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17_Microbiological laboratory diagnostics

17_Microbiological laboratory diagnostics

Acta Tropica 165 (2017) 40–65

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Acta Tropica

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Microbiological laboratory diagnostics of neglected zoonotic diseases


Norbert Georg Schwarz a,∗, Ulrike Loderstaedt b, Andreas Hahn a, Rebecca Hinz c,
Andreas Erich Zautner b,d, Daniel Eibach a, Marcellus Fischer c, Ralf Matthias Hagen c,
Hagen Frickmann c,e

a Bernhard Nocht Institute for Tropical Medicine, Bernhard Nocht street 74, D-20359 Hamburg, Germany
b Institute for Clinical Chemistry, University Medical Centre, D-37075 Göttingen, Germany
c Department of Tropical Medicine at the Bernhard Nocht Institute, German Armed Forces Hospital of Hamburg, Bernhard Nocht street 74, D-20359
Hamburg, Germany
d Institute for Medical Microbiology, University Medical Centre, D-37075 Göttingen, Germany
e Institute for Microbiology, Virology and Hygiene, University Medicine Rostock, D-18057 Rostock, Germany

article info abstract

Article history: This review reports on laboratory diagnostic approaches for selected, highly pathogenic neglected
Received 6 March 2015 zoonotic diseases, i.e. anthrax, bovine tuberculosis, brucellosis, echinococcosis, leishmaniasis, rabies,
Received in revised form 3 August 2015 Taenia solium-associated diseases (neuro-/cysticercosis & taeniasis) and trypanosomiasis.
Accepted 4 September 2015
Available online 21 September 2015 Diagnostic options, including microscopy, culture, matrix-assisted laser-desorption-ionisation time-
of-flight mass spectrometry, molecular approaches and serology are introduced. These procedures are
Keywords: critically discussed regarding their diagnostic reliability and state of evaluation. For rare diseases reliable
Neglected zoonotic disease evaluation data are scarce due to the rarity of samples.
Neglected tropical disease If bio-safety level 3 is required for cultural growth, but such high standards of laboratory infrastruc-
Laboratory ture are not available, serological and molecular approaches from inactivated sample material might be
Diagnostic alternatives.
PCR Multiple subsequent testing using various test platforms in a stepwise approach may improve sensi-
Molecular diagnosis tivity and specificity.
Sensitivity Cheap and easy to use tests, usually called “rapid diagnostic tests” (RDTs) may impact disease control
Specificity measures, but should not preclude developing countries from state of the art diagnostics.
© 2015 The Authors. Published by Elsevier B.V. This is an open access article under the CC BY license

1. Introduction the World Health Organisation (WHO) Neglected Tropical Dis-
eases (NTDs) Initiative, the recognition of special requirements for
Zoonoses are diseases that are transmitted from animals (in research and control programme planning brought about by the
some definitions vertebrates) to humans. Most newly emerging zoonotic transmission pattern lead to the assembling of a list of
infections in humans are of zoonotic origin (Quammen, 2012) and neglected zoonotic diseases (NZDs) as a subset of the NTD list.
may be transmitted to humans via infected meat during prepara-
tion and ingestion or close contact to animals, e.g. when hunting, Some zoonotic diseases are difficult to handle due to their high
slaughtering or herding animals (Karesh et al., 2012). Following infectivity requiring special facilities (e.g. biosafety level 3, see
Table 1). Without claiming to be exhaustive, the WHO initially
∗ Corresponding author. Fax: +49 40 42818 512. listed 7 diseases as NZDs: Anthrax, bovine tuberculosis, brucel-
E-mail addresses: [email protected] (N.G. Schwarz), losis, cysticercosis and neurocysticercosis, cystic echinococcosis
or hydatid disease, rabies, zoonotic sleeping sickness or human
[email protected] (U. Loderstaedt), [email protected] African trypanosomiasis (HAT) (WHO, 2014). Different other dis-
(A. Hahn), [email protected] (R. Hinz), [email protected] (A.E. Zautner), eases are mentioned in different releases on NZDs. We decided
[email protected] (D. Eibach), marcellusfi[email protected] (M. Fischer), to focus our review on diagnosis of the 7 diseases from the initial
[email protected] (R.M. Hagen), [email protected] (H. Frickmann). list, additionally addressed Chagas disease, which like HAT is also
0001-706X/© 2015 The Authors. Published by Elsevier B.V. This is an open access article under the CC BY license (

