). The study design and the questionnaire used in UN Administered 

Province of Kosovo (Serbia) are included in Annex F. 

Confirmation of the outbreak by verification of cases 

The confirmation of tularaemia cases should be based on the case definitions given in section 

6.1. Any epidemiological investigations should be based on these case definitions. Depending 

on the situation, it may be necessary to send samples for confirmation to a specialized laboratory in a different country. Specimens to be shipped to diagnostic laboratories require special 

attention to the safe packing and transportation of the material (see Annex E). 

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7. surveillAnce And outBreAk mAnAgement 

7.4.4	 Sample	collection 

In outbreak situations, an extended sample collection should be realized beyond the material 

that is required for direct diagnostic purposes. The aim is to detect the source(s) of infection, to determine the transmission route(s), and to perform the molecular characterization of 

the pathogen. Therefore, human diagnostic specimens as well as animal and environmental 

samples are helpful (see section 6.2). 

Dead animals 

When an outbreak is suspected or when an epizootic is recognized, collection of dead animals after die-offs becomes an important measure. The public should be encouraged by local 

authorities to report sick and dead animals observed. However, before examining dead animals for tularaemia, deliberate poisoning by humans must be ruled out as the cause of the 

die-off. When collecting dead animals, the following steps are recommended: 

1. Handle animals wearing thick rubber gloves. 

2. Place the carcass in double plastic bag. 

3. Label the bag containing the carcass. 

4. Transport the bag containing the carcass to the field station or to a laboratory authorized to 

accept specimens for tularaemia analyses. If a fresh carcass cannot be investigated within 

24 h, it should be frozen, thereby possibly enabling the isolation of F. tularensis later. 

Necropsy specimens from animals 

1. Tissues (fresh frozen, unpreserved) from lung, liver, spleen, lymph nodes, skin lesions and/ 

or kidney: suitable for culture in order to recover live organisms and for identifying DNA 

by PCR. When animals have died from tularaemia, the pathogen can be isolated most 

often from liver or spleen; tissues which are easy to obtain. 

2. Formalin-fixed tissue (not to be frozen) is suitable for histopathology, immunohistochemistry and PCR. Formalin-fixed specimens must be packaged separately from unpreserved autopsy specimens for bacterial isolation. 

Other samples 

Several other animal or environmental specimens have been subjected to attempts to identify 

the source of infection with tularaemia. However, systematic studies of the best methods of 

specimen collection and of the performance of the diagnostic methods applied to such specimens have not yet been done. Observations at the location where dead animals are found are 

often as important as the carcass itself to determine the source of tularaemia. Observations at 

the site of collection of a sample should be recorded with reference to collected samples, such 

as filters, surface swabs, dry materials, etc. In addition, the following may be helpful: 

1. Capture of animals/trapping of animals – this can be helpful in outbreak investigations to 

identify sources of infection and to type the pathogen when dead animals are not found. 

2. Animal excrement: extensive die-offs of tularaemia-susceptible animals during epizootics 

may render the capture of living animals and collection of sufficient samples more difficult. In this situation, collection and examination of animal excrement for detection of 

F. tularensis-specific antigen may be performed. 

3. Ticks from captured animals – for detection of F. tularensis, ticks may be collected from 

host mammals or with a drag. Removing ticks from captured animals is relatively simple 

and can be carried out simultaneously when collecting fleas and other ectoparasites. Whenever possible, ticks should be transported alive in moisturized tubes in an insulated box 

cooled with ice or cooling packs. If live transportation is not possible, transport ticks on dry 

ice. This may facilitate culture of F. tularensis. 

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wHo guidelines on tulArAemiA 

4. Water samples from streams and wells in the affected area – when carcasses of infected 

mammals that have died of tularaemia remain in streams or wells, the water may become 

contaminated with F. tularensis and may remain contaminated for more than a month. 

Mammals drinking the water may become infected and local epizootics may occur. Contaminated water may also be highly infectious for humans and domestic animals. 

7.4.5	 Laboratory	methods	suitable	under	field	conditions 

Hand-held assays for detection and identification of F. tularensis are currently being developed (Grunow et al., 2000; Emanuel et al., 2003). 

Depending on the location, size and duration of the outbreak, it can be helpful to establish 

a field laboratory. This may be a mobile laboratory which is brought in or a local laboratory which is additionally equipped for diagnosing tularaemia. A field laboratory for tularaemia should be able to perform microagglutination and/or ELISA for antigen and antibody 

detection (Syrjälä et al., 1986; Sjöstedt et al., 1997; Grunow et al., 2000; Porsch-Ozcurumez 

et al., 2004). In addition, simple instruments for genetic detection of pathogens under field 

conditions and biosensors are currently under development. The laboratory may also isolate 

F. tularensis, provided appropriate safety measures are in place (see section 8.1). In addition, 

the field laboratory will be the focal point for the collection and registration of the samples. It 

may also prepare samples for shipment to specialized laboratories for confirmation or further 

research such as molecular characterization (see section 6.4.4). 

7.4.6	 Measures	for	preventing	the	spread	of	tularaemia 

During a tularaemia outbreak, the following precautions may be taken: 

Oropharyngeal tularaemia 

Waterborne infection: 

— avoid drinking of unboiled water 

— disinfect (chlorinate) water which is used for washing or for brushing teeth 

— protect water sources from contact with animals such as rats, mice,: etc. 

Foodborne infection: 

— protect food stores from contact with animals (e.g. trap mice, rats or other rodents) 

— avoid eating food which may be contaminated with animal faeces 

— wash food with care as the aerosols and dust created can be infectious. 

Respiratory or ulceroglandular tularaemia 

Infectious aerosols, direct contact with infectious animals, and arthropod bites: 

— avoid the hunting of hares and rabbits and consumption of hare and rabbit meat; 

— wash hands after contact with wild and domestic animals; 

— regularly inspect domestic animals for signs of disease; in outbreak situations, avoid 

close contact with domestic animals such as dogs and cats; 

— avoid dust and aerosols (especially relevant for farmers and landscapers) by closing 

doors of tractors and avoid rooms where aerosols or dust are generated; if such exposure 

cannot be avoided, wear respiratory masks (protection class FFP3); 

— avoid exposure to blood-sucking arthropods by wearing long-sleeved clothing, and 

using repellents or mosquito nets. 

Although currently not available, vaccination of the human population against tularaemia in 

endemic regions would be most effective. 

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8. considerAtions For HAndling F. tularensis

8.	 Considerations	for	handling	F. tularensis 

may chu, karen elkins, Francis nano, richard titball 

8.1	 Safety	measures	in	the	laboratory 

Laboratory-acquired infections 

Although most naturally-occurring cases of tularaemia are the consequence of vector-borne 

transmission or from contact with infected animal tissues, the form of the disease which is 

of by far the greatest concern in the laboratory is respiratory tularaemia. This pathogen is so 

infectious by the airborne route that it has been stated that ”The hazard of infection with 

Francisella tularensis is well recognized; few persons escape illness if they continue to work 

with the organism” (Overholt et al., 1961). Studies in the USA with human volunteers during the 1950s and 1960s showed that the infectious dose of a strain of F. tularensis subspecies 

tularensis was between 10 and 50 CFU (McCrumb, 1961), making this bacterium one of the 

most infectious pathogens by the airborne route (Feldman et al., 2001). Prior to 1970, the 

incidence of laboratory-acquired tularaemia was high. For instance, in one laboratory working with F. tularensis subspecies tularensis the incidence of typhoidal tularaemia was reported 

to be 5.7 cases per 1000 employees at risk (Burke, 1977). In a detailed analysis of 34 cases of 

laboratory-acquired infection it was reported that 20 showed pulmonary involvement (Overholt et al., 1961). For the vast majority of these cases, there was no obvious previous exposure 

of the individual to the pathogen, but the high incidence of pulmonary involvement clearly 

suggest that most of these individuals contracted the disease after exposure to airborne bacteria (Overholt et al., 1961). The reductions in the incidence of laboratory-acquired tularaemia 

since the 1970s appear to be due partly to the use of more specialized laboratories with appropriate microbiological containment cabinets, partly to the increased awareness of the risk of 

infection, and partly to the use of the LVS vaccine (Rusnak et al., 2004). 

8.1.1	 Laboratory	biosafety	level 

There is no global agreement on the assignment of organisms to laboratory biosafety levels. 

WHO recommends that countries (and regions) should draw up a national (and regional) 

classification of microorganisms taking into account: (i) the pathogenicity of the organism; 

(ii) the mode of transmission and host range of the organism; (iii) the local availability of 

effective preventive measures; and (iv) the local availability of effective treatment. In most 

countries, the highly virulent subspecies tularensis will be classified as risk group 3 (high risk 

for the laboratory worker, but low community risk) while the other subspecies will be classified as risk group 2 (moderate risk for the laboratory worker, low community risk (World 

Health Organization, 2004b). 

The degree of risk varies not only with the virulence of the organism but also with the 

material being handled. We recommend that all strains of F. tularensis subspecies tularensis 

should be handled at biosafety level 3 while strains of subspecies novicida and the LVS 

strain of F. tularensis can generally be handled at biosafety level 2. It is recommended that all 

handling of clinical samples and cultures suspected to be F. tularensis should be performed 

under biosafety level 3 conditions. In addition, any work which involves the culture of humanvirulent strains of F. tularensis subspecies holarctica or subspecies mediasiatica where aerosols 

of the bacteria may be generated should be carried out at a minimum of biosafety level 3. 

A key consideration when handling F. tularensis is not only the physical construction of the 

laboratory, but also the experience of the individuals who will be handling the bacterium. 

Because of the high infectivity of F. tularensis by the airborne route, we recommend that any 

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individuals who plan to work with F. tularensis receive formal training in the methodologies 

and procedures required for the safe handling of this pathogen. In particular, individuals 

should be made aware of the possible sources of aerosols and the procedures which minimize 

the generation of aerosols. In addition, we recommend that new workers are supervised by an 

experienced individual during their learning phase. 

Since F. tularensis is one of the most infectious pathogens known, the importance of using 

appropriate biosafety practices and facilities cannot be overemphasized. Each laboratory 

should have defined procedures addressing the use of equipment (especially equipment that 

may generate aerosols); disinfection of equipment and contaminated materials; handling and 

processing samples; spill containment and clean-up; and waste handling. These procedures 

should be clearly and concisely written, easily accessible and rigorously followed. Guidance 

on biosafety level 3 containment and practices and disinfection is provided in the WHO 

Laboratory Biosafety Manual (World Health Organization, 2004b). 

8.1.2	 Physical	requirements	for	a	laboratory	handling	pathogenic	F. tularensis 

When handling cultures of any pathogenic F. tularensis subspecies, biosafety level 3 is prudent. A separate room is required with only one entrance; a biohazard notice prohibiting the 

entry of unauthorized persons should be prominently displayed at the entrance. Ideally, the 

room should have a double-door entrance designed to provide an airlock. The ventilation 

should be arranged to maintain the air pressure within the room at a slightly lower level than 

its surroundings. Air from the room should be discharged to the exterior, well away from air 

intakes and opening windows, otherwise it must be sterilized by filtration or heat treatment. 

The walls should be impermeable and all windows sealed to allow disinfestation and fumigation; it should be safeguarded against infestation with rodents or insects. The room must 

have a properly installed and tested Class II or III biological safety cabinet. The air exhaust 

from the cabinet should be so arranged as to avoid interference with the air balance in the 

room or within the cabinet when it is switched on. The room should have a sink, an autoclave 

and enough incubator space for all culture requirements. Hand-washing facilities must be 

provided near the exit. 

Biosafety cabinets should be used for all procedures involving human pathogenic strains 

(including F. tularensis strain LVS) and especially when aerosols of bacteria might be generated. 

These procedures include the growth of bacteria in liquid culture, the growth of large numbers 

of bacteria (e.g. for research purposes), procedures which couple significant mechanical energy 

into cells (for example sonication and centrifugation of cultures), or animal experimental 

work. See the WHO Laboratory Biosafety Manual (World Health Organization, 2004b) for 

appropriate and useful information on selection and use of biological safety cabinets. Ideally, 

items of equipment which might generate aerosols should be contained within a purposebuilt cabinet. 

Centrifuges may cause dangerous aerosols, especially when tubes containing virulent 

bacteria break. Glass tubes should not be used for virulent materials, instead polycarbonate 

tubes with tightly-fitting screw-capped lids and rubber O-rings are recommended. When 

centrifuges are located outside a biosafety cabinet, the centrifuge rotors should incorporate 

some form of seal that allows the rotor to be removed and opened only within a safety cabinet. 

Additionally, during centrifugation procedures access to the laboratory should be limited only 

to individuals wearing some form of respiratory protection such as a respirator. Respiratory 

protection should only be removed when the rotor has been opened, and the integrity of the 

centrifuge tubes has been established. 

8.1.3	 Decontamination	and	sterilization 

Materials which are potentially contaminated with F. tularensis should be sterilized before 

their disposal. The bacterium does not form resistant structures and is relatively sensitive to 

all standard inactivation procedures. Therefore the destruct cycle of inactivation used in auto42
 
8. considerAtions For HAndling F. tularensis 

claves is suitable for the inactivation of F. tularensis. The bacterium is sensitive to hypochlorite 

and other commonly-used chemical decontaminants. The thermal inactivation of the bacteria at temperatures below those experienced in an autoclave is possible, for example for the 

generation of killed cells for immunization, ELISA plate coating or the isolation of DNA or 

polysaccharides. Typically, heating to 60 °C for 1 h will inactivate bacteria in suspension, but 

the precise conditions for complete killing need to be ascertained in the laboratory. The temperature and time required for bacterial killing will depend not only on the strain being used 

but also on the density of the cell suspension and the suspending medium. Care must be taken with this type of low-temperature inactivation to ensure the statistically-significant possibility of killing all bacteria. Therefore it is necessary to first construct a killing curve which 

will inform the researcher of the conditions required for bacterial killing. Whatever method is 

used to inactivate F. tularensis, it is essential to ensure that samples leaving a biosafety level 3 

laboratory, for handling at a lower level of containment, are free of viable F. tularensis. 

F. tularensis is readily inactivated on exposure to UV irradiation. However, the sterilization 

of safety cabinets by exposure to an internal UV light source is not recommended as the 

primary method of bacterial killing. If there is a massive spill, safety cabinets used for the 

handling of F. tularensis should be sterilized using formaldehyde vapour, in accordance with 

the manufacturer’s instructions. Formaldehyde decontamination is not necessary for routine 

work. 

8.1.4	 Health	monitoring	of	laboratory	workers 

Some form of health monitoring is essential for staff working with F. tularensis. Baseline blood 

samples should be taken before any work commences and staff should carry an “at risk” card 

indicating that in the event of a febrile illness, accidental exposure to F. tularensis should be 

considered by the attending clinician. In the event of possible exposure to F. tularensis, the 

subsequent development of a fever is likely to provide one of the earliest indications of infection. For more details on the symptoms of tularaemia see chapter 4. 

8.1.5	 Treatment	following	a	suspected	exposure 

Following likely exposure of an individual to F. tularensis, appropriate chemotherapy should 

commence immediately; at least 14 days of treatment is required. However, the isolation of 

the individual is not necessary since human-to-human transmission of the disease has not 

been reported. For details of chemotherapeutic regimes for the treatment of tularaemia see 

chapter 5. 

8.2	Vaccines	and	vaccination 

One focus of current research work in the USA and in Europe is to devise a vaccine which could 

be used to protect individuals from disease caused by the deliberate release of F. tularensis. 

This vaccine would also have a utility for the protection of researchers who are working with 

highly-virulent strains of F. tularensis. Vaccine development was initiated in the 1930s in the 

former Soviet Union. The most widely studied and used vaccine strain in recent times has 

been the attenuated Live Vaccine Strain (LVS), which was derived from a virulent strain of 

F. tularensis subspecies holarctica. The precise history of the LVS strain is uncertain but it is 

known to have been derived from strain 15, which was originally generated at the Gamaleya 

Institute in the former Soviet Union. A mixed ampoule of vaccine strains, probably including 

strain 15, was transferred to the USA in 1956, and the LVS strain was probably derived from 

this strain in the USA (Eigelsbach & Downs, 1961; Sandström, 1994). In the past, F. tularensis LVS has been studied as an investigational vaccine by the United States Department of 

Defence (Waag et al., 1992; Waag et al., 1995; Waag et al., 1996) but is not licensed for general 

use in the USA (Dennis et al., 2001). It is also unavailable in Europe and most other parts of 

the world, but a live attenuated vaccine is still in use some parts of the former Soviet Union 

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where it has been used to immunize millions of people against tularaemia (Sjöstedt, Tärnvik 

& Sandström, 1996). It is possible that future studies will allow either the LVS vaccine or 

alternative attenuated strains to be licensed. Therefore it is appropriate to provide a summary 

of the properties of the LVS vaccine. 

Of the animal species which have been evaluated as models of disease, the non-human 

primates and rabbits appear to mimic human disease most accurately. In contrast, mice appear 

to be uniformly susceptible to disease caused by strains belonging to all subspecies, including 

F. tularensis subspecies novicida. Much of the research on mechanisms of virulence and 

protection which has been carried out in recent years has used the LVS strain of F. tularensis in 

mice. This model of disease is valuable because protective immune responses can be induced 

by the delivery of this strain intradermally or by scarification while mechanisms of virulence 

can be investigated when the strain is given by the intraperitoneal or intravenous routes. 

Many workers have combined these different routes to investigate the protective responses 

induced by LVS against a subsequent intraperitoneal or intravenous challenge with LVS. In 

addition, the risk of serious laboratory-acquired infection is significantly reduced when the 

LVS strain is used rather than a highly-virulent strain. 

However, it is also important to consider the limitations of this disease model. There is 

good evidence that protection against an intraperitoneal or intravenous challenge with LVS 

can be mediated solely by antibody directed against LPS, whereas protection against highlyvirulent strains of F. tularensis in mice requires CD4+ and or CD8+ T-cell responses. There is 

also good reason to believe that these findings reflect the nature of the protective responses 

in humans; previous studies have shown that the passive transfer of immune serum provides 

only very limited protection against tularaemia. 

The nature of the attenuating lesion(s) in the LVS strain is not known. When delivered 

intradermally into the mouse, LVS is relatively avirulent and by the subcutaneous route 

the median lethal dose is approximately 107 CFU (compared with < 10 CFU reported for 

virulent F. tularensis subspecies tularensis and subspecies holarctica strains). However, by the 

intraperitoneal route, LVS has a median lethal dose of < 10 CFU (Fortier et al., 1991). The 

reasons for this route-specific attenuation in the mouse are not known. In addition, different 

batches of the vaccine show different immunogenicity, a property ascribed to the different 

proportions of so-called blue and grey colony types (Eigelsbach & Downs, 1961). 

Clinical trials with the LVS vaccine have not been carried out, but there are some data from 

studies with small numbers of individuals which suggest that vaccination by the intradermal 

route is able to either provide protection against a subsequent challenge with fully virulent 

F. tularensis or to reduce the severity of the disease (Saslaw et al., 1961a; Saslaw et al., 1961b). 

There is some evidence that the vaccine is effective when given by the airborne and oral routes 

of delivery (Saslaw et al., 1961a; Saslaw et al., 1961b; Hornick & Eigelsbach, 1966; Hornick et al., 

1966). Trials in the USA during the 1950s and 1960s involved volunteers who were immunized 

with LVS and approximately one year later challenged by the airborne route with 200–20 000 

CFU of F. tularensis strain SCHU S4. All of the control subjects developed tularaemia and at the 

earliest indication of disease were treated with streptomycin or tetracycline. All made a complete 

recovery from the disease. The majority of vaccinees challenged with up to 2000 organisms 

escaped major clinical illness. Immunized volunteers challenged with 20 000 organisms 

showed modified disease symptoms compared to non-immunized volunteers infected with a 

similar dose of SCHU S4 (Eigelsbach, Hornik & Tulis, 1967). In addition to these studies there 

is evidence that the LVS vaccine has had an impact on the incidence of laboratory-acquired 

tularaemia. A study published by Burke in 1977 reported the incidence of laboratory-acquired 

tularaemia before (1950–1959) and after (1960–1969) routine immunization of workers. The 

incidence of typhoidal tularaemia fell from 5.7 to 0.27 cases per 1000 employees at risk. The 

incidence of ulceroglandular tularaemia was unchanged but the clinical signs and symptoms 

of this form of the disease were moderated in vaccinated individuals. Vaccination of laboratory 

workers may reduce the risk of laboratory-acquired infections (Rusnak et al., 2004). 

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8. considerAtions For HAndling F. tularensis 

At this time it is not clear whether or when the LVS vaccine will be licensed. However, 

the data presented above indicate the value of this vaccine for the prevention of laboratoryacquired tularaemia. Should the LVS vaccine not be licensed, then development of an 

alternative licensable vaccine may require considerable time. The protective components of 

the LVS vaccine have not yet been identified, and very little is known about mechanisms of 

virulence. A rationally attenuated mutant to replace the LVS vaccine seems to be the most 

likely solution and some of the pathways that might be interrupted to provide attenuation 

have already been identified (Karlsson et al., 2000). However, techniques for the construction 

of defined mutants of F. tularensis are at an early stage and will need to be further developed 

before a rationally attenuated mutant can be devised and tested. 

8.2.1	 Genetic	manipulation	of	F. tularensis 

Although techniques for the genetic manipulation of F. tularensis are in their relative infancy, it 

is important to consider which replicons and antibiotic markers are acceptable for use. Genes 

encoding resistance to antibiotics that might be used prophylactically or therapeutically 

should be used with caution, and a full assessment of the risk associated with such manipulations carried out with special reference to the availability of alternative antibiotics for the 

treatment of disease. F. tularensis is naturally resistant to many beta-lactam antibiotics and 

therefore ampicillin resistance is not a useful marker for genetic manipulation of strains. 

Similarly, many strains are resistant to erythromycin, although this marker may be useful in 

erythromycin-susceptible strains. The markers used most frequently in F. tularensis include 

chloramphenicol resistance, tetracycline resistance and kanamycin resistance. Of these, tetracycline and kanamycin resistance markers offer the best selection. Because the tetracyclines 

are often used as drugs of first choice, the use of tetracycline resistance as a marker may not be 

acceptable in all countries. Additionally, care should be taken over the selection of kanamycin 

resistance markers. It is important to use a gene that encodes kanamycin-specific resistance 

rather than one which provides resistance to a range of aminoglycosides since this might 

provide resistance to streptomycin and gentamicin. The kanamycin resistance gene found in 

transposon Tn5 is kanamycin-specific and can be used in F. tularensis. 

8.2.2	 Sources	of	strains	for	research 

Although strains of F. tularensis are in principle available from a number of national culture 

collections, the release of strains from these sources is now carefully controlled. In some 

countries additional legislation operates to regulate the acquisition of strains of F. tularensis 

from any source. For example, in the Russian Federation the export of killed or live bacterial 

cells or even of DNA to any other country is prohibited. The stringent controls on access are 

currently the subject of considerable discussion. For example, in the USA the American Type 

Culture Collection (ATCC) is currently reviewing the procedures for the release of F. tularensis 

to researchers. It is recommended that researchers should attempt to obtain strains from culture collections or other researchers working in their own country in the first instance. 

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9. reFerences

9.	 References 

Abd H et al. (2003). Survival and growth of Francisella tularensis in Acanthamoeba castellanii. 

Applied and Environmental Microbiology, 69:600–606. 

Akdis AC, et al. (1993). Immunological evaluation of erythema nodosum in tularaemia. British 

Journal of Dermatology, 129:275–279. 

Algazin IP, Bogdanov II (1978). The role of lemmings in natural tularemia foci in the Trans 

Polar region. Zhurnal mikrobiologii, epidemiologii, i immunobiologii, 2:93–97. 

Anda P et al. (2001). Waterborne outbreak of tularemia associated with crayfish fishing. 

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Annexes
 

A. diAgnostic Protocols

A.	 Diagnostic	protocols 

The following protocols have been provided by the University of Umeå, Umeå, Sweden, Centers for Disease Control and Prevention, Fort Collins, CO, USA, the Bundeswehr Institute of 

Microbiology, Munich, Germany, and the Robert Koch Institute, Berlin, Germany. 

Biosafety. Personnel handling diagnostic cultures of F. tularensis are at considerable risk of 

infection and need to take precautionary measures. Biosafety Level 3 and Animal Biosafety 

Level 3 practices, containment equipment, and facilities are recommended, respectively, for 

all manipulations of suspect cultures, animal necropsies and animal studies. 

Collection and transport of samples. Every effort should be made to collect and preserve specimens so that viable bacteria can be recovered. Decontaminate the surface area prior to specimen collection. Ensure that adequate volumes are collected to avoid false negatives as a result 

of insufficient sample volume. To minimize loss in viability, specimens should be transported 

to the laboratory within 24 h. For collection and transportation procedures for specific specimen types, see section 6.2.5. 

Note. Some of the protocols described below have been developed for use with the reagents 

listed. Reagents other than those listed in the protocol should be properly validated prior to 

use. 

A.1	Gram	stain 

Perspective. The Gram stain can be used for suspect cases of Francisella tularensis. Poorlystaining, Gram-negative (pink) short rods or coccoid forms from a patient with exposure and 

clinical symptoms compatible with tularaemia are considered suspect for F. tularensis. 

Application. Smears may be prepared from fresh, unpreserved specimens including cultures, 

tissues, and primary specimens in which the number of organisms is expected to be high. 

Reagents. Gram stain kit. 

Equipment/supplies 

1. Microscope slides. 

2. Gas or alcohol burner. 

3. Staining rack for slides. 

4. Microscope with high power and oil immersion objectives. 

Procedure 

1. Smear a thin layer of the specimen onto the slide. Smears must be made with a light concentration of cells in order to visualize individual cells clearly. Thick smears containing too 

many cells may give an indistinct reading. 

2. Let the smear air dry, then heat-fix the smear by passing the slide through a flame, then let 

the slide cool. 

3. Cover the smear with crystal violet for 1 min, followed by iodine for 2 min; rinse with tap 

water. 

4. Decolorize the smear with ethanol for 10 s; rinse with tap water. 

5. Counterstain the smear with safranin for 1 min; rinse with tap water. 

6. Allow the slide to air dry then examine using the oil immersion objective on the microscope. 

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Interpretation. In Gram-stained smears, F. tularensis appears as a very tiny (0.2–0.7 μm x < 1.0 

μm), pleomorphic, poorly-staining Gram-negative (pink) short rods or coccoid forms usually 

seen as single cells. 

Quality control. Staining of test specimens should be carried out in parallel with known Grampositive and Gram-negative organisms to ensure proper staining results. Reagents should be 

fresh and the crystal violet stain kept free of contamination. Microscope lenses and objectives 

should be kept dust and oil-free. 

A.2	Culture	on	agar 

Perspective. Culture is considered the “gold standard” for F. tularensis. Cystine heart agar 

enriched with chocolatized red blood cells (CHAB) is a practical medium for culture of F. tularensis as the organism displays characteristic morphology (green, opalescent, raised, shiny 

colonies). Other enriched media such as GCII agar base with 1% haemoglobin and 1% IsoVitaleXTM, Mueller-Hinton with 1% IsoVitaleXTM, buffered charcoal yeast extract (BCYE), or 

thiogylcollate-glucose blood agar (TGBA) can also be used for the cultivation of F. tularensis, 

however, the colony morphology is not distinctive on these media. For contaminated specimens, antibiotic-supplemented media are recommended. 

Application. All clinical specimens. 

Media 

1. Cystine heart agar base with 9% chocolatized sheep blood (see section C.1). 

2. Mueller-Hinton agar with 1% IsoVitaleXTM. 

3. Thioglycollate-glucose blood agar (TGBA). 

4. GCII agar base with 1% haemoglobin and 1% IsoVitaleXTM (see section C.2). 

5. Chocolate agar. 

6. Buffered charcoal yeast extract (BCYE). 

7. CHAB-A: CHAB supplemented with colistin 7.5 mg, amphotericin 2.5 mg, lincomycin 

0.5 mg, trimethoprim 4 mg and ampicillin 10 mg per l. 

Equipment/supplies 

1. Sterile bacteriologic loops. 

2. Incubator: 37 °C. 

Procedure 

1. Use established inoculation and plating procedures for various clinical specimens. 

2. Incubate cultures at 35–37 °C; ambient atmosphere or 5% CO2

 is acceptable. 

3. Incubate the primary plates for 7–10 days. 

Interpretation. On general media such as chocolate agar, or BCYE, F. tularensis is a greywhite, 

opaque colony, usually too small to be seen at 24 h on primary isolation from clinical specimens. After incubation for 48 h or more, colonies are about 1–2 mm in diameter, white to grey 

to bluish-grey, opaque, flat, with an entire edge, smooth, and have a shiny surface. On CHAB, 

after 48 h the colonies are 2–4 mm in size, greenish-white, dense with a butyrous consistency. 

A characteristic opalescent sheen is evident on the surface of the colonies, especially if the 

plate has been incubated for 48–72 h. F. tularensis will not grow on MacConkey agar. 

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A. diAgnostic Protocols 

Figure	A.1	 F. tularensis on CHAB agar after 48 h incubation. Image provided by the Centers for Disease Control 

and Prevention, Fort Collins, CO, USA 

Quality control. Each batch of plates must be checked for sterility by incubating plates at 37 °C 

for at least 24 h. Each batch of plates must be checked for the ability to support growth of 

F. tularensis. 

A.2.1	 Mouse	inoculation 

Perspective. Inoculation of laboratory mice is a useful method for recovery of F. tularensis from 

contaminated specimens. Prior to any work with animals in the laboratory, protocols must be 

reviewed and approved by the appropriate animal use committee. 

Application. Specimens or cultures contaminated with other organisms. 

Materials 

1. Specimen suspected to contain F. tularensis. 

2. Pathogen-free mice (Swiss-Webster outbred mice, 6–9 weeks old). 

3. Saline (0.85% NaCl). 

4. Sterile sand or latex particles. 

Facilities/supplies/equipment 

1. Animal-care facilities (negative-pressure room). 

2. Sterile mortar and pestle. 

3. Syringes (1 ml) with 25 x 1/2” needles. 

4. Sterile gloves. 

5. Discard container (’Sharps bin’) for needles. 

6. Sterile scissors, forceps. 

7. Alcohol, 70%. 

8. Sterile Petri dish or shallow container. 

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wHo guidelines on tulArAemiA 

Procedure 

1. Sample preparation: tissue specimens are prepared by excising small pieces into a mortar 

with a few drops of saline. A small amount of sterile abrasive material (sand or latex) is 

added to the mixture in the mortar before grinding with the pestle. Cultures are resuspended in saline and mixed prior to inoculation. Aspirates and respiratory specimens should 

be diluted in saline only if too viscous to inoculate. 

2. Inoculation of the animal: inject 0.1–0.5 ml of the prepared samples subcutaneously in the 

lower abdomen. Typically, two mice are each injected with 0.1 ml of prepared inoculum 

from culture or other purified material, whereas for tissue specimens, up to 0.5 ml of suspension should be injected. 

3. Observation: mice are checked twice daily for signs of illness. Surviving mice are kept for 

3 weeks (21 days) before reporting as negative. 

4. Processing mouse tissue: all moribund mice are sacrificed. Animals are necropsied and 

spleen and liver tissues are removed for culture on agar plates. 

5. Interpretation: F. tularensis is easily isolated when passaged through susceptible, laboratory-raised, specific-pathogen free mice. Inoculated mice typically become ill within the 

first 3–4 days post-inoculation. Tissue samples (liver, spleen) taken from the mice are good 

sources of nearly pure cultures of F. tularensis as mice will selectively amplify pathogenic 

strains of F. tularensis. Recovered liver and spleen are enlarged and haemolytic and sometimes contain multiple focal abscesses. 

Quality control. Since the mouse inoculation test is dependent on visible morbidity, mouse colonies should be inoculated with a known virulent F. tularensis isolate to test their sensitivity 

to infection prior to use. Sacrificed mice must be processed as quickly as possible or stored 

in the freezer at -20 °C to prevent deterioration of the material. When performing necropsy 

of the mice, take care not to cause unwanted enteric contamination by touching the lower 

abdominal surfaces. Use a freshly cut tissue for culture. 

A.3	Antigen	detection 

A.3.1	 Direct	fluorescence	assay	(DFA) 

Perspective. The direct fluorescent antibody (DFA) assay is based on the interaction between 

fluorescently (FITC)-labelled rabbit polyclonal anti-F. tularensis antibody and F. tularensisspecific antigens. This antibody is generated against whole killed F. tularensis cells. DFA is a 

rapid assay that can be used for both presumptive and confirmatory detection of F. tularensis. 

The DFA test is considered presumptive when a primary specimen is DFA-positive and confirmatory when a recovered isolate is DFA-positive. 

Application. Slides may be prepared from cultures, ulcer swabs, tissues, blood and sputum 

specimens. 

Reagents 

1. FITC-labelled rabbit anti-F. tularensis conjugate (CDC, Fort Collins, CO; section D, Table 

D.1). 

2. Phosphate-buffered saline (PBS). 

Equipment/supplies 

1. Microscope slides, two-etched ring. 

2. Gas or alcohol burner. 

3. Coplin jars or other jars capable of holding slide/s. 

4. Fluorescence microscope, with FITC filter. 

64
 
A. diAgnostic Protocols 

Procedure 

1. Primary specimens: smear a thin layer of the specimen onto the slide. Smears must be 

made with a light concentration of cells in order to visualize individual cells clearly. Thick 

smears containing too many cells may give an indistinct reading. 

2. Cultures: from a 24 h culture, use a 1 μl inoculating loop to pick up cells in the 3rd quadrant 

of growth. Prepare a suspension equivalent to 0.5 McFarland standard in saline. Mix well. 

Use a sterile swab to smear onto the slide. 

3. Let the smear air dry. 

4. Heat-fix the smear by passing the slide through a flame and then allowing it to cool. 

5. Add 20 μl of conjugate (enough to cover the smear area) and incubate for 30 min at ambient 

temperature. Use a humidified chamber for incubation to prevent evaporation. 

6. Fill a Coplin jar with PBS and soak the slide for 10 min. Discard the PBS from the container 

and refill with fresh PBS. Soak the slide for a further 10 min. 

7.	 Remove the slide from PBS and rinse in distilled H2O to remove remaining salt. 

8. Allow the slide to air-dry. 

9.	 View the slide using a fluorescence microscope and FITC filter; slides can be analysed using 

40X, 63X or 100X objectives. 

Interpretation. A positive DFA result is seen as bright, intense green staining over the entire 

surface of the bacterial cell. F. tularensis cells are tiny, (0.2–0.7 x ≤ 1.0 μm) pleomorphic cells 

and generally appear as single cells in short rod or coccoid form. 