N.G. Schwarz et al. / Acta Tropica 165 (2017) 40–65 41

Table 1 2.2. Disease
Attribution of described pathogens in alphabetical order to bio-safety levels accord-
ing to German National Guidelines, i.e. Technical guidelines for biological agents Cutaneous anthrax, comprising 95% of cases, is induced by
462, 464, and 466 (“Technische Richtlinien für biologische Arbeitsstoffe”). Bio-safety infected skin lesions and characterised by red skin efflorescence,
level 3** allows for handling in facilities with reduced structural safety infrastructure oedematous swelling, secretion filled vesicles and necrotic ulcers,
but with reliable personal protective equipment. the so-called “pustula malignans”. Systemic spread with lethal out-
come results in about 20% of non-treated patients. Due to fear of
Pathogen or pathogen group Bio-safety level bioterrorism, non-cutaneous forms are a main concern today. Gas-
trointestinal intake of insufficiently boiled infected meat may lead
Echinococcus spp. 3** to gut-necrosis. Pulmonary anthrax is extremely rare and results
Bacillus anthracis from inhalation of contaminated aerosols, e.g. in workers deal-
Brucella spp. 3 ing with animal skins. Case-fatality rate is nearly 100%, in spite
Leishmania spp. 3(1), and 2(2) of antibiotic therapy and intensive care medicine (Sweeney et al.,
Lyssaviruses 3**(3), 2(4), and 1(5) 2011). Vaccinations are not routinely recommended for travellers
Mycobacterium bovis ssp. bovis/caprae 3(6), and 3**(7) (Beyer, 2004) but restricted to veterinarians, outbreak situations or
Taenia solium specialised armed forces personnel.
Trypanosoma brucei gambiense/rhodesiense; 3
2.3. Diagnostic procedures
Trypanosoma cruzi 3**
3(8), and 3**(9)

(1)Brucella spp. other than (2), (2)B. ceti, B. microti, and B. pinnidedalis, (3)L. braziliensis,
L. donovani, L. guyanensis, L. infantum, and L. panamensis, (4)Leishmania spp. other
than (3) and (5), (5)L. enriettii, and L. tarentolae, (6)Mokola virus, and Lagos bat virus,
(7)lyssaviruses other than (6), (8)T. cruzi, (9) T. brucei gambiense/rhodesiense.

caused by trypanosomes, and furthermore added leishmaniasis on 2.3.1. Sampling and pre-analytic considerations
request by the Acta Tropica editor. Clinical samples comprise swabs from skin lesions and in case of