Quality control. Each DFA test should be performed in parallel with a positive and negative 

control to ensure proper interpretation. Positive and negative controls can be prepared in 

advance and stored at -20 °C for future use. Process controls and test samples together. To 

maintain integrity of the conjugate, avoid freeze-thawing FITC-labelled antibodies. Upon 

receiving reagents, small aliquots should be made and the stock stored (in the dark) at -20 °C. 

The reagent should be checked annually to ensure reactivity. 

Figure	A.2	 Isolate of F. tularensis stained by the direct fluorescence method (magnification 630X). Image provided by Centers for Disease Control and Prevention, Fort Collins, CO, USA. 

65
 
wHo guidelines on tulArAemiA 

A.3.2	 Slide	agglutination 

Perspective. The slide agglutination assay is based on the interaction between high-titre antiF. tularensis antibody and F. tularensis-specific antigens. This interaction produces immediate 

clumping of the cells, visible to the naked eye. A positive slide agglutination test on an isolate, 

with characteristic F. tularensis growth on agar, is confirmatory for F. tularensis. 

Application. Suspect cultures of F. tularensis. 

Reagents 

1. Hyperimmune rabbit-anti F. tularensis antibody (Difco, Becton Dickinson and Company; 

Annex D). 

2. Normal rabbit serum. 

3. Suspect culture. 

Equipment/supplies 

1. Bacteriological loops, 1 μl size. 

2. Glass slides, two-etched ring. 

3. Pipettor, 10–200 μl. 

Procedure 

1. Mark and section slide with wax pencil into two compartments. 

2. Place one drop of positive serum in one compartment and one drop of negative serum in 

the next compartment. 

3. Add equal amounts (i.e. one small loopful) of isolate to the positive and negative control 

serum; make sure cells are resuspended evenly. 

4. Rock slide back and forth for 1 min, or according to manufacturer’s recommendations. 

5. Examine spots for visible clumping. 

6. Discard the used slide in the proper disposal container and decontaminate by autoclaving. 

Interpretation. A positive reaction (clumping of cells) is visible if the suspect culture is F. tularensis. A negative result is visualized as an absence of clumping. Each test should be performed in parallel with a positive and negative control to ensure proper interpretation. Process 

controls and test samples together. 

Quality control. To ensure specificity, positive and negative reagents may be tested against 

other killed bacterial cell preparations such as Brucella spp., Yersinia spp., Bacillus spp., and 

Escherichia coli. 

A.3.3	 Capture	enzyme-linked	immunosorbent	assay	(cELISA) 

Perspective. The capture enzyme-linked immunosorbent assay (cELISA) is based on antigen 

binding to a monoclonal antibody (mAb) coated onto a solid phase (microtitre plate) and 

detection of the bound antigen using the same mAb (peroxidase labelled) (Grunow et al., 

2000). The mAb is specific for the LPS of F. tularensis, including subspecies tularensis, holarctica, and mediasiatica. The cELISA is considered confirmatory when a recovered isolate, with 

characteristic growth on agar, is positive. Either whole cells or LPS extracted cells (final dilution of at least 1:4) can be used in the assay. The sensitivity of the assay is about 104

–103 CFU/ 

ml. 

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A. diAgnostic Protocols 

Figure	A.3	 Slide agglutination for F. tularensis +, positive serum; –, negative serum. Image provided by Centers 

for Disease Control and Prevention, Fort Collins, CO, USA. 

Application. This protocol has been developed primarily for confirmation of suspect cultures 

and presumptive testing of animal tissues. 

Reagents (sufficient for one microtitre plate, i.e. 40 unknown samples) 

1.	 Monoclonal antibody anti-F. tularensis (e.g. Senova GmbH, Germany, Annex D). 

2.	 Monoclonal antibody anti-F. tularensis-HRP conjugate (e.g. Senova GmbH, Germany, 

Annex D). 

3.	 Positive control (e.g. F. tularensis LVS OD = c. 109 cells/ml), 0.01% thiomersal added 560nm

for long-term storage, optional: solubilization of LPS using LPS extraction buffer. 

4.	 Tetramethyl benzidine (TMB) ready to use (12 ml). 

5.	 Phosphate buffered saline (PBS), isoton, pH 7.2, 1l. 

6.	 Dry skimmed milk. 

7.	 Tween-20. 

8.	 Coating buffer: sodium carbonate pH 8.5–9, 0.025 M. 

9.	 0.25 M sulphuric acid. 

10.	 Optional: thiomersal 1%. 

11.	 Optional: LPS extraction buffer 10 ml (Chlamydia Assay, Abbott, Solna, Sweden, or prepared to the following formulation: for 100 ml, chenodeoxycholic acid 0.099 g EDTA trisodium salt 0.354 g, sodium azide 0.100 g, sodium chloride 0.231 g, disodium hydrogen 

phosphate-7-hydrate 1.764 g, sodium hydrogen phosphate 0.454 g, add to 100 ml distilled 

water). 

Equipment/supplies 

1.	 ELISA-reader. 

2.	 8 or 12 multi-channel pipette. 

3.	 Reservoirs to handle fluids for multi-channel pipette. 

4.	 1 channel pipette 10–100 μl. 

6.	 Microcentrifuge tubes 1.5 ml. 

7.	 1l glassware, 1 l and 10 ml beakers or tubes. 

8.	 10 ml pipettes and appropriate pipettor. 

9.	 ELISA-Washer (optional). 

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wHo guidelines on tulArAemiA 

10. Incubator at 37 °C. 

11. Water-bath at 8 °C. 

12. Water-bath at 60 °C. 

13. pH meter. 

Preparation of solutions (to process one plate) 

1. The wash buffer is prepared by mixing PBS powder in 1 l distilled water (0.15 M) and 0.5 ml 

Tween-20, and adjusting the pH to 7.4 (this buffer can be stored at room temperature for 3 

days). 

2. The blocking buffer is prepared by adding 1.0 g of dry skimmed milk (4%) to 25 ml wash 

buffer (prepared fresh before use). 

3. The dilution buffer is prepared by mixing 10 ml of blocking buffer with 30 ml of wash 

buffer (prepared fresh before use). 

4. The coating buffer is prepared by adding 0.8 g sodium carbonate and 1.46 g sodium hydrogen carbonate to 250 ml distilled water, 0.025 M. The pH should be adjusted to pH 8.5– 

9.0. 

Procedure (for one plate, up to 40 unknown samples) 

1. Dilute stock solution of mAb to a final concentration of 5μg/ml in 10.5 ml coating buffer. 

2. Add 100 μl of diluted mAb per well and cover with cover tape or plate seal. 

3. Incubate for 1 h at 37 °C or overnight at 4 °C. 

4. Preparation of unknown samples: Liquid samples should be diluted 1:2 with dilution 

buffer. Solid samples should be homogenized in PBS and after sedimentation the liquid 

phase should be diluted 1:2 with dilution buffer. There are several protocols to improve the 

homogenization of solid samples and the extraction of bacterial antigens. Here, a simple 

protocol is proposed (optional): 

Liquid sample:
 

a) mix 150 μl unknown sample with 150 μl LPS extraction buffer in an microcentrifuge 


tube 

b) heat the tubes for 30 min in a water-bath at 60 °C. 

Solid sample: 

a) place sample (0.2 g) in a Petri dish or other appropriate container 

b) add 0.5 ml PBS 

c) slice the sample into fine pieces, homogenize and pipette the supernatant into a 1.5 ml 

microcentrifuge tube 

d) add 0.7 ml LPS extraction buffer 

e) incubate samples at 60 °C for 30 min 

f) vortex, and allow the sediment to settle; if the sediment does not settle, filter or spin at 

low speed (< 100 x g) 

g) carefully remove 200 μl of the sample and dilute with 200 μl (1:2) dilution buffer (make 

further dilution, if required) 

h) samples are now ready for analysis. 

68
 
A. diAgnostic Protocols 

5.	 Prepare positive and negative controls in microcentrifuge tubes: 

a) mix 30 μl of positive control with 270 μl dilution buffer (equivalent 108

 CFU/ml) 

b) take 10 μl from 108 CFU/ml and mix with 990 μl dilution buffer (equivalent 106 CFU/ 

ml) 

c) take 200 μl from 106 CFU/ml and mix with 400 μl dilution buffer (equivalent 3 x 105 

CFU/ml = positive control (PC1) 

d) take 50 μl from PC1 and mix with 450 μl dilution buffer (equivalent 3 x 104 CFU/ml = 

PC3) 

e) take 50 μl from PC3 and mix with 450 μl dilution buffer (equivalent 3 x 103 CFU/ml = 

PC5) 

f) take 50 μl from 106 bacteria/ml and mix with 450 μl dilution buffer (equivalent 1 x 105 

bacteria/ml = PC2) 

g) take 50 μl from PC2 and mix with 450 μl dilution buffer (equivalent 1x 104 CFU/ml = 

PC4) 

h) take 50 μl from PC4 and mix with 450 μl dilution buffer (equivalent 1x 103 CFU/ml = 

PC6) 

i) dilution buffer only = negative control (NC). 

6.	 Wash the plate twice using 300 μl washing buffer per well. 

7.	 Block the plate with 125 μl blocking buffer per well for 30 min at 37 °C. 

8.	 Wash the plates again as in step 6. 

9.	 Add 100 μl volumes of the controls and unknown samples as duplicates to the appropriate 

wells. 

10.	 Incubate for 1 h at 37 °C. 

11.	 Wash the plates four times as in step 6. 

12. Add 100 μl of detector mAb-HRP-conjugate (10μg/ml) to all wells except blank wells. 

13.	 Incubate for 1 h at 28 °C. 

14.	 Wash the plates five times as in step 6. 

15.	 Add the substrate TMB 100 μl per well and observe for approx. 7 min (up to 15 min) for 

colour reaction. 

16.	 Stop the reaction by adding 100 μl of sulphuric acid. 

17.	 Read the optical density (OD) of the samples at 450 nm (reference wavelength 620 nm). 

18.	 Create a standard curve from the control samples and read the bacterial load of the 

unknown samples, if required. 

69
 
 

wHo guidelines on tulArAemiA 

Plate	scheme 

Unknown samples 

1	 2	 3	 4	 5	 6	 7	 8	 9	 10	 11	 12 

A Blank Blank 

B PC1 PC1 

C PC2 PC2 

D PC3 PC3 

E PC4 PC4 

F PC5 PC5 

G PC6 PC6 

H NC NC 

Interpretation. Negative specimens will show an OD close to that of the negative control (usual 

cut-off OD < 0.2). OD of positive specimens and controls ranges from 0.2–1.5. A quantification of the bacterial load can be done using the standard curve. 

Quality control. Perform each test in parallel with appropriate controls as described above 

to ensure proper performance. Selected specimens may be tested in duplicate to ensure 

reproducibility. The description of the method is given as an example. When establishing 

the assay under given laboratory conditions, further adaptation, validation, and optimization 

could be required. 

A.4	Molecular	detection 

A.4.1	 Gel-based	PCR	for	detection	of	the	species	F. tularensis	and	genus	 

Francisella 

The multiplex PCR is based on detection of the 16S rRNA gene specific for strains of the genus 

Francisella and the lpnA encoding a 17-kDa lipoprotein (TUL4; amplicon size 0.4 kb) specific 

for the species F. tularensis. PCR is a rapid assay that can be used for both presumptive and 

confirmatory detection of F. tularensis. 

Application. The protocol has been developed primarily to detect F. tularensis in clinical samples obtained by taking swabs of ulcers (Johansson et al., 2000). DNA is recovered from the 

ulcer swabs and put in guanidine thiocyanate (GuSCN) buffer, designated L6. The swabs are 

transported in tubes containing 700 μl of the L6 buffer. 

Supplies and equipment for preparation of DNA 

1. Autoclaved 1.5 ml microcentrifuge tubes. 

2. Autoclaved 0.5 ml microcentrifuge tubes. 

3. Sterile pipette tips. 

4. Heating block. 

5. Sterile pipette tips. 

6. Pipette (10 μl–1000 μl, adjustable). 

7. Microcentrifuge. 

8. PCR equipment – thermocycler. 

9. Gel box. 

10. UV light box and camera. 

70
 
A. diAgnostic Protocols 

Reagents for nucleic acid extraction (by the method of Boom et al., 1990) 

1.	 L6-buffer, composed of: 

a) guanidine thiocyanate: 30 g 

b) 0.1M Tris pH 6.4: 25 ml 

c) 0.5 MEDTA pH8.0: 5.5ml 

d) Triton X-100: 650 mg 

Dissolve the guanidine thiocyanate in the Tris buffer by heating at 60 °C in a waterbath. 

Shake the bottle repeatedly. Add the EDTA and the Triton X-100. The L6 buffer can be 

dispensed into 700 μl aliquots in 1.5 ml microcentrifuge tubes and stored in the dark for 

3 months. 

2.	 L2-buffer, composed of: 

a) guanidine thiocyanate: 60 g 

b) 0.1 M Tris pH 6.4: 50 ml 

Dissolve the guanidine thiocyanate in the Tris buffer by heating at 60 °C in a waterbath. 

Shake the bottle repeatedly. The L2 buffer can be stored in the dark for 3 months. 

3.	 Ethanol (70 %). 

4.	 Acetone. 

5.	 Glassmilk® (MP Biomedicals, Irvine CA, USA). 

6.	 1x Tris EDTA (TE) buffer, pH 8.0. 

7.	 Distilled water. 

8.	 Agarose. 

9.	 1x Tris borate EDTA (TBE). 

10.	 Ethidium bromide. 

Reagents for PCR 

1.	 Distilled water. 

2.	 Ampli Taq Gold 5U/μl (Applied Biosystems, Foster City, California, USA). 

3.	 10x PCR-buffer II (Applied Biosystems). 

4.	 25 mM magnesium chloride (MgCl2

). 

5.	 10x dNTP stock solution (Amersham Biosciences, Buckinghamshire, UK). 

6.	 Stock solution of PCR primers (10 μM). 

Target	gene	 Primer	 Primer	sequences	(5’	to	’3) 

16S rRNA F5 cct ttt tga gtt tcg ctc c 

16S rRNA F11 tac cag ttg gaa acg act gt 

17-kDa lipoprotein TUL-435 gct gta tca tca ttt aat aac tgc tg 

17 kDa lipoprotein TUL-863 ttg gga agc ttg tat cat ggc act 

71
 
wHo guidelines on tulArAemiA 

PCR mixes
 

Master mix preparation:
 

1.	 Thaw master mix reagents. 

2.	 Spin down the contents of the tubes briefly. 

3.	 Mix the components of the master mix in 1.5 ml microcentrifuge tubes. 

4.	 Transfer 20 μl of the master mix to 0.5 ml microcentrifuge PCR tubes. 

5.	 Add 5μl of distilled water to the control tube. 

Master	mix	for	PCR	mixes	 μl 

10x dNTP 2.5 

25 mM MgCl2

 Solution 3.0 

10x PCR buffer II 2.5 

10 μMF5 1.5 

10 μM F11 1.5 

10 μM TUL-435 1.5 

10 μM TUL-863 1.5 

Distilled water 5.6 

Ampli Taq Gold 5U/μl 0.4 

Total	 20.0 

Further preparation pre­PCR 

1.	 Switch on heating block at 56 °C. 

2.	 Vortex the tube containing Glassmilk® until the suspension is dissolved. 

3.	 Check that there are no crystals in the L6 buffer; if there are, heat the tube at 45 °C until 

they have dissolved, then cool to room temperature. 

Positive control preparation 

1.	 Suspend the positive control strain in distilled H2

O to give an OD600 of 1.3 (approximately 

2 x 109 CFU/ml). 

2.	 Make a dilution of this suspension to give 2 x 105 CFU/ml in distilled H2

O. This will result 

in 150 CFU per PCR reaction. This is the positive control. 

3.	 Transfer 5μl of the positive control to 700 μl of L6 buffer and add 300 μl of distilled 

water. 

Negative control preparation. The negative control is composed of 700 μl of L6-buffer + 300 μl 

of distilled water. 

Sample preparation 

1.	 To each patient sample (700 μl), add 300 μl of L6. 

2.	 Add 5μl of glassmilk to each tube (samples and controls), vortex for 5 s. 

3.	 Leave for 10 min at room temperature. 

4.	 Vortex for 5 s. 

5.	 Centrifuge at 10 000 x g for 15 s. 

6.	 Discard the supernatant. 

7.	 Wash the deposit twice with 1 ml volumes of L2 buffer, twice with 1 ml volumes of ethanol and finally once with 1.0 ml of acetone. Between each step, vortex briefly and centrifuge at 10 000 x g for 15 s and discard the supernatant. 

72
 
 

A. diAgnostic Protocols 

8.	 Dry the pellets, with the lids of the tubes open, at 56 °C for 10 min. 

9.	 Add 30 μl of 1 x TE-buffer, vortex briefly. 

10.	 Incubate for 10 min at 56 °C. 

11.	 Vortex briefly. 

12. Centrifuge at 12 000 x g for 2 min. 

13.	 Transfer the supernatant to another 1.5 ml microcentrifuge tube. 

14.	 The tubes can be stored at 4 °C if the PCR is performed the same day, otherwise they 

should be stored at -18 °C overnight. 

15.	 Add 5μl of purified DNA from each sample to the tubes containing the PCR mixes (addition of DNA to PCR mixes should be performed in a designated pre-PCR area in a laminar flow hood). 

16.	 Vortex briefly and rapidly spin down. 

PCR programme 

1.	 95 °C for 10 min 

2.	 94 °C for 30 s 

3.	 60 °C for 1 min 

4.	 72 °C for 1 min 

5.	 perform 29 more cycles of steps 2–4 (30 cycles in total) 

6.	 hold at 4 °C. 

Electrophoresis. To analyse the PCR products, run 5μl on a 2% agarose gel with ethidium 

bromide. Include an appropriate molecular weight ladder on the gel. Use a UV light box to 

visualize the gel and take a picture to document the results. 

Interpretation. The amplicon size for TUL4 is 386 bp. No TUL4 product is amplified for F. philomiragia. The amplicon size for 16S rDNA is 1104 bp. 

Quality control. Positive (subspecies tularensis and subspecies holarctica) and negative (no template) controls should be included each time the PCR is performed. Dilutions of template 

DNA should be tested, as inhibition can be caused by high template DNA concentrations. 

A.5	Serology 

A.5.1	 Microagglutination 

Perspective A single serum/plasma specimen with a titre ≥1:128 is presumptive for F. tularensis. Paired serum specimens, taken 14 days apart, giving a 4-fold increase or decrease in titre 

with at least one titre being ≥1:128, are confirmatory for F. tularensis. A tube agglutination 

format with the same reagents also exists. The tube agglutination cut-off is ≥1:160. 

Application. Serum samples. 

Reagents 

1.	 Microagglutination (MA) diluent: phosphate-buffered saline with 1% normal rabbit 

serum, 0.4% formaldehyde; pH 7.2. 

2.	 Formalinized killed safranin-stained F. tularensis cells (CDC, Fort Collins, USA; Annex 

D). 

3.	 Positive control serum, high titre (> 1:128). 

73
 
wHo guidelines on tulArAemiA 

4. Positive control serum, low titre (1:128) (CDC, Fort Collins, USA; Annex D). 

5. Negative control serum, (< 1:16), matched by species to positive control serum. 

Equipment/supplies 

1. Microtitre plates, 96-well ”U” bottom (Costar, Cambridge, MA, USA). 

2. Sterile pipette tips. 

3. Reagent reservoir. 

4. Plate-sealing tape. 

5. Multichannel pipette (10 μl–100 μl, adjustable). 

6. Light box or mirror reader. 

7. Water-bath at 56 °C. 

Procedure 

1. Add 25 μl of MA buffer to each of the 96 wells of a microtitre plate. 

2. Add 25 μl of test serum to the first column of 8 wells (well 1): a total of 5 test samples and 

3 controls (high and low positive, negative) can be done on each plate. 

3. Carry out serial 2-fold dilutions of sera from well 1 to well 12, i.e. transfer 25 μl from well 1 

to well 2 and so on to well 12; discard the last 25 μl. 

4. Add 25 μl of antigen preparation to each well; mix gently. 

5. Seal plate to prevent evaporation, and incubate at room temperature for 12–18 h. 

6. Read and record the results. 

7. Discard microtitre plate in an appropriate container for decontamination and disposal. 

Interpretation. Negative specimens will have a button of cells at the bottom of the well. Positive specimens agglutinate the antigen, thus there will be a mat that looks like a diffused net 

of cells. There should be no cell button at the bottom of the well. Titres of ≥1:128 are considered positive. Prozone may be observed in cases of high titre serum when antibody is in 

excess of antigen. 

Quality control. Perform each test in parallel with high-, low-positive and negative control 

sera to ensure proper performance. Selected specimens may be tested in duplicate to ensure 

reproducibility. If Brucella infection is suspected, microagglutination should be performed in 

parallel with Brucella antigen to exclude cross-reactions. 

A.5.2	 Enzyme-linked	immunosorbent	assay	(ELISA) 

Perspective. This procedure determines the titres of anti-Francisella immunoglobulins, isotype 

G and M. A single serum/plasma specimen with a positive ELISA titre is presumptive for F. 

tularensis. Paired serum specimens, with at least one having a significant IgG or IgM titre 

is considered confirmatory for F. tularensis. In part, this procedure might be automated i.e. 

washing the plates. 

Application. Human serum samples. 

Consumables 

1. 96-well microtitre plates, flat-bottomed (Nunc, Immunoplate MaxiSorp). 

2. 15 ml conical tubes (Falcon). 

3. Plate seal/adhesive cover (Nunc, can be replaced by parafilm). 

4. Pipette tips (Sarstedt). 

74
 
 

 

 

 

 

 

 

 

A. diAgnostic Protocols 

Solutions (see recipes in section A.5.3) 

1. Coating buffer. 

2. Francisella tularensis LPS. 

3. Washing buffer. 

4. Incubation buffer. 

5. Substrate buffer. 

6. Substrate tablets (Sigma 104 phosphatase substrate tablets number S0942). 

7. 5M and 3M NaOH. 

8. Standard and positive control (titrated patient sera, possibly pooled (see Titration of standards and controls, section A.5.3). 

9. Anti-IgM/anti-IgG. 

Equipment 

1. ELISA reader (Tecan Sunrise). 

2. Multichannel pipette. 

3. Pipette charger. 

4. Single channel pipette. 

5. If applicable: automated plate washer (Anthos fluido). 

Procedure. All buffers and solutions should be at room temperature. 

DAY 1 

1. Coating: 

a) Take out plates from fridge. Coat the plates according to the coating scheme below; note 

lot numbers of reagents, coating date and assign consecutive numbers to the plates. 

b) Dilute the antigen in coating buffer to a concentration of 3μg LPS/ml. Allow at least 4 

ml per plate.
 

c) Load 100 μl per well according to the coating scheme below.
 

d) Cover plate with plate seal and incubate overnight at 20–26 °C.
 

Coating	scheme	(CB	=	coating	buffer,	LPS	=	F. tularensis	LPS 

	 1	 2	 3	 4	 5	 6	 7	 8	 9	 10	 11	 12 

A CB CB CB CB CB CB CB CB CB CB CB CB 

B CB LPS LPS LPS LPS LPS LPS LPS LPS LPS LPS CB 

C CB LPS LPS LPS LPS LPS LPS LPS LPS LPS LPS CB 

D CB CB CB CB CB CB CB CB CB CB CB CB 

E CB CB CB CB CB CB CB CB CB CB CB CB 

F CB LPS LPS LPS LPS LPS LPS LPS LPS LPS LPS CB 

G CB LPS LPS LPS LPS LPS LPS LPS LPS LPS LPS CB 

H CB CB CB CB CB CB CB CB CB CB CB CB 

75
 
 

 

 

 

 

 

 

 

 

wHo guidelines on tulArAemiA 

DAY 2 

2. Blocking: 

a) Empty the wells, blot the inverted plate on absorbent paper to remove any residual 

buffer. Load 200 μl washing buffer per well in rows B-G, columns 2–11. 

b) Cover plate with plate seal. Incubate plates at 20–26 °C for at least 30 min. After this 

step the plates can be stored at either 4 °C or -70 °C. 

3. Preparations: 

a) Design loading scheme for the plates in form of a computer protocol, Word or Excel file 

format. The outermost wells are left empty. Include standards on all plates (column 2 

and 3) but only include a positive control on the first plate (column 3). 

b) Take out patient sera and standard/positive control and allow to thaw. Do not inactivate.
 

c) Note the lot numbers of reagents, solutions and controls.
 

d) Label two test tubes per patient serum: 1/100 and 1/1000.
 

4. Rinse: 

a) Empty the wells, blot the inverted plate on absorbent paper to remove any residual 

buffer. Fill the wells with washing buffer (c. 300 μl/well) and wait one minute. Repeat 

five times. 

b) Place the plates upside-down on moist tissue; they should not dry out! 

5. Dilute samples: 

a) Dilute the serum samples 1/1000, in two steps: 

i. 25 μl serum + 2.5 ml incubation buffer, mix carefully (1) 

ii. 400 μl of (1) + 3.6 ml incubation buffer, mix. 

b) Dilute standard/control according to titration (see Titration of standards and controls, 

section A.5.3). 

6. Incubate samples: 

a) Load 100 μl per well according to the loading scheme below. Place standard/control (C) 

in column 2, 3 and 4. Place each sample (the 1/1000 dilution) into four antigen-coated 

wells and into two wells with coating buffer. The latter serve as control for unspecific 

reactions. 

Loading	scheme	(IB	=	incubation	buffer) 

	 1	 2	 3	 4	 5	 6	 7	 8	 9	 10	 11	 12 

A IB IB IB IB IB IB IB IB IB IB IB IB 

B IB C C C 01 02 03 04 05 06 07 IB 

C IB C C C 01 02 03 04 05 06 07 IB 

D IB C C C 01 02 03 04 05 06 07 IB 

E IB C C C 01 02 03 04 05 06 07 IB 

F IB C C C 01 02 03 04 05 06 07 IB 

G IB C C C 01 02 03 04 05 06 07 IB 

A IB IB IB IB IB IB IB IB IB IB IB IB 

b) Cover plate with plate seal and incubate for 3–5 hours at 20–26 °C. 

76
 
 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

A. diAgnostic Protocols 

7.	 Rinse: 

a) Repeat rinse steps as above. 

8.	 Incubate conjugate: 

a) Every sample is tested with two conjugates, anti-IgG and anti-IgM. Allow at least 3 ml 

of each conjugate per plate. 

b) Immediately before use dilute the conjugates in incubation buffer (according to lotspecific instructions from the manufacturer). 

c)	 Load 100 μl per well according to the loading scheme below; anti-IgG in rows B-D; 

anti-IgM in rows E-G. 

Conjugate	loading	scheme	(IB	=	incubation	buffer) 

	 1	 2	 3	 4	 5	 6	 7	 8	 9	 10	 11	 12 

A IB IB IB IB IB IB IB IB IB IB IB IB 

B IB antiG antiG antiG antiG antiG antiG antiG antiG antiG antiG IB 

C IB antiG antiG antiG antiG antiG antiG antiG antiG antiG antiG IB 

D IB antiG antiG antiG antiG antiG antiG antiG antiG antiG antiG IB 

E IB antiM antiM antiM antiM antiM antiM antiM antiM antiM antiM IB 

F IB antiM antiM antiM antiM antiM antiM antiM antiM antiM antiM IB 

G IB antiM antiM antiM antiM antiM antiM antiM antiM antiM antiM IB 

H IB IB IB IB IB IB IB IB IB IB IB IB 

d) Cover plates with plate seal and incubate overnight at 20–26 °C. 

DAY 3 

9.	 Repeat rinse steps as above. 

10.	 Prepare substrate: 

a) Prepare at least 6 ml substrate dilution per plate. Dissolve one substrate tablet in 5 ml 

substrate buffer. Incubate for 30 min at room temperature. 

b) For blanks mix 50 μl 5M NaOH with 5 ml substrate buffer. Allow 800 μl per plate. 

11. Incubate substrate: 

a) Load 100 μl per well according to the loading scheme below. 

Loading	scheme	 

(B	=	blank,	SB	=	substrate	buffer,	SS	=	substrate	solution) 

	 1	 2	 3	 4	 5	 6	 7	 8	 9	 10	 11	 12 

A B SB SB SB SB SB SB SB SB SB SB SB 

B B SS SS SS SS SS SS SS SS SS SS SB 

C B SS SS SS SS SS SS SS SS SS SS SB 

D B SS SS SS SS SS SS SS SS SS SS SB 

E B SS SS SS SS SS SS SS SS SS SS SB 

F B SS SS SS SS SS SS SS SS SS SS SB 

G B SS SS SS SS SS SS SS SS SS SS SB 

H B SB SB SB SB SB SB SB SB SB SB SB 

b) Once you have finished loading the first plate start the timer to record the incubation 

time. 

77 
wHo guidelines on tulArAemiA 

12. Read and stop the reaction: 

a) Read absorbance at 405 nm. Stop the reaction in the first plate when the standards/ 

calibrator (columns 2 and 3) have an absorbance of approx. 1. 

b) Stop the reaction by adding 50 μl 3M NaOH. 

c) Read absorbance at 405 nm. Save the results as an Excel file connected to your loading 

scheme. Use this reading for calculations. 

13. Calculation: 

a) This calculation can be done automatically by setting up an Excel file. Single wells are 

identified by the combination of a letter from A-H and a number from 1–12, according 

to the rows and columns of the plate. 

b) Calculate the titre of anti-F. tularensis LPS IgG: 

i.	 mean of the controls: 

(B2 + C2)/2 = mean1
 

(B3 + C3)/2 = mean2
 

ii.	 background correction: 

mean1

 – D2 = background corrected mean 1 (bcm) 

mean2 – D3 = bcm2 

calibrator = (bcm1 +bcm2)/2 

correction factor = 1/calibrator 

Due to previous titration of the standards, the expected absorbance (A405) in wells B2, 

C2, B3 and C3 is OD 1. This calculation is done to adjust the expected value to the real 

value of this reading. 

c)	 Calculate the IgG titre in patient sera. Repeat the following steps for each sample. For 

example: 

mean = (B5 + C5)/2 

background correction: mean - D5 = background corrected value 

relative IgG titre = background corrected value x correction factor x dilution of the 

serum. 

d) Calculate the anti-F. tularensis LPS IgM titres in patient sera. Repeat the previous steps 

as described for IgG titres but use absorbance values for wells F2, G2, F3 and G3 for 

the correction factor, row E for background correction and sample absorbance values 

in rows F and G for the relative IgM titre. 

Interpretation. Negative blood samples from humans who had no record of tularaemia or vaccination against tularaemia typically result in IgG and IgM titres of 0.2–0.4. 

Quality control. Perform each test in parallel with positive and negative control sera to ensure 

proper performance. Document all test results. 

A.5.3	 Buffer	recipes	and	storage	conditions	 

Coating buffer 

Ingredients 

a) Sodium hydrogen carbonate, NaHCO3

: 3.47 g.
 

b) Sodium carbonate, NaCO3

: 1.18 g.
 

c) Sodium azide, NaN3

: 0.20 g.
 

d) Ultra-pure water to total volume: 1000 ml.
 

78
 
A. diAgnostic Protocols 

Preparation 

a) Dissolve the substances in water to approximately three-quarters of the total volume.
 

b) Adjust the pH to 9.5–9.7.
 

c) Dilute with water to 1000 ml.
 

d) Autoclave at 121 °C for 15 min.
 

Storage. 2–8 °C.
 

Best before. 3 months.
 

Environmental and safety instructions
 

a) Sodium azide is a highly toxic substance that can be absorbed through the skin. If 

sodium azide gets in contact with acids a toxic gas develops. If sodium azide gets in 

contact with lead or copper explosive metal azides develop. 

b) Work in a fume hood and wear gloves when handling sodium azide. Excess sodium 

azide solutions must not be poured down the sink but should be disposed of in accordance with national regulations. 

Washing buffer (PBS with 0.5% BSA) 

Ingredients 

a) Bovine serum albumin (BSA): 5.00 g.
 

b) Sodium azide, NaN3

: 0.20 g.
 

c) PBS: 1000 ml.
 

Preparation 

a) Dissolve the substances in PBS.
 

b) Adjust the pH to 7.2–7.4.
 

c) Filter the solution through a 0.2 μm membrane filter.
 

Storage. Room temperature, dark. 


Best before. 2 months.
 

Environmental and safety instructions. As for sodium azide (see coating buffer above).
 

Incubation buffer (PBS with 0.05% Tween 20) 

Ingredients 

a) Sodium chloride, NaCl: 8.00 g.
 

b) Disodiumhydrogenphosphate Na2

HPO4 x 2H2O:1.43g.
 

c) Potassium chloride, KCl: 0.20g.
 

d) Potassium dihydrogenphosphate KH2

PO4 x 2H2O2: 0.20g.
 

e) Sodium azide, NaN3

: 0.20 g.
 

f) Tween-20: 0.5 ml.
 

g) Ultra-pure water to a total volume: 1000 ml.
 

Preparation 

a) Dissolve the substances in water to approximately three-quarters of the total volume. 

b) Adjust the pH to 7.3–7.4. 

79
 
wHo guidelines on tulArAemiA 

c)	 Dilute with water to 1000 ml. 

d) Autoclave at 105 °C for 15 min. 

Storage: Room temperature, dark. 

Best before: 1 month. 

Environmental and safety instructions: As for sodium azide (see above; coating buffer). 

Substrate buffer 

Ingredients 

a) Diethanolamine: 97 ml.
 

b) Magnesium chloride MgCl2 x 6H2O2: 0.101 g.
 

c) Sodium azide, NaN3: 0.20 g.
 

d) Ultra-pure water to a total volume: 1000 ml.
 

Preparation 

a) Dissolve the substances in water to approximately three-quarters of the total volume.
 

b) Adjust the pH to 9.8 with HCl.
 

c) Add water to give a total volume of 1000 ml.
 

Storage: Room temperature, dark. 

Best before: 2 months. 

Environmental and safety instructions 

a) For sodium azide, see above (coating buffer).
 

b) Diethanolamine can be harmful to the eyes.
 

Titration of standards and controls 

a) Test patient serum or alternatively serum from vaccinees for anti-F. tularensis IgG and 

IgM according to the ELISA protocol. 

b) Select a serum that is positive for both immunoglobulin isotypes and gives absorbance 

of > 1 at 450 nm. 

c)	 Perform serial two-fold dilutions of the selected serum and perform the ELISA protocol 

again. 

d) An appropriate standard/control should reach an absorbance of 1 within 20–40 min. 

The dilution of the selected serum which fulfils this criterion is used as standard in 

future experiments. 