For diseases that can cause outbreaks, diagnosing the causative lung anthrax or gastrointestinal anthrax blood, sputum, and stool
agent can be essential for infection control, and for the fate of an samples. Post-mortal, heart blood, lung and spleen tissue should be
individual with a suspected infection the result of a diagnostic test considered. Samples should be transported cooled but not frozen in
may have immediate consequences: sterile vessels, blood additionally in inoculated blood culture media
(Rao et al., 2010) to BSL-3 facilities.
Diagnostic tests can provide incorrect results with serious con-
sequences for the individual or the community. Somebody who is 2.4. Microscopic approaches
not infected by the suspected agent but tests falsely positive may
suffer from treatment side effects, stigmatisation or be exposed to 2.4.1. Procedures
health facilities and acquire an infection after being admitted with If bacterial vitality is not required, fixation procedures should
other (really) infected individuals. On the other hand, false negative
results may hinder targeted treatment and infectious individuals be performed, e.g. heat fixation or fixation in 10% formalin solution
may infect other people. for light microscopy or fixation in 4% formaline / 0.5% glutaralde-
hyde in 0.5 M HEPES buffer, pH 7.2, for electron microscopy for 1
We provide an overview on laboratory diagnostic tests for 8 hour. Light microscopic approaches comprise Gram stain as well
highly pathogenic neglected zoonotic diseases (hp-NZDs): anthrax, as capsule staining according to Foth or M’Fadyean (M’Fadyean,
bovine tuberculosis, brucellosis, echinococcosis, leishmaniasis, 1903). Spores are stained with malachite-green / safranin or car-
rabies, Taenia solium-associated diseases (neuro-/cysticercosis & bol fuchsin / methylene blue (Hamouda et al., 2002). Microscopic
taeniasis) and trypanosomiasis. Many of the causative agents of mobility testing, using the “hanging drop” procedure (Tittsler and
these diseases require biosafety level 3 (BSL-3) facilities for safe Sandholzer, 1936), requires vital bacteria and BSL-3 facilities.
handling. We discuss strengths and weaknesses of different tests
for each disease, also keeping resource-limited settings in mind and For electron microscopy, fixed suspension is put on a carrier
impact of test results on patient care and management. Finally, we net and analyzed after a negative contrast procedure (Gelderblom,
reflect on laboratory diagnostic options in regions with no station- 1991).
ary BSL-3 facilities (Discussion I), i.e. the use of deployable BSL-3
labs or diagnostic approaches from inactivated sample material. 2.4.2. Reliability and critical interpretation of test results
Our closing remarks address general aspects on interpretation of Cylindrical, rod-shaped (1–1.25 × 3–5 ␮m) B. anthracis bacteria
diagnostic tests (Discussion II).
are usually arranged in chains. In vivo growth with demonstration
2. Anthrax of the capsule serves to discriminate B. anthracis from other spore-
forming bacteria. Elliptic, light-breaking spores are in the middle
2.1. Pathogen of the spore-forming cell and do not protrude from the cell. Vital
cells are immobile. Immunofluorescence procedures (Phillips et al.,
Bacillus anthracis is a spore-forming, world-wide distributed 1983) are not yet sufficiently standardised.
bacterium whose spores can long persist in soil. Virulent pathogens
harbour two plasmids, encoding the components of the synthesis 2.4.3. Value of microscopy in the diagnostic process
apparatus for the forming of a poly-gamma-D-glutamic acid cap- Microscopy can contribute to but not ensure the diagnosis of
sule on the plasmid pXO2 and the protective antigen (83.5 kDa),
the edema factor (an adenylylcyclase of 88.8 kDa), and the lethal anthrax.
factor (a metalloprotease of 90.2 kDa) on plasmid pXO1. Infections
of humans occur by direct contact to infected animals and alimen- 2.5. Cultural approaches with and without consecutive
tary intake of spores or vegetative cells (Mitscherlich and Marth, matrix-assisted laser desorption ionisation time-of-flight mass
1984; Sweeney et al., 2011). Detection of spores in environmental spectrometry
samples like powders or clothes is particularly challenging (Irenge
and Gala, 2012). 2.5.1. Procedures
Blood containing standard agars can be used in BSL-3 facilities

to demonstrate lacking haemolysis around B. anthracis colonies.
Pre-incubation at 70 ◦C for 30 minutes may inactivate vegeta-
tive, thermo-labile colonisers. Chromogenic agars like, e.g. “Cereus

42 N.G. Schwarz et al. / Acta Tropica 165 (2017) 40–65 Table 2 Probe 2 5 -CCT-GTA-TCC-ACC-CTC- ACT-CTT-CCA-TTT-TC-3 5 -TAT-TGT-TAT-CCT-GTT- ATG-CCA-TTT-GAG-ATT-TTT-3 5 -AAT-TCC-GTG-GTA-TTG- GAG-TTA-TTG-TTC-C-3 5 -GCG-CAA-GCT-TCT-GGT- GCT-AGC-3
Primer and probes of a well-characterised (Beyer et al., 2003) real-time multiplex PCR (förster resonance energy transfer (FRET) principle) for the identification of Bacillus anthracis. Reporters and quenchers can be chosen according
ident” agar (Heipha, Eppelheim, Germany), are in use (Rao et al., to requirements of the used PCR device. Probe 1 5 -TGC-GGT-AAC-ACT-TCA- CTC-CAG-TTC-GA-3 5 -GCT-AGT-TAT-GGT-ACA- GAG-TTT-GCG-AC-3 5 -CCA-TAA-CTA-GCA-TTT- GTG-CTT-TGA-AT-3 5 -CAA-GCA-AAC-GCA-CAA- TCA-GAA-GCT-AAG-3
2010; Tomaso et al., 2006).
Biochemical testing by API 50CH kits (bioMérieux, Nürtingen,
Germany) cannot discriminate B. anthracis from B. cereus but from Target Primer 1 5 -ACG-TAT-GGT-GTT-TCA- AGA-TTC-ATG-3
various other Bacillus spp. (Rao et al., 2010). Diagnostic approaches
comprise MALDI-TOF-MS, (Dybwad et al., 2013; Lasch et al., 2009)
gamma phage lysis of capsule-free, vegetative B. anthracis cells
(Schuch et al., 2002), capsule induction through addition of 0.7%
sodium bicarbonate to the agar with subsequent incubation at
12-15% CO2, capsule staining according to M’Fadyean (M’Fadyean,
1903), and proof of penicillin-sensitivity (Doganay and Aydin,