A.6	References 

Boom et al (1990). Reagents for nucleic acid extraction by the Boom method. Journal of Clinical 

Microbiology, 28:495–503. 

Grunow et al (2000). Detection of Francisella tularensis in biological specimens using a capture 

enzyme-linked immunosorbent assay, an immunochromatographic handheld assay, and a 

PCR. Clinical and Diagnostic Laboratory Immunology, 7:86–90. 

Johansson et al. (2000). Comparative analysis of PCR versus culture for diagnosis of ulceroglandular tularaemia. European Journal of Clinical Microbiology, 38:22–26. 

80
 
B. tests For suPlementAl cHArActerizAtion

B.	 Tests	for	supplemental	characterization 

The following protocols have been provided by the University of Umeå, Umeå, Sweden and 

the Centers for Disease Control and Prevention, Fort Collins, CO, USA. 

Once an isolate has been confirmed as F. tularensis, additional methods can be used for supplemental characterization. The following protocols should be considered only supplementary 

and not diagnostic for F. tularensis. 

B.1	 Antimicrobial	susceptibility 

B.1.1	 E-test 

Perspective. The E-test (AB Biodisk, Solna, Sweden) method is based on the use of plastic 

strips containing a predefined antibiotic gradient. The primary reason for performing this 

supplementary test is to determine the antibiotic susceptibility of the recovered isolate. 

Application. Isolates confirmed as F. tularensis. 

Procedure. Consult the E-test® Technical Manual (AB Biodisk) for more detailed information. 

Agar medium 

1. Cysteine heart agar + 10% blood (Tomaso et al., 2005) or + 2% haemoglobin (Ikäheimo et 

al., 2000). 

2. Glucose cysteine blood agar (Scheel et al.,1993). 

3. GCII agar with 1% haemoglobin and 1% IsoVitaleXTM (Johansson et al., 2000) (See section 

C.2 ). 

Inoculum 

Suspend 48-h colonies from chocolate agar in broth to give an opacity equivalent to 1 McFarland standard. 

Incubation 

37 °C in 5% CO2

 or ambient atmosphere for 48 h. 

Caution 

1. Testing should only be performed if appropriate biosafety measures are available (see section 6.6). 

2. Public health officials should be notified regarding all isolates presumptively identified as 

F. tularensis. 

B.1.2	 Quality	control 

Quality control (MIC mg/l). The values given for the reference strains below are based on the 

CLSI broth microdilution method using supplemented cation-adjusted Mueller-Hinton broth 

(CAMHB) and 48 h incubation at 37 °C (Clinical and Laboratory Standards Institute, 2005). 

81
 
 

 

 

wHo guidelines on tulArAemiA 

	 Staphylococcus aureus Pseudomonas aeruginosa	 

	 Atcc 29213 Atcc 27853 

Ciprofloxacin 0.25–1 0.25–1 

Chloramphenicol 4–32 not tested 

Doxycycline 0.25–2 4–32 

Gentamicin 0.25–1 0.5–4 

Levofloxacin 0.12–0.5 0.5–4 

Streptomycin 8–64 32–256 

Tetracycline 0.5–4 8–64 

Interpretation: CLSI MIC criteria (mg/l) for F. tularensis. Values are based upon broth microdilution using supplemented CAMHB and 48 h incubation at 37 °C (Clinical and Laboratory 

Standards Institute, 2005). 

	 	 MIC	criteria	(mg/l) 

Antibiotic	 Susceptible	 Intermediatea	 Resistant 

Ciprofloxacin 

Chloramphenicol 

Doxycycline 

Gentamicin 

≤ 0.5

≤ 8.0

≤ 4.0

≤ 4.0

 –

 –

 –

 –

 – 

– 

– 

– 

Levofloxacin ≤ 0.5 – – 

Streptomycin 

Tetracycline 

≤ 8.0

≤ 4.0

 –

 –

 – 

– 

a The absence of resistant strains precludes defining any result categories other than ’susceptible’. For strains 

yielding results suggestive of a ’nonsusceptible’ category, organism identification and antimicrobial susceptibility 

test results should be confirmed. 

Precautions when reading results 

1. For bactericidal drugs, read at complete inhibition of growth (microcolonies, hazes and 

isolated colonies). 

2. For bacteriostatic drugs, read at 80% inhibition when trailing is seen. 

3. Please consult the E-test Technical Manual for more information, downloadable at www. 

abbiodisk.com. 

B.2	Molecular	biotyping 

The methods below allow the discrimination of different subspecies of F. tularensis. 

B.2.1	 Gel-based	PCR	for	identification	of F. tularensis	subspecies	holarctica		 

(type	B)	strains	(Byström	et	al.,	2005) 

The targeted F. tularensis DNA marker has been denoted (Ft-M19) and is present in all four 

F. tularensis subspecies: tularensis (type A), holarctica (type B), mediasiatica, and novicida. The 

genomes of subspecies holarctica strains universally exhibit a specific 30-bp deletion at the 

targeted genomic locus (Johansson et al., 2004; Byström et al., 2005). The PCR assay relies 

on amplification of Ft-M19 that is located between the genes deaD and ppiC using a pair of 

amplification primers that are located inside the flanking genes. 

Application. The protocol has been developed for rapid subtyping of F. tularensis cultures. The 

Ft-M19 assay can be used after identification to the species level using diagnostic methods 

outlined in Annex A. 

Supplies and equipment. See protocol A.4.1 for listing of supplies and equipment needed. 

82
 
 

B. tests For suPPlementAl cHArActerizAtion 

Preparation of DNA 

1.	 Grow bacteria for 24–48 h on an appropriate solid agar medium at 37 °C. 

2.	 Collect 5–10 bacterial colonies on a sterile bacteriological loop and prepare a suspension 

of bacteria in 1 ml of water. 

3.	 Kill the bacteria by incubating the suspension at 65 °C for 2 h. 

4.	 Transfer 100 μl of the heat-killed bacterial suspension to 900 μl of buffer L6 (see Reagents 

for nucleic acid extraction, section A.4.1). 

5.	 Add 5 μl of Glassmilk® to each tube with samples or controls, vortex for 5 s. 

6.	 Leave for 10 min in room temperature then vortex 5 s. 

7.	 Centrifuge at 10 000 x g for 15 s and then discard the supernatant. 

8.	 Wash the deposit twice with 1 ml of L2 buffer (see Reagents for nucleic acid extraction, section A.4.1), twice with 1 ml of ethanol and finally once with 1 ml of acetone. Between each 

step, vortex briefly, centrifuge at 10 000 x g for 15 s and discard the supernatant. 

9.	 Dry the pellets with the lids of the tubes open, at 56 °C for 10 min. 

10.	 Add 30 μl of 1x TE-buffer, vortex briefly. 

11.	 Incubate for 10 min at 56 °C, vortex briefly then centrifuge at 12 000 x g for 2 min. 

12.	 Transfer the supernatant to another 1.5 ml micro centrifuge tube. The tubes can be stored 

at 4 °C if the PCR is performed the same day, otherwise they should be stored at -18 °C 

overnight. 

B.2.2	 PCR 

1.	 For PCR reagents, see Reagents for PCR, section A.4.1. 

2.	 PCR primers: 

Forward: 5’-aggcggagatctaggaaccttt-3’ 

Reverse: 5’-agcccaagctgactaaaatcttt-3’ 

PCR mixes. For PCR mix set-up and positive and negative control preparation, see section 

A.4.1. 

Master mix preparation 

Master	mix	for	PCR	 μl 

dNTP (2.5 mM) 2.0 

MgCl2

 solution (25 mM) 1.2 

10xPCR buffer II 2.5 

Forward primer (5 pmol/μl) 2.0 

Reverse primer (5 pmol/μl) 2.0 

Distilled water 14.1 

Ampli Taq Gold 5 U/μ 24.0 

Total	 25.0 

83
 
 

wHo guidelines on tulArAemiA 

Sample preparation 

1. Add 1 μl of purified DNA sample to each PCR mix to a final volume of 25 μl. This should 

be performed in a designated pre-PCR area in a laminar flow hood. 

2. Vortex briefly and spin down the contents of the tubes without delay. 

3. Immediately place the PCR tubes in the cycling equipment and start the PCR. 

PCR programme 

1. 95 °C for 10 min 

2. 94 °C for 30 s 

3. 62 °C for 30 s 

4. 72 °C for 30 s 

5. repeat steps 2–4 for 29 cycles 

6. hold at 4 °C. 

Electrophoresis 

1. To analyse the PCR products, load 3 μl sample mixed with loading dye on a 3 % NuSieve 

3:1 agarose gel (Cambrex Bioscience Rockland, Inc., Maine) with ethidium bromide. 

2. Include an appropriate molecular weight ladder on the gel and run in 1x TBE buffer at 5 

V/cm for 3 h. 

3. Use a UV light box to visualize the DNA and photograph to document the results. 

Interpretation. The PCR fragment size of Ft-M19 is 220 bp for all strains of subspecies holarctica 

(type B). PCR amplifications of DNA from strains of subspecies tularensis (type A), mediasiatica, and novicida each result in a 250 bp product. 

Quality control. Positive (subspecies tularensis and subspecies holarctica) and negative (no template) controls should be included each time the PCR is performed. Dilutions of template 

DNA should be tested, as inhibition can be caused by high-template DNA concentrations. 

B.2.3	 Subspecies-specific	real-time	PCR	assays	for	subspecies	tularensis	and	 

subspecies	holarctica	(Kugeler	et	al.,	2006) 

Perspective. The subspecies tularensis (type A) specific assay is targeted against the pdpD gene 

and the subspecies holarctica (type B) specific assay detects a unique location of the ISFtu2 

insertion-like element. The type A and type B assays are TaqMan® assays utilizing two primers and a fluorogenic probe for target detection. These assays were developed and evaluated 

on the LightCycler® 1.2 (Roche Applied Science, Indianapolis, IN) real-time PCR instrument. 

Application. The protocol has been developed for rapid subtyping of bacterial F. tularensis cultures. These assays can be used after identification to the species level using diagnostic methods outlined in Annex A. 

Note. Due to the sensitive nature of real-time PCR, care should be taken to ensure that false 

positive results do not occur as a result of cross-contamination. 

Supplies and equipment 

1. Lightcycler capillaries. 

2. Lightcycler carousel. 

3. Lightcycler centrifuge adapters. 

4. Lightcycler carousel centrifuge (optional). 

84
 
 

B. tests For suPPlementAl cHArActerizAtion 

5. Sterile microcentrifuge tubes (1.7 ml). 

6. Sterile PCR tubes (0.5 ml). 

7.	 Pipettors. 

8. Pipette tips (aerosol barrier). 

9. Microcentrifuge. 

Reagents 

1. Qiagen QIAamp® DNA Mini Kit (Valencia, CA). 

2. LightCycler® Fast Start DNA Master Hybridization Probes (Roche). 

Components: 

a) PCR grade water (#3)
 

b) MgCl2

 (#2)
 

c) 10x reaction mix (#1)
 

3. Uracil DNA glycosylase (2U/μl) (Roche). 

4. TaqMan® primers and probes 

Type A assay: 

a) Forward primer (stock = 50 μM) (final= 750 nM per reaction) 


5’ gagacatcaattaaaagaagcaatacctt 3’
 

b) Reverse primer (stock = 50 μM) (final= 750 nM per reaction)
 

5’ ccaagagtactatttccggttggt 3’
 

c)	 Probe (stock = 10 μM) (final= 200 nM per reaction) 


5’ aaaattctgc*t*cagcaggattttgatttggtt 3’
 

Type B assay: 

a) Forward primer (stock = 50 μM) (final= 750 nM per reaction)
 

5’ cttgtacttttatttggctactgagaaact 3’
 

b) Reverse primer (stock = 50 μM) (final= 750 nM per reaction)
 

5’ cttgcttggtttgtaaatatagtggaa 3’
 

c)	 Probe (stock = 10 μM) (final= 200 nM per reaction)
 

5’ acctagttcaacc*t*caagacttttagtaatgggaatgtca 3’
 

Probes are synthesized with 5’ FAM (6-carboxyl-fluorescein) and internal dark quenchers 

(either BHQ1 or QSY-7) at the nucleotide positions indicated by the *. 

DNA template preparation 

1. Grow bacteria for 24–48 h at 37 °C on an appropriate agar medium. 

2. Collect 1 μl of cells using a sterile bacteriological loop and add to 180 μl of Buffer ATL 

(QIAamp DNA Mini Kit). 

3. Purify DNA according to the QIAamp® tissue protocol. 

4. As a final step, elute DNA from the QIAamp® column in 200 μl of Qiagen elution buffer. 

Negative control preparation 

1. The negative control should be prepared using sterile water distributed in aliquots into 

sterile tubes in a clean DNA-free area. 

2. Aliquots may be prepared and stored at -20 °C until use. 

85
 
wHo guidelines on tulArAemiA 

Positive control preparation 

1. Both a type A and a type B strain are needed for positive controls. 

2. The control strains should be diluted to approximately 1000–10 000 genomic equivalents 

per μl (between 2–20 pg/μl) to minimize risk for cross-contamination. 

3. Aliquots may be prepared and stored at -20 °C until use. 

Procedure. In a DNA-free area (clean room with hood, with dedicated gloves, pipettors, tips, 

lab coat, etc.). 

1. Calculate how many reactions are going to be performed for each assay (type A and type B) 

including an appropriate number of positive and negative controls. In calculating volumes 

for master-mix preparation, add extra reactions (10%) for pipetting errors. For example, if 

you have 10 samples, 1 positive and 1 negative control (12 capillaries total) to run with the 

type A assay, base the master-mix calculation on 15 capillaries. 

Reagent	 Volume	per	reaction	(μl) 

Water (#3) 3.3 

MgCl2 (#2) 3.2 

10X reaction mix (#1) 2.0 

Uracil DNA glycosylase (2 U/μl) 0.5 

Forward primer (50 μM) 0.3 

Reverse Primer (50 μM) 0.3 

Probe (10 μM) 0.4 

Total	 10.0 

2. Prepare the master mix for each assay separately. Prepare one sterile tube with the master 

mix for the type A assay, and one tube with the master mix for the type B assay. 

3. Move master mixes to a separate area for loading and addition of DNA. 

4. Working in a separate area or hood for DNA work (with dedicated gloves, pipettors, tips, 

lab coat, etc.): 

a) Add capillaries either directly to a LightCycler® carousel or LightCycler® centrifuge 

adapters with clean gloves, taking extra care to not touch the opening of the capillaries. 

b) The final reaction volume for each capillary is 20 μl. 

c)	 Add 10 μl of master mix to each capillary. 

d) Prepare negative control capillaries first. Add 10 μl of sterile water to the negative control capillary and cap. 

e) Prepare sample capillaries next. Add 9 μl of sterile water and 1 μl of samples to be tested 

and then cap. If necessary, more sample DNA can be added (up to 9 μl) and water volume adjusted accordingly for the final reaction volume of 20 μl. 

f)	 Prepare positive control capillaries last. Add 9 μl of sterile water and 1 μl of DNA and 

then cap. 

g) Gently centrifuge (either in carousel with LightCycler® carousel centrifuge or in an allpurpose microcentrifuge using LightCycler® centrifuge adapters) to collect contents at 

the bottom of the capillaries. 

h) Load carousel into LightCycler® and run programme. 

86
 
B. tests For suPPlementAl cHArActerizAtion 

PCR programme 

1. 50 °C for 2 min 

2. 95 °C for 10 min 

3. 95 °C for 10 s 

4. 65 °C for 30 s 

5. repeat steps 3–4 for 44 cycles 

6. 45 °C for 5 min. 

Results and interpretation. After the programme has finished, proceed to analysis by adjusting 

the y-axis to channel F1/F3. Baseline adjustment should be selected as arithmetic. The 

threshold-crossing values are then automatically calculated using the second derivative 

maximum method. A sample is considered positive if it crosses the threshold during the 45 

cycles of the programme AND displays logarithmic amplification. 

Quality control. Any PCR run should be repeated if the negative and positive controls do not 

perform as intended. PCR inhibition can be caused by numerous factors. Negative samples 

can be spiked to check for PCR inhibition. It is important to ensure weak positive results (high 

Ct values) are not due to cross-contamination from positive control samples. 

B.3	References 

AB Biodisk, Solna, Sweden: Package insert, product labels, E-test Technical Guide, Media/ 

Inoculum/Incubation, Reading and Trouble Shooting charts, CIS 007; also see www. 

abbiodisk.com. 

Byström M et al. (2005). Tularaemia in Denmark: identification of a Francisella tularensis subsp. holarctica strain by real-time PCR and high-resolution typing by multiple-locus variablenumber tandem repeat analysis. Journal of Clinical Microbiology, 543:5355–5358. 

Clinical and Laboratory Standards Institute (2005). M100-S15, Performance Standards for 

Antimicrobial Susceptibility Testing; Fifteenth Informational Supplement. Table 2K:136– 

137,144. 

Ikäheimo I et al. (2000). In vitro antibiotic susceptibility of Francisella tularensis isolated from 

humans and animals. Journal of Antimicrobial Chemotherapy, 46:287–290. 

Johansson A et al. (2000). Ciprofloxacin for treatment of tularaemia in children. Pediatric Infectious Disease Journal, 19:449–453. 

Johansson et al. (2004). Worldwide genetic relationships among Francisella tularensis isolates 

determined by multiple-locus variable-number tandem repeat analysis. Journal of Bacteriology, 

186:5808–5818. 

Kugeler KJ et al. (2006). Real-time subspecies-specific PCR for Francisella tularensis type A 

and type B. Emerging Infectious Diseases, 12:1799–1801. 

Scheel O et al. (1993). Susceptibility pattern of Scandinavian Francisella tularensis isolates 

with regard to oral and parenteral antimicrobial agents. Acta Pathologica, Microbiologica, et 

Immunologica Scandinavica, 101:33–36. 

Tomaso et al. (2005). Antimicrobial susceptibilities of Austrian Francisella tularensis holarctica 

biovar II strains. International Journal of Antimicrobial Agents, 26:279–284. 

87
 

c. Protocols For PrePArAtion oF selected F. tulArensis culture mediA

C.	 Protocols	for	preparation	of	selected		 

	 F. tularensis	culture	media 

The following protocols have been provided by the University of Umeå, Sweden and the Centers for Disease Control and Prevention, Fort Collins, CO, USA. 

C.1	Cysteine	heart	agar	base	with	9%	chocolatized	sheep	blood 

Cystine heart agar (CHA) with blood (CHAB) is a nonselective medium used for primary isolation and cultivation of F. tularensis. Additionally, growth on CHAB provides a presumptive 

identification of F. tularensis as the organism shows characteristic growth on this medium 

(green, opalescent, raised, shiny colonies at 24–48 h). The recipe can be supplemented with 

antibiotics for the isolation of F. tularensis from materials that may be contaminated with other 

organisms. 

C.1.1	 Materials	(for	1	litre	of	medium) 

1. Cysteine heart agar powder (Becton Dickinson Diagnostic Systems). 

2. Defibrinated sheep blood. 

3. For preparation of antibiotic supplemented CHA (CHAB-A) (per litre of medium): 

a) 7.5 mg colistin 

b) 2.5 mg amphotericin 

c) 0.5 mg lincomycin 

d) 4 mg trimethoprim 

e) 10 mg ampicillin. 

C.1.2	 Procedure 

1. Suspend 51 g of CHA powder in 1 litre of purified water. 

2. Mix thoroughly, heat with frequent agitation, bring to a boil, and gently swirl to completely 

suspend the powder. 

3. Autoclave at 121 °C for 15 min. 

4. Cool to 72 °C. 

5. Aseptically add 90 ml of defibrinated sheep blood (9% final) and mix on heated stirrer until 

blood becomes chocolatized. 

6. If preparing CHAB-A, cool to 55 °C and add antibiotics. 

7.	 Mix gently but carefully using a magnetic stirrer and avoid the formation of air bubbles (i.e. 

foam) in the agar. 

8. Dispense 20 ml volumes of medium into sterile Petri dishes. 

9.	 Replace the lids on the Petri dishes, and allow the medium to stay at room temperature for 

several hours. Place plates in a plastic bag and store at 4 °C. 

89
 
 

 

 

 

 

 

 

wHo guidelines on tulArAemiA 

C.1.3	 Quality	control 

1. Incubate several uninoculated plates at 37 °C for 24 h and check for contamination. 

2. Inoculated CHAB should support the growth of F. tularensis (e.g. LVS) after incubation at 

37 °C in ambient air or in a 5% CO2

 humid atmosphere for 48 h. 

3. F. tularensis should appear as white to greenish colonies, usually too small to be seen as 

individual colonies at 24 h. After incubation for 48 h, colonies are about 2–4 mm in diameter, opaque and exhibit greenish-lavender colour. An opalescent sheen is apparent on the 

surface of the colonies if incubated 48–72 h. 

C.2	GCII	agar	with	1%	haemoglobin	and	1%	IsoVitaleX 

GCII agar with haemoglobin and IsoVitaleX is a nonselective medium used for primary isolation 

and cultivation of fastidious microorganisms; especially Francisella, Neisseria and Haemophilus 

species. To obtain a suitable level of standardization for qualitative microbiological procedures, 

haemoglobin powder has replaced whole blood products in this ’chocolate agar’ medium. GCII 

agar with haemoglobin was originally developed for standardized culture of Neisseria gonorrhoeae and can be used with various supplements that enhance the growth of fastidious bacteria 

(e.g. IsoVitaleX) and/or to suppress contaminant bacteria (e. g. various antimicrobial agents). 

The following methods allow for the production of 200 ml of medium (five plates, 90 mm 

diameter); adjust quantities proportionately for the production of larger volumes of medium. 

C.2.1	 Haemoglobin	solution 

1. Add 2 g of soluble bovine haemoglobin powder (Becton Dickinson Diagnostic Systems, 

Sparks, MD, USA) to 100 ml of ultrapure water and mix using a magnetic stirrer. Store 

overnight at 4 °C. 

2. Mix thoroughly the following day using the magnetic stirrer until a smooth suspension is 

achieved. 

3. Autoclave the haemoglobin solution at 121 °C for 15 min. Cool in a water-bath set to 50– 

60 °C. 

C.2.2	 GCII	agar 

1. Suspend 7.2 g of GCII agar base (Becton Dickinson Diagnostic Systems) in a small amount 

of ultrapure water and add water to a final volume of 100 ml. 

2. Mix thoroughly, heat with frequent agitation, bring to a boil, and gently swirl to completely 

suspend the powder (approximately 1 min). 

3. Autoclave the medium at 121 °C for 15 min. Cool in a water-bath set to 52 °C. 

C.2.3	 Enrichment	supplement	(IsoVitaleX) 

IsoVitaleX, a registered trademark of Becton Dickinson, is a chemically defined mixture of 

supplements that enhances the growth of fastidious bacteria. IsoVitaleXTM (Becton Dickinson) 

comes in vials of lyophilized substance. 

1. Aseptically open the vial containing the lyophilized growth supplement. 

2. Use a sterile needle and syringe to aseptically transfer 10 ml of the accompanying diluent 

to the vial. 

3. Shake to ensure complete solution. After reconstitution use the growth supplement immediately, or store at 4 °C and use within 2 weeks. 

4. The composition of an alternative supplement is detailed in Table C.1. 

90
 
c. Protocols For PrePArAtion oF selected F. tularensis culture mediA 

C.2.4	 Preparation	of	plates 

1. Aseptically add 100 ml sterile haemoglobin solution and 2 ml of reconstituted growth 

supplement to 100 ml of GCII agar base medium. 

2. Mix gently but carefully using a magnetic stirrer and avoid the formation of air bubbles (i.e. 

foam) in the agar. 

3. Dispense 20 ml volumes of medium into each sterile Petri dish to achieve a uniform agar 

depth of 3–4 mm. 

4. Replace the lids on the Petri dishes and allow the medium to stay at room temperature for 

several hours. Place plates in a plastic bag and store at 4 °C. 

C.2.5	 Quality	control 

Inoculated GCII agar with haemoglobin and IsoVitaleX should support the growth of F. tularensis (e.g. the avirulent and fastidious strain ATCC 6223) after incubation at 37 °C in ambient 

air or in a 5 % CO2

 humid atmosphere for 48 h. 

Table	C.1 

Chemically-defined enrichment supplement for F. tularensis (Department of Clinical 

Bacteriology, Umeå University Hospital, Sweden) 

Ingredient	 Weight	(g)	per	litre	ultrapure	water 

Vitamin B12 0.01 

L-Glutamine 10.0 

Glucose 100.0 

Nicotinamide adenine dinucleotide 0.25 

Co-carboxylase 0.10 

Thiamine hydrochloride 0.003 

Ferric nitrate 0.02 

L-Cysteine hydrochloride 25.9 

Adenine sulphate 1.5 

Guanine hydrochloride 0.03 

L-Cystine 1.10 

p-Aminobenzoic acid 0.013 

91
 

d. reAgent list

 

D.	 Reagent	list 

Table	D.1 

Diagnostic reagents for F. tularensis 

Reagents:	commercially	available	 	 

Francisella tularensis antiserum (for slide and Becton Dickinson Diagnostic Systems 

tube agglutination) Loveton Circle Sparks MD USA 21152 

Phone: 001 410 316 4000; 

www.bd.com/ds/index.asp: Catalogue number: 240939 

Anti-Francisella tularensis LPS (clone FB11) 	 Advanced ImmunoChemical Inc 

Anti-Francisella tularensis LPS (clone T14)	 105 Claremont Ave Long Beach CA USA 90803 

Phone: 001 562 434 4676; 

www.advimmuno.com; Catalogue number: G3-T4E1014 

Hybridoma against F. tularensis	 American Type Culture Collection (ATCC) 

subsp. holarctica (LVS)	 P.O.Box 1549 Manassas VA 20108 USA 

Phone: 001 703 365 2700; 

www. atcc. org; Catalogue number: HB-10830 

Monoclonal antibody anti-F. tularensis Senova GmbH 

Monoclonal antibody anti-F. tularensis-HRP Winzerlaer Strasse 2 07745 Jena Germany 

conjugate Phone: +49 (0) 3641 508 508; 

www. senova. de 

Diagnostic Kit for Tularemia (Souprava k Bioveta a.s. 

diagnostice tularemie) including F. tularensis Komenskeho 212 CZ-68323 Ivanovice na Hane 

antigen for agglutination and F. tularensis Czech Republic 

control antiserum Phone: +420 517 318 599; www.bioveta.cz 

Reagents:	available	from	reference	laboratories 

FITC-conjugated rabbit-anti F. tularensis Centers for Disease Control and Prevention (CDC) 

polyclonal serum (DFA); formalin-fixed Division of Vector-Borne Infectious Diseases, Bacterial 

F. tularensis cells (microagglutination); Zoonoses Branch, Diagnostic and Reference Laboratory 

positive control anti-F. tularensis serum Fort Collins CO USA 

(microagglutination) Phone: 001 970 221 6400 

F. tularensis clones (positive controls) fopA, 	 Dr. Fran Nano University of Victoria, Victoria, 

iglC, pdpD BC V8W 3P6 Canada 

Phone: 001 250 721 7074 

Strains	 Source 

ATCC 6223 Francisella tularensis subsp. American Type Culture Collection (ATCC) 


tularensis (B-38); avirulent strain P.O. Box 1549 Manassas VA 2010
 

ATCC 25015; ATCC 25016; ATCC 25017; 


ATCC 25018 Francisella philomiragia
 

93
 

E.	PACkAGInG	AnD	TRAnSPORT	OF	InFECTIOUS	SUBSTAnCES	AnD	DIAGnOSTIC	SPECIMEnS	

E	 Transport	of	specimens	and		 

	 cultures	of	F. tularensis 

The transport of infectious substances, including F. tularensis, is highly regulated. Implementation of regulations is mandatory. To help understand the current regulatory framework 

and support compliance with current international regulations for the transport of infectious 

substances and patient specimens by all modes of transport, WHO has developed the document Guidance on regulations for the Transport of Infectious Substances 2007–2008, World Health 

Organization, 2007, applicable as from 1 January 2007. We recommend the reader to consult 

the above document for details regarding the transport of infectious substances. 

Infectious substances are divided into two categories, A and B. A flowchart for the classification of infectious substances and patient specimens is given in Figure E.1. 

According to applicable transport regulations, cultures (as defined in the transport regulations) of F. tularensis are classified as Category A, while specimens of F. tularensis not in the 

form of cultures are assigned to Category B. 

E.1	 Shipping	F.	tularensis 

It is the responsibility of the shipper to ensure the correct classification, packaging, labelling 

and documentation of all samples containing F. tularensis destined for transport. 

All specimens to be transported must be packaged according to applicable regulations, as 

described in Guidance on regulations for the Transport of Infectious Substances 2007–2008, World 

Health Organization, 2007, applicable as from 1 January 2007. 

Infectious substances in Category A may only be transported in packaging that meets the 

United Nations class 6.2 specifications and complies with UN Packing Instruction P620 (PI602 

air mode). This ensures that strict performance criteria are met. There is no comprehensive 

list of suppliers for packaging that complies with these packing instructions; however, an 

Internet search using search terms such as ’UN Packaging’ and ’UN infectious substance 

packaging’ provides the appropriate information. 

Maximum net quantities for packages vary depending on the transport mode. For surface transport, there is no maximum quantity per package. For air transport, the limits per 

package are 50 ml or 50 g for passenger aircraft and 4l or 4 kg for cargo aircraft. Note: hand 

carriage of Category A and Category B infectious substances and transport of these materials 

in diplomatic pouches is strictly prohibited by international air carriers! 

For substances in Category B, it may be possible to source packaging locally rather than 

finding an authorized supplier, provided that the packaging manufacturer and the shipper 

can comply fully with the requirements of P650. Testing documents are not required, however. Dangerous goods documentation (including a shipper’s declaration) is not required. 

For surface transport, there is no maximum quantity per package. For air transport, no 

primary receptacle shall exceed 1l (for liquids) or 1 kg (for solids) and the volume shipped per 

package shall not exceed 4l or 4 kg. Note: hand carriage of Category A and Category B infectious substances and transport of these materials in diplomatic pouches is strictly prohibited 

by international air carriers! 

E.2	Training 

Persons engaged in the transport of dangerous goods, including infectious substances, shall 

have received training commensurate with their responsibilities. For transport of Category 

A infectious substances, personnel must undergo training in accordance with the modal 

requirements. This can include attendance at approved courses and passing examinations. 

Further information should be obtained from the local transport authorities. 

95
 
 

wHo guidelines on tulArAemiA 

Figure	E.1	 Flowchart	for	the	classification	of	infectious	substances	and	patient	specimens 

	 

Not subject to the 

transport requirements 

for dangerous goods 

unless meeting the 

criteria for another 

division or class 

YES NO	OR	 

UNKNOWN 

Has an informed professional judgment based on 

the known medical history, symptoms and individual 

circumstances of the source, human or animal, and 

endemic conditions determined that there is only 

minimal likelihood that pathogens are present? 

NO YES	OR	 

UNKNOWN 

Does it meet the definition of a 

Category A substance? 

NO	OR	 

UNKNOWN

Is it known not to contain infectious substances? 

Have any pathogens present been neutralized or inactivated, so that they no longer pose a health risk? 

May it contain microorganisms that are non-pathogenic to humans or animals? 

Is it in a form in which any pathogens present have been neutralized or inactivated such that they no 

longer pose a health risk? 

Is it an environmental sample (including food and water sample) that is not considered to pose a 

significant risk of infection? 

Is it a dried blood spot? 

Is it a faecal occult blood screening test? 

Is it decontaminated medical or clinical waste? 

Is it for transfusion or transplantation? 

YES 

Substance for classification 

Reproduced from the Guidance on regulations for the transport of infectious substances 2007–2008, WHO, 2007. 

Subject to 

‘Exempt human or 

animal specimen’ 

provisions 

UN 3373 Biological 

substance, Category B 

UN 2814 Infectious 


substance, affecting 


humans, or
 

UN 2900 Infectious 


substance, affecting 


animals only
 
e. trAnsPort oF sPecimens And cultures oF F. tularensis 

E.3	References 

Guidance on regulations for the Transport of Infectious Substances 2007–2008, Geneva, World 

Health Organization, 2007 (http://www.who.int/csr/resources/publications/biosafety/WHO_ 

CDS_EPR_2007_2/en/index.html). 

97
 

F. cHecklist For outBreAk investigAtion

F.	 Checklist	for	outbreak	investigation 

F.1	 Preparations 

F.1.1	 Conditions	under	which	a	tularaemia	outbreak	investigation	may	be	useful 

1. Clinical manifestations support tularaemia. 

2. Single cases have been confirmed by laboratory methods. 

F.1.2	 Outbreak	investigation	team 

An outbreak investigation team could be composed of the following members covering different tasks: 

1. Team leader: coordinates team activities, communicates (information, agreements) with 

the national/international health authorities. 

2. Epidemiologist: designs epidemiological study design, ensures that the right type of data is 

collected in a manner that is suitable for epidemiological investigation, analyses the epidemiological data, prepares a report to convey the information. 

3. Microbiologist: supports appropriate collection, storage and shipment of clinical and environmental specimens; undertakes microbiological analyses. 

4. Ecologist 	 and/or veterinarian and/or entomologist: assesses environmental risks for 

Francisella infection and evaluates the involvement of animal species and vectors in the 

tularaemia outbreak. 

5. Physician: supports the treatment of patients and preventive measures to reduce the risk of 

exposure to infection sources, collects clinical specimens for further analyses. 

6. Laboratory staff: support first field analyses, preparation of specimens, etc. 

7.	 Local health authorities and personnel: of high priority for inclusion as they usually know 

the local situation and specific circumstances. 

8. Translator, if needed. 

F.1.3	 Coordination	with	local	health	authorities 

1. Existence of a request for support made by local authorities. 

2. Consideration 	 of local specificities, e.g. political and social environment, safety considerations in crisis-afflicted areas, laboratory and clinical capacities. 

3. Agreements by local authorities to undertake an outbreak investigation in compliance with 

regulations for investigations of humans and collection of environmental samples, including animal carcasses. 

4. Local conditions for supporting the outbreak investigation including personnel to undertake 

field investigation, cars, laboratory facilities (with appropriate safety level), established 

laboratory methods for diagnosis of tularaemia and detection of Francisella, possibilities for 

storage of samples, accommodation for the investigation team. 

The investigation must be adapted to local prerequisites and regulations. Instruments and 

materials need to be transported into the affected region. Administrative requirements need 

to be initiated, including regulations for transportation of specimens to specialized laboratories. 

99
 
wHo guidelines on tulArAemiA 

F.1.4	 Action	to	be	taken	by	the	investigation	team	before	going	to	the	field 

1. Communicate with local authorities (objectives of the planned investigation, timeframe, 

type of samples to be collected, number of samples to be collected, expected results, ownership of samples etc.). 