2.5.2. Reliability and critical interpretation of test results
Neither selective agars nor biochemical approaches can unam-

biguously discriminate between B. anthracis and closely related
Bacillus spp. (Rao et al., 2010). Phage-resistant and penicillin-
resistant B. anthracis strains have been infrequently described
(Abshire et al., 2005). MALDI-TOF-MS from culture material is sen-
sitive and specific and provides reliable results (Lasch et al., 2009).

2.5.3. Value of cultural diagnostic approaches in the diagnostic

Biochemical techniques require BSL-3 laboratory conditions and
usually do not allow for unambiguous results (Rao et al., 2010).
MALDI-TOF-MS from colonies may be used as a diagnostic stand-
alone technology (Lasch et al., 2009).

2.6. Molecular diagnostic approaches

2.6.1. Procedures
PCR-protocols for the identification of B. anthracis preferably tar-

get sequences on both virulence plasmids pXO1 and pXO2 and on
the bacterial chromosome (Agren et al., 2013; Antwerpen et al.,
2008; Beyer et al., 2003; Easterday et al., 2005; Ellerbrok et al.,
2002; Hoffmaster et al., 2002; Marston et al., 2006; Qi et al., 2001)
(Table 2). For PCR from agar colonies, simple boiling releases suffi-
cient DNA while more complex sample materials may require DNA
preparation kits (Beyer et al., 2003). Multi-locus variable-number
tandem repeat analysis (MLVA) or single nucleotide polymorphism
(SNP) can differentiate species (Lista et al., 2006; Stratilo et al.,
2006; Van Ert et al., 2007).

2.6.2. Reliability and critical interpretation of test results
Multiplexing various PCR targets (Beyer et al., 2003) can avoid

non-specific reactions (Helgason et al., 2000)(Hoffmaster et al.,
2002). Only few PCR targets (Agren et al., 2013) are not affected
by this problem. In some Bacillus anthracis strains one or even both
virulence plasmids are missing, potentially leading to false negative
results in multiplex assays (Turnbull et al., 1992). As such strains
lack their virulence genes, negative PCR results are acceptable if
technically adequate execution is confirmed by positive and nega-
tive controls. Positive PCR signals from primary patient materials,
however, should trigger isolation of the causative agent to ensure
specificity. In doubtful instances, diagnostic inoculation in animals
might be considered (Rao et al., 2010).

2.6.3. Value of molecular diagnostic approaches in the diagnostic

PCR can be applied for confirmation testing from colony mate-
rial, in case of false negative or non-interpretable cultural results
from spiked control samples and as a parallel diagnostic procedure
in case of non-selective enrichment (Hoffmaster et al., 2002).