2. According to the identified aims, set up the investigation team (see above). 

3. Vaccinate the investigation team against tularaemia, if possible. 

4. If no biosafety level 3 laboratory facility is available in the affected area, only preliminary 

investigations or investigations with inactivated specimens can be done. For further investigation, including isolation of the pathogen by cultivation, the specimens have to be shipped to specialized laboratories. 

5. Full protection is not needed for epidemiological investigations of, for example, households 

or patients; for the latter, normal hygienic measures should be sufficient. 

6. Respect visa and vaccination regulations when the outbreak investigation is international. 

F.1.5	 Equipment 

1. Personal computer (including power adaptor, storage media (Floppy-disks, CD-ROM, 

USB-key). 

2. Systems for drawing blood samples. 

3. Swabs and appropriate transportation materials. 

4. Instruments to handle carcasses and environmental samples. 

5. Sufficient freezing capacity (dry ice) to store and ship samples, in appropriate containers (in 

compliance with regulations for shipment of dangerous goods). 

6. Materials and instruments for laboratory investigations when these are to be performed 

locally. It should be recognized that in certain regions technical support is completely unavailable. 

F.2	 Outbreak	investigation	in	the	field 

Data available from epidemiological and laboratory investigations should be analysed 

promptly to obtain, as a minimum, information on the size and duration of the outbreak, on 

the source of infection and suspected transmission routes of the pathogen. 

Once it has been decided that an outbreak investigation for tularaemia is to be carried out, the 

following steps may be undertaken. 

1. Contact local authorities: the team leader communicates with the local authorities regarding the coordination of work with local staff, procedures for reporting and the communication of results. 

2. Enquire about the epidemiological situation: contact patients and local physicians (see 

sample questionnaire for case-control study), analyse laboratory data (for evidence of their 

plausibility). 

3. Formulate a hypothesis on source and cause of the outbreak, e.g. who was affected; where 

and when did the suspected outbreak occur? 

4. Construct a time schedule for the investigation. 

5. Make preliminary recommendations for treatment and prevention of the disease. 

6. Set up facilities for diagnosis and/or for shipping specimens to a different laboratory for 

confirmation/further analyses (see chapter 6). 

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F. cHecklist For outBreAk investigAtion 

7.	 Take samples (clinical and environmental). All samples should be frozen as soon as possible if not analysed within 24 hours. An appropriate system for labelling and identification 

of the samples should be used, e.g. barcodes, numbering. 

8.	 Analyse samples. 

9.	 Report the results according to the plan of action agreed upon with the local authorities. 

10.	 Write up recommendations for: the limitation of the outbreak; avoiding new cases; adequate treatment of patients; prevention of further outbreaks; and in the longer term for 

improvement of the surveillance and diagnosis of tularaemia. 

11.	 Agree on further analysis of specimens and epidemiological data. 

12.	 Educate local health workers and authorities if requested. Especially in poorly developed 

areas, long-term support could be important to prevent or recognize early new outbreaks 

of tularaemia. This could include training of physicians for recognition of the disease, 

training of health-care workers in epidemiological analysis and of laboratory staff in 

diagnostic methods for tularaemia. 

13.	 Publish the results to a wider audience (public media, scientific literature). 

F.3	 Example	for	a	tularaemia	outbreak,	Kosovo	(Serbia) 

An outbreak of tularaemia in UN Administered Province of Kosovo (Serbia) in 2000 was 

investigated by an international team brought together by the WHO Regional Office for 

Europe (Reintjes et al., 2002). The following information formed part of the report of the outbreak investigation which gathered together useful practical experiences. The main part was 

a case-control study to assess the risk factors for infection with F. tularensis. 

F.3.1	 Protocol	for	case-control	study	of	tularaemia	infection	used	by	the	 

investigation	team 

1.	 After arrival at a case household: 

a) Ask whether the case was living in this house in the month before the illness started. 

If yes, please continue the questionnaire and ask for control households (see definition 

below). If no, please continue, but do not select controls for the case household. 

b) Ask whether either the person with suspected tularaemia infection or the person 

preparing the food is available to give a blood specimen; if not available, visit again 

another time/day. 

c)	 In the household of a case, a blood specimen should be taken from all family members 

who have experienced fever and cervical lymphadenitis after 1 November 1999 (the 

probable date of onset of the outbreak). 

d) After taking blood samples and completing the questionnaire, enquire about the two 

closest neighbours. If these neighbours use the same water supply, do not visit them 

but ask for other neighbours. 

2.	 After arrival at a control household: 

Ask whether anybody in the family has had fever plus lymphadenitis colli since the first 

identification of index case. If no, continue with questionnaire and take blood from person preparing food. If yes, use as new case – take blood from all family members with 

symptoms. Select new control for both cases (you now have two cases and need four 

controls). 

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wHo guidelines on tulArAemiA 

102
 

Case definition 

Any person who has had lymphadenitis colli after 1 December 1999, with laboratory confirmation of tularaemia (see section 6.1). 

Definition of a case household 

A household with one or more family members with laboratory confirmation of tularaemia 

infection and with lymphadenitis colli after confirmation of tularaemia. 

Definition of a control household 

Closest household to a case household during one month before onset of illness in the case, 

but with different water supply, with all family members free of high fever (at least 38 °C) 

and lymphadenitis colli, or, if any family member with lymphadenitis colli, with laboratory 

confirmation that all family members are negative for tularaemia infection. 

Questionnaire	for	case-control	study:	risk	factors	for	tularaemia	in	 

households 

CONTACT DETAILS: 

Name: 

Date of birth (day; month; year): 


Village:
 

Case household (number):
 

FOOD CONSUMPTION:
 

In the month before case became ill; did you eat any of the following? 


m Uncooked vegetables
 

m Meat of hares
 

m Pershut (dried ham)
 

m Fruits (e.g. apples)
 

m Milk/cream
 

WATER SUPPLY
 

In the month before illness what water did you use for drinking? (boiled;chlorinated):
 

m Central water (piped) 


m Bottled water
 

m Personal well-water
 

m Water from collective well
 

m Water from tanks
 

m Streams
 

In the month before illness what water did you use for food preparation? (boiled; 


chlorinated) 


m Central water (piped)
 

m Bottled water
 

m Personal well-water
 

m Water from collective well
 

m Water from tanks 


m Streams
 
F. cHecklist For outBreAk investigAtion 

ANIMALS
 

In the past year have you seen large numbers of rodents? If yes, when?
 

m Inside house:
 

m Outside house: 

What kind of rodent did you see?
 

m Mouse in the field
 

m Mouse in the house
 

m Rat in the field
 

m Rat in the house or yard
 

In the past year have you seen sick or dead rodents? If yes
 

m Inside the house?
 

m Outside the house in the private yard?
 

m In your water supply?
 

m Do you have any domestic animals (pigs, cow, hens)? 


m Did you use flour from a humanitarian organization?
 

m If yes, which organization?
 

m What country of origin is this flour?
 

m Do you have dogs or cats in your household?
 

m Did you have contact with wild animals or carcasses?
 

m Did you get bites by ticks, fleas, bugs, mosquitoes, flies, etc.?
 

m If yes; did you get inflammation or an ulcer at the site of the bite?
 

OBSERVATIONS OF WATER SUPPLY (ASK TO BE SHOWN THE WATER SUPPLY) 


m Well?
 

m Piped water?
 

m If well; is it protected against rodents?
 

m If well; is it protected against water leakage?
 

FOOD STORAGE (PLEASE ASK TO BE SHOWN THE FOOD STORAGE ROOM) 


m Is the food storage room protected against rodents?
 

m Do you see faeces of rodents near the food or food touched/partly eaten by rodents?
 

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g. Protocols For trAPPing And sAmPling smAll mAmmAls

105
 

G.	Protocols	for	trapping	and	

	 sampling	small	mammals 

Field investigations involve capturing the small mammal hosts of tularaemia for collection of 

blood, ectoparasites (ticks and fleas), and occasionally tissue samples. The primary goal is to 

collect sufficient numbers of animals representative of small mammal communities at study 

sites to determine which species of animals are infected with F. tularensis. The surveys include 

collection of live rabbits and rodents, and ectoparasites from these animals. 

Much of the information below is derived from Sudia et al., (1970), Mills et al., (1995) and 

Dennis et al., (1999). National regulations on the ethical prerequisites for animal research 

need to be considered. 

G.1	Trapping	of	small	mammals 

1.	 Set a combination of Tomahawk and Sherman live traps in areas of special interest. Check 

the proper functioning of the traps before setting. 

2.	 Place upholstery cotton in each trap during cold weather to reduce the potential for hypothermia. Cover traps with canvas on top and sides to provide additional protection during 

adverse weather. 

3.	 Bait traps with rolled oats, peanut butter, or other appropriate items that are attractive to 

the animals. 

4.	 Check traps at 1–2-hour intervals to ensure that animals do not suffer from prolonged 

exposure to heat or other threatening conditions. Check the traps as early in the morning 

as possible. 

5.	 Wear rubber gloves while handling the traps with captured animals. 

6.	 Label traps with captured animals with information about the location of trap on a strip 

of tape attached to the top of the trap. 

7.	 Place the traps with captured animals in double plastic collecting bags. Place collecting 

bags with the captured rodents in a cool area out of the sun until all traps have been 

checked. 

8.	 Replace collected traps with spare baited traps. 

9.	 Transport the captured rodents to the field station for processing as soon as possible. 

10.	 Surviving animals should be released as soon as possible, at or near the original point 

of capture (usually within 1–2 hours after the traps have been brought to the processing 

site). 

11.	 Traps should be decontaminated after each trapping session. 

G.2	Protocols	for	collecting	diagnostic	samples	from	small	mammals 

G.2.1	 Anaesthesia 

1.	 Captured animals are anaesthetized with Isoflurane or Ketamine depending on the animal species. All handling of animals and other possibly contaminated materials should 

be done with caution and using personal protection such as gloves, full face protection 

(FFP3) masks and other measures according to the risk assessment. All manipulations 

with sharp tools such as dissection of carcasses or taking blood samples with needles 

must be done by trained personnel and with special caution. 
wHo guidelines on tulArAemiA 

2. Small rodents captured in Sherman traps can be easily removed to 12” x 12” ziplock bags 

containing a cotton or gauze with Isoflurane. Animals in Tomahawk traps may be anaesthetized by placing the entire trap into a large plastic bag containing anaesthetic-soaked 

gauze or in a tool box. Alternatively, insert a large pair of forceps through the mesh of the 

trap, grasp the skin of the animal, and inject it with ketamine/xylazine (10:1) with a needle 

and syringe. 

3. If an animal begins to recover during collection of ectoparasites or during collection of 

blood samples, it should be re-anaesthetized immediately. If an animal escapes while 

attempting to anaesthetize it, personnel should not try to recapture it with their hands 

because of the high risk of being bitten. 

G.2.2	 Collecting	ectoparasites 

1. The anaesthetized animals are placed in a pan and processed for fleas and ticks. 

2. Fleas are removed from anaesthetized animals by brushing with a fine toothed comb. 

Attached ticks are removed with fine forceps. 

G.2.3	 Taking	blood	samples 

1. Label the cryovials for blood samples before collecting blood. Only screw-cap plastic cryovials with external thread should be used. 

2. Blood samples are collected from anaesthetized animals by bleeding from the retroorbital 

plexus or by cardiac puncture. Use a sterile heparinized tube or Pasteur pipette for retroorbital bleeding. Use only gentle pressure to avoid breaking the tube. Use small hematocrit 

bulbs to eject the blood that may remain in the capillary tube into a cryovial. 

3. If the animal has died only recently, blood can be obtained directly from the heart by cardiac puncture. This procedure should be also done when retroorbital bleeding is impractical. For example, large animals can be difficult to bleed by the retroorbital technique, and 

cardiac puncture is likely to be a more satisfactory procedure. 

The cardiac puncture procedure is as follows: 

a) Choose an appropriately sized syringe and needle based on the size of the animal. 

For mice, a 1 cc syringe with a 25G x 5/8 inch needle (or similar small gauge needle) 

is preferable. Larger syringes are likely to create too much vacuum and collapse the 

heart chambers of these small animals. For larger animals, such as squirrels and rabbits, a 3 cc syringe can be used, although a 1 cc syringe fitted with a 23G needle that is 

1–1.5 inches in length is satisfactory. 

b) Lay the animal out flat in a white enamel pan. 

c)	 Insert the needle just below the base of the sternum and slightly to the left of centre. As 

you insert the needle, gently pull back on the plunger until blood appears at the base of 

the needle. Once the needle is correctly positioned and blood is entering the syringe, 

gently pull back on the plunger, being careful not to withdraw the needle from the 

heart. Pulling back too quickly on the plunger will create excessive vacuum and collapse 

the heart, interfering with the collection of blood and increasing the chance that a blood 

clot will form in the needle. Continue to remove blood until it no longer flows freely into 

the syringe. 

d) Carefully squirt the blood into a cryovial or onto a Nobuto strip. 

106
 
g. Protocols For trAPPing And sAmPling smAll mAmmAls 

G.2.4	 Dissection	for	organ	tissue	recovery 

1. Flame a pair of scissors and forceps over an alcohol burner. 

2. Place the animal ventral side up in a clean pan. 

3. Squirt the ventral surface with 70% ethanol from a squeeze bottle and wipe with a gauze 

square. 

4. Pinch the skin of the lower part of the abdomen with forceps and lift it. Place the scissors 

below the forceps and cut through the skin and abdominal musculature. Insert one blade 

of the scissors into the incision and make cuts on each side of the abdominal wall, and 

then pull the cut skin and musculature back above the diaphragm to completely expose the 

abdominal cavity. 

5. Using blunt-ended, nontoothed, sterile forceps, lift the stomach to expose the spleen. Grasp 

the spleen with the forceps and gently pull to remove it and place it in a cryovial. Using 

the same forceps (flame sterilized between each organ; ensure that the forceps are cooled 

between each usage) grasp the kidneys, one at a time, and place them into a second cryovial. With the forceps, grasp a portion of liver of appropriate size to fit into the third cryovial. Additional samples such as enlarged lymph nodes or liver lesions may be collected as 

needed. 

6. After the processing of rodents has been completed, the processing area should be carefully 

decontaminated and carcasses must be collected for later incineration or other appropriate 

disposal. 

G.3	References 

Dennis D et al. (1999). Plague manual: epidemiology, distribution, surveillance, control. Geneva, 

World Health Organization (WHO/CDS/CSR/EDC/99/2). 

Mills J et al. (1995). Methods for Trapping & Sampling Small Mammals for Virologic Testing. Atlanta, US Department of Health & Human Services. 

Sudia W et al. (1970). Collection & Processing of Vertebrate Specimens for Arbovirus Studies. 

Atlanta, US Department of Health, Education, and Welfare. 

107
 

index

 

 

 

 

Index 

Advanced ImmunoChemical Inc 93
 

aerosols
 

avoidance of exposure 40
 

disease contracted from 14–15
 

infective dose 1, 41
 

laboratory biosafety 41, 42
 

post-exposure prophylaxis 22
 

agar, culture on 30, 62–63
 

see also culture media
 

age distribution 4
 

agglutination tests 30, 73–74
 

air systems, laboratory 42
 

air transport 95, 96, 97
 

American Type Culture Collection (ATCC) 


45, 93
 

aminoglycosides 21, 22–23
 

ampicillin 47
 

anaesthesia 105, 106
 

animals (mammals)
 

avoidance of contact 40
 

capture/trapping 39, 105
 

dead see dead animals
 

disease and pathology 8–9, 10
 

dissection for organ tissue recovery 107
 

ectoparasite collection 39, 106
 

human contact 3
 

models of disease 44
 

reservoirs 5, 6–7
 

sample collection 39, 105–106
 

serology in wild 9
 

surveillance 36
 

transmission by 7, 11, 13
 

anthrax 18
 

antibiotics 21–25
 

choice of agent 22–25
 

minimal inhibitory concentrations 22, 


26
 

post-exposure prophylaxis 22, 43
 

recommended treatment 21–22
 

resistance 25, 45
 

antibodies 11, 30, 44
 

see also serology
 

antigen detection assays 31, 64–70
 

antimicrobial susceptibility tests 32, 81–82
 

arthropods
 

avoidance of exposure 40
 

diagnostic tests 28
 

vectors 5–6
 

aspirates 27, 28, 29
 

Australia 4, 5
 

autoclaves 42, 43
 

autopsy specimens
 

animals 28, 39, 107
 

humans 28, 29
 

bactericidal agents 22, 23, 24, 82
 

bacteriology 30
 

bacteriostatic agents 23–24, 82
 

beavers 7, 9, 10
 

Becton Dickinson Diagnostic Systems 93
 

beta-lactam antibiotics 21, 25, 45
 

biochemical biotyping 32
 

biological warfare 3, 32
 

biopsies, tissue 27–28, 29
 

biosafety 41–43, 61
 

levels 34, 41–42
 

bioterrorism 3, 25, 38
 

biotyping
 

biochemical 32
 

molecular 32–33, 82–87
 

Bioveta a.s. 93
 

blood chemistry 1.0
 

blood samples
 

animals 28, 105–107
 

human 28, 29, 30
 

brucellosis 19
 

buffer recipes/storage 78–80
 

buffered charcoal yeast extract (BCYE) 31, 


62
 

C-reactive protein 17
 

cabinets, biosafety 42, 43
 

Caedibacter taeniospiralis 4
 

Canada 3, 5, 7
 

capture, small mammals 39, 105
 

carcasses, animal see dead animals
 

cardiac puncture procedure 106–107
 

case–control studies 38, 101, 102
 

109
 
 