N.G. Schwarz et al. / Acta Tropica 165 (2017) 40–65 43

2.7. Serological approaches Cultural growth requires BSL-3 conditions. Sample preparation
under a laminar flow workbench is advisable for samples from sus-
2.7.1. Procedures pected tuberculosis patients, as long as inactivation is not possible.
Apart from in-house ELISAs and Western blots for the detection If diagnostic procedures allow prior inactivation, it should be con-
sidered, e.g. by heating to 65 ◦C for at least 30 minutes (Best et al.,
of antibodies against protective antigen (PA), edema factor (EF), and 1990).
lethal factor (LF), a validated, commercially available anti-PA-IgG-
ELISA (Virion/Serion Ltd., Würzburg, Germany) can be purchased. 3.4. Microscopic approaches
Serological antigen detection kits (Burans et al., 1996) with suffi-
cient sensitivity and specificity are not available at present. 3.4.1. Procedures
Mycobacteria can be visualised as acid-fast rod-shaped bacteria
2.7.2. Reliability and critical interpretation of test results
Specific antibodies are not detectable in the critical early stages after Ziehl-Neelsen or Kinyoun staining (Fandinho et al., 1990) by
light microscopy at 1000× magnification. Auramine stained slides
of the disease. Therefore, serological testing to diagnose acute (Holani et al., 2014) are analysed under a fluorescence microscope
anthrax is futile. at 400× magnification.

2.7.3. Value of serological approaches in the diagnostic process 3.4.2. Reliability and critical interpretation of test results
Serological antibody assessment may help for retrospective Errors, deviation from protocols, and lacking experience with

assessments, differential diagnostic considerations or post- autofluorescent detritus after Auramine staining can lead to
vaccination assessments. false results. In case of doubtful Auramine results, decolorisation
and subsequent Ziehl-Neelsen staining allows for morphologi-
3. Bovine tuberculosis cal assessment of suspicious structures at 1000x magnification
(Fandinho et al., 1990). Bacteria other than mycobacteria are at
3.1. Pathogen least partially acid-fast, e.g. Nocardia spp. or Rhodococcus spp., so
confusion might occur (Jones et al., 2004).
The main causative agents of bovine tuberculosis are M. bovis
ssp. bovis and – to a lesser extent – M. bovis ssp. caprae (Ayele et al., 3.4.3. Value of microscopic approaches in the diagnostic process
2004; Cosivi et al., 1998). The demonstration of acid-fast rod-shaped bacteria by

Together with vaccination strain Bacillus Calmette-Guérin and Ziehl–Neelsen or Auramine staining is used for rapid assessment of
M. tuberculosis, M. africanum, M. microti, M. pinnipedii, and infectiousness, even in resource-limited settings (Sawadogo et al.,
M. canetti, they form the Mycobacterium tuberculosis complex. 2012), but fails to discriminate on species level.
Like Corynebacteriaceae, Nocardiaceae, and Tsukumurellaceae,
Mycobacterium spp. are Actinomycetales (Tortoli, 2003; Tortoli 3.5. Cultural approaches with and without consecutive
et al., 2006a; Tortoli et al., 2006b). matrix-assisted laser desorption ionisation time-of-flight mass
spectrometry (MALDI-TOF-MS)
Cattle represent the major reservoir of bovine tuberculosis,
while human-to-human transmission is rare (Evans et al., 2007). 3.5.1. Procedures
Reported incidences are 2/100,000 in WHO region Africa, 7/100,000 Cultural growth of mycobacteria at 37 ◦C and 30 ◦C (Pfyffer,
in WHO region America and 30/100,000 in WHO region Europe
(Muller et al., 2013). Diagnosis is mainly of epidemiologic inter- 2007; Richter, 2009) is possible from solid media and broths.
est. Diagnosis is done most of the time on clinical criteria and Growth on solid media, e.g. egg-based according to Löwenstein-
sometimes by direct microscopic examination with and without Jensen or agar-based according to Middlebrook, allows for
previous culture. macroscopic assessment of colonies (Muddaiah et al., 2013;
Sanders et al., 2004). Incubation should be performed for 8 weeks,
3.2. Disease if clinical suspicion persists for 12 weeks (Pfyffer, 2007; Richter,
2009). Increased CO2 concentration of 5-10% during the incubation
Bovine tuberculosis is clinically and radiologically indistin- of agar media facilitates mycobacterial growth and thus increases
guishable from M. tuberculosis-induced tuberculosis in humans. sensitivity (Isenberg, 2004; Pfyffer, 2007).
Granulomatous lesions manifest in a variety of organs, in partic-
ular infections of the gastrointestinal tract occur. Symptoms are Fluid culture media, e.g. Middlebrook 7H9, Middlebrook 7H12,
non-specific like chronic cough, fever, weight loss and sweating at BACTEC MGIT (BD, Heidelberg, Germany), or BacT/Alert MP
night (Cassidy, 2006). (bioMérieux, Marcy l’Étoile, France) (Pfyffer, 2007), are recom-
mended by WHO (WHO, 2007) even for low-income countries due
3.3. Diagnostic procedures to the shortened time-to-positivity by 1-2 weeks (Pfyffer, 2007).

3.3.1. Sampling and pre-analytic considerations Material from unsterile compartments should be decontami-
In case of suspected pulmonary tuberculosis, 3 putrid morning nated from accompanying bacterial flora prior to mycobacterial
culture by N-actetyl-l-cysteine (NALC) or by sodium dodecylsul-
sputa should be collected in sterile tubes. If fasting secretion from fate (SDS, unsuitable for cultural growth in several fluid media with
the stomach is collected via endoscopy, e.g. in children, it has to be automated detection), both in combination with NaOH (Pfyffer,
buffered prior to transport to maintain viability of mycobacteria. 2007).
Alternative sample materials, depending on the site of infection,
comprise urine, menstrual blood, lymphatic node tissue, bioptic Culture is followed by molecular methods or MALDI-TOF-MS
materials, sperm, bone marrow, blood culture material in case (Quinlan et al., 2014). If none is available, M. bovis spp. bovis and
of systemic spread, stool, and cerebrospinal fluid (CDC, 1999). M. bovis spp. caprae share the biochemical features negative niacin
Fluid samples should be enriched by centrifugation at 3300 × g. testing, negative nitrate reduction, low need for O2 on Lebek agar
Saliva, dried swabs, pooled sputum or urine, and sample storage (Meissner and Schroder, 1969), sensitivity against 1 mg/L furoic
or transport time >7 days are inacceptable for diagnostic purposes acid hydrazide, and sensitivity against cycloserine (Lumb et al.,
(Paramasivan et al., 1983). 2007). While M. bovis spp. bovis is resistant to pyrazinamide,

44 N.G. Schwarz et al. / Acta Tropica 165 (2017) 40–65

M. bovis spp. caprae is sensitive (Lumb et al., 2007). Subspecies- repetitive unit-variable number tandem repeat (MIRU-VNTR)
depending natural resistance against this first line tuberculostatic (Ramos et al., 2014).
drug (de Jong et al., 2005) makes reliable discrimination useful.
3.7. Serological approaches
3.5.2. Reliability and critical interpretation of test results
Culture in fluid media is the most sensitive method for the 3.7.1. Procedures
Antibody testing is irrelevant due to insufficient sensitivity and
detection of mycobacteria (CLSI, 2008; Dinnes et al., 2007) with a
detection limit of 10–100 bacteria (Pfyffer, 2007). MALDI-TOF-MS specificity (Steingart et al., 2011; Steingart et al., 2007).
recently showed 82.8% correct identifications (Quinlan et al., 2014). Methods of choice are Interferon gamma release assays (IGRA),
Identification of M. bovis spp. bovis by its pyrazinamide-resistance
is unreliable (Hlavsa et al., 2008). commercially available in an ELISPOT format (T-SPOT-TB, Oxford
Immunotec, Oxford, UK) and in an ELISA variant (QuantiFERON-TB
3.5.3. Value of cultural approaches in the diagnostic process Gold in tube, Cellestis Ltd., Carnegy, Australia). The antigens ESAT
In spite of its long incubation time, cultural growth under BSL- 6 und CFP 10, which are used for IGRA testing, can be found in M.
tuberculosis complex but not in the Bacillus Calmette Guerin vac-
3 conditions remains the diagnostic gold standard (Richter, 2009; cination strain (Ferrara et al., 2006; Mack et al., 2009; Pai et al.,
Rusch-Gerdes and Hillemann, 2008). 2008).