 

 

 

 

 

 

 

 

 

 

wHo guidelines on tulArAemiA 

case definitions 27, 102
 

case descriptions, naturally-infected 


animals 8
 

case households 101, 102
 

cat-scratch disease 18
 

cation-adjusted Mueller-Hinton broth 


(CAMHB) 32, 82
 

cats 8
 

ceftriaxone 25
 

cell-mediated immunity 11, 23, 44
 

Centers for Disease Control and 


Prevention (CDC) 93
 

centrifuges 42
 

CHAB see cystine heart agar with 


chocolatized blood
 

chest X-rays 14, 16
 

childhood tularaemia
 

clinical manifestations 15
 

treatment 21, 24, 25
 

China 5
 

chloramphenicol 22, 23, 45, 82
 

chocolate agar 30, 62, 81, 90
 

Chrysops flies 5, 6
 

ciprofloxacin 21, 22, 24, 25, 82
 

citrulline ureidase 32, 33
 

clindamycin 25
 

Clinical and Laboratory Standards 


Institute (CLSI) 82
 

co-trimoxazole 25
 

coating buffer 78
 

complications 15
 

confirmed cases 27
 

conjunctivitis 14
 

coyotes 9
 

culture 30–31, 62–64
 

on agar 31, 62–63
 

mouse inoculation 31, 63–64
 

culture media 31, 62–63
 

antimicrobial susceptibility testing 81
 

growth characterization 32
 

preparation protocols 89–91
 

cysteine/cystine-supplemented media 31, 


32, 33
 

cystine heart agar with chocolatized blood 


(CHAB) 31, 62, 63
 

antibiotic-supplemented (CHAB-A) 


31, 62, 89
 

preparation 89–91
 

cytomegalovirus (CMV) disease 18
 

dead animals
 

blood sampling 105–107
 

collection and transport 39
 

disease transmission 1, 7, 14
 

dissection for organ tissue recovery 107
 

observations at location 39
 

specimens 28, 39
 

surveillance for 36
 

decontamination 42
 

deer flies 6, 28
 

Dermacentor ticks 5, 6, 28
 

diagnosis 27–34
 

case definitions 27
 

differential 17, 18–19
 

specimens see specimens, diagnostic
 

diagnostic tests 3, 30–32
 

field conditions 40
 

protocols 61–80
 

reagents 93
 

differential diagnosis 17, 18–19 

direct fluorescent antibody (DFA) assay 

31, 64–65, 93
 

disasters, natural 3
 

documentation, transport 95
 

dogs 8
 

dormouse 10
 

doxycycline 21, 22, 24, 82
 

E-test 32, 81–82
 

ecologist 99
 

ectoparasites, collecting 39, 106
 

ELISA see enzyme-linked immunosorbent 


assay
 

endemic areas 1, 5
 

endocarditis 15
 

entomologist 99
 

environmental reservoirs 7
 

environmental specimens 28, 39
 

enzyme-linked immunosorbent assay 


(ELISA) 

antibody-directed 30, 74–78 

antigen capture (cELISA) 31, 66–70 

buffer recipes/storage conditions 78–80 

epidemic disease intelligence 36
 

epidemics see outbreaks
 

epidemiologist 99
 

epidemiology 5–9
 

epizootics 7, 37
 

Epstein-Barr virus (EBV) infection 18
 

110
 
 

 

 

 

 

 

 

 

 

 

 

index 

equipment, outbreak investigation 100
 

erythema multiforme 17
 

erythema nodosum 17
 

erythrocyte sedimentation rate 17
 

erythromycin 25, 45
 

Europe 5, 6, 10
 

excrement, animal 7, 39
 

experimental tularaemia 8–9
 

faeces, animal 7, 39
 

farming activities 14, 40
 

fatality rates 11, 14
 

fatty acids 29
 

fever 11, 12
 

of unknown origin 15–17, 21
 

field investigations 38–39, 100
 

example 101–103
 

laboratory methods 40
 

sample collection 39–40
 

trapping/sampling small mammals 


105–107
 

field laboratory 40
 

Finland 5, 6, 14, 15, 17
 

fleas 6, 106
 

flies 5, 6
 

foodborne infection 13, 14, 40
 

formalin-fixed specimens 29, 39
 

Francisella novidica see Francisella 


tularensis subspecies novidica
 

Francisella philomiragia 3, 4
 

characterization 31, 32
 

human disease 12
 

Francisella tularensis 1, 3–4
 

bacteriology 29
 

in environment 7
 

genetic manipulation 45
 

handling 41–45
 

identification see diagnostic tests
 

inactivation methods 42
 

isolates 4
 

strains see strains
 

subspecies 4, 32–33
 

supplemental characterization 31–33, 


81–87
 

taxonomy 3–4
 

Francisella tularensis subspecies holarctica
 

(type B) 4
 

antibiotic sensitivity 22, 24–25, 26
 

experimental infections 9
 

human disease see type B tularaemia
 

identification 32–33, 82–87
 

laboratory biosafety 41
 

supplemental tests 31–33
 

transmission 5, 11
 

Francisella tularensis subspecies 


mediasiatica 4
 

characterization 32, 33
 

human disease 12
 

laboratory biosafety 41
 

Francisella tularensis subspecies novidica
 

4, 5
 

characterization 32, 33
 

experimental studies 44
 

human disease 12
 

laboratory biosafety 41
 

Francisella tularensis subspecies tularensis
 

(type A) 1, 4
 

antibiotic sensitivity 22, 25, 26
 

experimental infections 8, 9
 

human disease see type A tularaemia
 

identification 32, 84–87
 

laboratory biosafety 41
 

supplemental tests 31–33
 

transmission 5, 7, 11
 

Francisellaceae 3–4, 7
 

frozen specimens 39
 

Ft-M19 assay 32–33, 82–84
 

fulminant disease 15, 17
 

gastrointestinal symptoms 11
 

GC II agar base with 1% haemoglobin and 


1% IsoVitaleX 30, 62, 81, 90–91
 

gender distribution 4
 

genetic manipulation 45
 

gentamicin 21, 22, 23, 82
 

geographical distribution 4, 5, 6
 

glandular tularaemia 12,13, 16
 

Global Outbreak Alert and Response 


Operations 36
 

glucose cysteine blood agar 81
 

glycerol fermentation 32, 33
 

Gram stain 61–62
 

grivet monkeys 9
 

ground squirrels 7
 

growth characteristics, bacterial 32
 

guanidine thiocyanate (GuSCN) buffers 


 70, 71
 

guinea-pigs 8
 

111
 
 

 

 

 

 

 

 

 

 

 

wHo guidelines on tulArAemiA 

hamsters 10
 

handling measures 41–45
 

hantavirus disease 19
 

hares 6–7, 10
 

health authorities/workers, local 99–100, 


101
 

health monitoring, laboratory staff 43
 

hedgehogs 10
 

hepatic failure 15
 

hilar lymphadenopathy 15, 16
 

HIV disease 15, 17, 18
 

horse flies 6
 

households, case and control 101
 

human-to-human transmission 5, 11, 35
 

humans
 

clinical expression 11–17
 

diagnostic specimens 27–28
 

disease surveillance 36
 

specimen collection and transport 

28–29
 

transmission to 7, 11
 

treatment 21–25
 

immunocompromised individuals 12, 


15–17, 21
 

immunohistochemical staining 31
 

immunological response 9, 11, 44
 

incubation buffer 79
 

incubation period 11
 

infectious substances
 

category A 95–96
 

category B 95–96
 

classification 95–96
 

defined 95
 

international regulations for transport 


95
 

packaging 95
 

infective dose 1, 41
 

influenza 19
 

international regulations, transport of 


infectious substances 95
 

international surveillance networks 36
 

ISFtu2 assay 32–33, 84–88
 

isoflurane 106
 

isolates 4, 31
 

IsoVitaleX 90
 

Japan 5
 

kanamycin 45
 

Kazakhstan 4, 5
 

ketamine 106
 

kit foxes 9
 

Kosovo, United Nations Administered 


Province of Kosovo (Serbia) 3, 12, 


37–38, 39, 101–102
 

laboratory
 

biosafety levels 34, 41–42
 

decontamination and sterilization 42, 


43
 

facilities, outbreaks 100
 

field 40
 

physical requirements 42
 

safety measures 41–43
 

laboratory-acquired infections 34, 41
 

LVS vaccine and 43–44
 

laboratory animals, course of disease 8–9
 

laboratory diagnostics 27–34
 

laboratory staff
 

health monitoring 43
 

outbreak investigation 99
 

post-exposure treatment 43
 

training 42
 

lemmings 10
 

leptospirosis 19
 

levofloxacin 24, 82
 

lipopolysaccharide (LPS) 30, 44, 66
 

antibody titres 30, 77–78
 

extraction buffer 67
 

live vaccine strain (LVS) 43–45
 

animal studies 9, 44
 

clinical trials 44–45
 

laboratory biosafety 41, 42
 

liver enzymes 17
 

local health authorities/workers 99–100, 


101
 

LPS see lipopolysaccharide
 

LVS see live vaccine strain
 

lymph node enlargement/lymphadenitis 


13, 15
 

cervical (colli) 14, 16, 102
 

hilar 15, 16
 

lymphocyte counts 17
 

lymphoma 19
 

mammals see animals
 

Martha’s Vineyard, USA 1, 5, 7
 

112
 
 

 

 

 

 

 

index 

meningitis 17, 22
 

Mexico 5
 

mice see mouse
 

microagglutination test 30, 73–74, 93
 

microbiologist 99
 

minimal inhibitory concentrations (MICs) 


22, 24, 25, 26
 

tests 32, 82
 

mink 6–7
 

models, animal 44
 

molecular biotyping 32–33, 82–87
 

molecular detection methods 30–31, 


70–73
 

moles 10
 

monkeys 9
 

monoclonal antibodies (mAb) 66, 68, 93
 

mortality rates 11, 14
 

mosquitoes 5–6, 28
 

mouse 7, 10
 

experimental infection 9, 44
 

inoculation test 31, 66–67
 

Mueller-Hinton agar with 1% IsoVitaleX 62
 

multi-locus variable number tandem 


repeat assay (MLVA) 32–33
 

muskrats 3, 7, 10
 

mycobacteriosis 18
 

Nano, Dr Fran 93
 

necropsy specimens see autopsy specimens
 

norfloxacin 24
 

notifiable disease reporting 36
 

oculoglandular tularaemia 13–14
 

Office International des Epizooties (OIE) 


37
 

oropharyngeal tularaemia 13, 14
 

outbreak investigation 100–103
 

prevention of spread 40
 

outbreaks
 

in biological warfare 38
 

field investigations see field 


investigations
 

historical 3
 

investigation 37–40, 99–103
 

management 35–40
 

natural 37
 

prevention of spread 40
 

response 36
 

rodent population size and 7
 

temporal variation 1
 

verification 36, 38
 

war-associated 3, 12
 

oxidase test 32, 33
 

packaging, for transport 95
 

pasteurellosis 18
 

pathology, in animals 8–9
 

PCR see polymerase chain reaction
 

penicillin 25
 

physician 99
 

plague 18
 

pneumonia 14–15, 16
 

atypical 19
 

polymerase chain reaction (PCR)
 

biotyping assays 32–33, 82–87
 

detection assays 31, 74–78
 

reagents 71
 

post-exposure prophylaxis 22, 43
 

prairie dogs 6–7, 8
 

pregnant women, treatment 21
 

presumptive cases 27
 

prevention
 

post-exposure chemotherapy 22, 43
 

spread of disease 40–41
 

protocols, diagnostic 61–80
 

protozoa 5, 11
 

pulsed-field gel electrophoresis (PFGE) 33
 

quinolone antibiotics 24–25, 26
 

rabbits 3, 7, 10
 

experimental infection 8, 44
 

rats 8, 10
 

reagents, diagnostic 93
 

relapse, disease 23–24
 

renal failure 17
 

reports, notifiable disease 36
 

reservoirs
 

environmental 7
 

mammals 6–7
 

respiratory specimens 27, 29
 

respiratory symptoms 11
 

respiratory tularaemia 13, 14–15, 17
 

laboratory acquired 41
 

prevention of spread 40
 

Rhesus macaques 9
 

rickettsiosis 18
 

rifampin (rifampicin) 22, 25
 

113
 
 

 

 

 

 

wHo guidelines on tulArAemiA 

risk factors, questionnaire 102
 

rodents 7
 

disease and pathology 9, 10
 

population size 7 


rRNA, 16S, gene sequences 3–4, 31–32
 

Russian Federation 3, 5, 7, 45
 

sample collection 61
 

captured animals 105–107
 

dead animals 39, 106
 

in the field 39–40
 

humans 28–29
 

samples, diagnostic see specimens, 


diagnostic
 

seasonality 4
 

Senova GmbH 93
 

sentinel surveillance 36
 

septicaemia 17
 

serology 30
 

protocols 73–80
 

studies in wild animals 9
 

serum specimens 27, 28, 29
 

sheep blood agar (SBA) 30, 32
 

Sherman traps 105, 106
 

shrews 10
 

signs, disease 12–17
 

16S rRNA gene sequences 3–4, 31–32
 

skin lesions 13, 17, 18
 

slide agglutination test 31, 66, 67
 

sodium azide 78
 

Soviet Union, former 3, 5–6, 7, 43
 

Spain 13, 16, 24
 

specimens, diagnostic 27–30
 

arthropods 28
 

classification 95, 96
 

collection see sample collection
 

environmental 28, 40
 

field 40
 

human 27–28
 

mammals 28
 

packaging and transport 95
 

transport 28, 29, 61
 

sputum specimens 29
 

staphylococcal skin lesions 18
 

sterilization 42–43
 

strains 4
 

identification 32–33
 

sources 45, 93
 

vaccine 43–45
 

streptococcal skin lesions 18
 

streptomycin 21, 22–23, 24, 82
 

resistance 25
 

subculture 32
 

subspecies 4
 

identification 32–33, 84–87
 

see also individual subspecies 

substrate buffer 80
 

subtyping see biotyping
 

supplemental characterization tests 


32–34, 82–87
 

surveillance 35–37
 

suspect cases 27
 

swabs 27, 29
 

Sweden 4, 5
 

human disease 15, 16, 17
 

transmission to humans 6, 7
 

symptoms 12–17
 

T-cell response see cell-mediated 


immunity
 

Tabanidae 6
 

talopoins 8
 

tamarins 8
 

TaqMan PCR assays 31, 84–87
 

taxonomy 3–4
 

team, outbreak investigation 99
 

temperature
 

bacterial inactivation 42
 

specimen storage 29
 

temporal variation, outbreaks 1
 

tetracycline 22, 23–24, 45, 82
 

resistance 25
 

thioglycollate-glucose blood agar (TGBA) 


30, 62
 

ticks 5–6, 7, 28
 

collecting 39, 106
 

tissue biopsies 27–28, 29
 

Tomahawk traps 105, 106
 

toxoplasmosis 18
 

training
 

laboratory workers 42
 

transport personnel 95
 

transmission 1, 5, 11, 38
 

airborne 13, 14–15, 40
 

by arthropods see vector-borne 


transmission
 

114
 
 

 

 

 

 

index 

by direct contact 11, 13
 

foodborne 13, 14, 40
 

by mammals 7, 11, 13
 

mechanical 6
 

prevention measures 40
 

waterborne see waterborne 


transmission
 

transport 95–97
 

dead animals 28
 

diagnostic specimens 28–29, 30, 61
 

international regulations 95–96
 

packaging 95
 

tick samples 39
 

trapping, small mammals 39, 105
 

treatment 21–25
 

post-exposure prophylactic 22, 43
 

tube agglutination test 30, 73
 

tul4 polymerase chain reaction assay 31, 


70–73
 

Turkey 14
 

Turkmenistan 4, 5
 

type A tularaemia 35
 

blood chemistry 17
 

clinical manifestations 11, 14–15
 

complications 15, 17
 

infectious agent see Francisella tularensis
 

subspecies tularensis
 

treatment 21, 25
 

type B tularaemia 35
 

blood chemistry 17
 

clinical manifestations 11–12, 15
 

complications 15, 17
 

infectious agent see Francisella tularensis
 

subspecies holarctica
 

treatment 21, 24–25
 

typhoidal tularaemia 13, 15, 41
 

ulceroglandular tularaemia 13, 16
 

complications 15, 17
 

prevention of spread 40
 

ulcers, primary 13, 16
 

ultraviolet (UV) irradiation 43
 

United States (USA) 3, 5
 

human disease 11, 12
 

vectors 5, 6
 

vaccination 21, 41, 43–45
 

vaccines 43–45 

vector-borne transmission 5–6, 11, 13
 

prevention 40
 

vectors 5–7 

ventilation, laboratory 42
 

veterinarian 99
 

virulence, subspecies 4
 

voles 7, 10
 

war-associated outbreaks 3, 12
 

washing buffer 79
 

water samples 40
 

waterborne transmission 5, 11
 

clinical form of disease 13, 14
 

prevention 40
 

white blood cell counts 17
 

WHO Laboratory Biosafety Manual 34, 42
 

World Animal Health Information System 


36, 37
 

World Health Organization (WHO)
 

laboratory biosafety 34, 41
 

surveillance network 36
 

transport regulations 95–96
 

World Organisation for Animal Health 


(OIE) 36, 37
 

X-rays, chest 15, 16
 

115