3.6. Molecular diagnostic approaches 3.7.2. Reliability and critical interpretation of test results
Regarding the clinical relevance of the results, T-SPOT and Quan-
3.6.1. Procedures
Numerous well validated commercial nucleic acid amplification tiferon were reported to be equivalent (Diel et al., 2010; Mack et al.,
2009). Infections with M. kansasii, M. marinum and M. szulgai were
(NAT) tests for the detection and identification of M. tuberculosis shown to be associated with reactive IGRA as well (Thijsen et al.,
complex are available, e.g. Amplified Mycobacterium Tuberculo- 2008). Specificity is considered to be 98% (Diel et al., 2011). Batch
sis Direct Test (AMTD) (Gen-Probe Inc., San Diego, CA, USA), BD specific specificity problems have been reported. Repeated tuber-
ProbeTec ET System (BD Diagnostics, Sparks, MD, USA), COBAS culin testing further reduces specificity of tuberculin testing due
TaqMan Mycobacterium tuberculosis MTB-Test (Roche Diagnostics, to booster effects (Menzies, 1999). Weakly positive results may not
Mannheim, Germany), Artus M. tuberculosis PCR Kit CE (Qiagen Ltd., always be reproducible (Felber and Graninger, 2013). False negative
Hilden, Germany), GenoType Mycobacteria Direct (GTMD) (HAIN IGRA results have been observed in about 10% of latently infected
Lifescience, Nehren, Germany), Loop-Mediated Isothermal Ampli- patients (Diel et al., 2009).
fication (LAMP) (Eiken Chemical Co. Ltd., Tokyo, Japan), RealArt M.
tuberculosis TM PCR Reagents (Abbott Laboratories, Abbott Park, A major disadvantage of IGRA is its unsuitability to discriminate
Illinois, USA), and GeneXpert System (Cepheid, Sunnyvale, CA, USA) active from latent tuberculosis (Whittaker et al., 2008).
(CLSI, 2008).
While anergy to tuberculin limits the diagnostic value of tuber-
Two variable regions of the 16S rRNA gene (Kirschner et al., culin testing in sarcoidosis patients with suspected tuberculosis,
1993; Springer et al., 1996) are best analysed for sequence-based the results of IGRA testing are not similarly affected (Gupta et al.,
species discrimination of mycobacteria (100% matching). For dis- 2011).
crimination within the M. tuberculosis complex, sequences of other
genes, like internal transcribed spacer (ITS), heat shock protein 3.7.3. Value of serological and IGRA approaches in the diagnostic
(hsp65), and rpoB gene have to be added (Adekambi et al., 2003). process
Alternative molecular discriminative approaches comprise the use
of gene probes like, e.g. AccuProbes (bioMérieux), strip hybridi- IGRA testing can detect latent tuberculosis prior to iatrogenic
sation assays like, e.g. GenoType CM/AS (HAIN LifeScience), and immunosuppression (Mack et al., 2009; Pai et al., 2008). Further
INNO-LiPA MYCOBACTERIA v2 (Innogenetics, Gent, Belgium) or the IGRA testing can identify early immunoconversions and should be
analysis of single nucleotide polymorphisms within the gyrase B performed in contact persons after potential transmission events
gene (gyrB) (Kasai et al., 2000; Niemann et al., 2000) and spoligo- to decide on preventive isoniazide therapy, although observed pro-
typing showing differences within the “direct repeat” (DR-) region gression rates in immunocompetent individuals are low (Nienhaus
(Kamerbeek et al., 1997). and Costa, 2013).

Next generation sequencing (NGS) makes differentiation of IGRA testing is not suitable for diagnosing acute tuberculosis
mycobacteria even on strain level possible (Outhred et al., 2014). due to lacking sensitivity and specificity (Diel et al., 2009; Sester
et al., 2011).
3.6.2. Reliability and critical interpretation of test results
A meta-analysis of commercially available NAT systems showed 4. Brucellosis

an average sensitivity of 85% for microscopically positive and neg- 4.1. Pathogen
ative samples, while the average specificity was 97% (Ling et al.,
2008). As applicable for all diagnostic procedures, positive and The causative agents of brucellosis are Brucella spp., Gram-
negative predictive values depend on prevalence. Accordingly, the negative, rod-shaped bacteria with close phylogenetic relationship
positive predictive of test results is low in non-selected groups in to Bartonella spp., Ochrobacterium spp., and Agrobacterium spp.
low-endemicity settings. (Moreno et al., 1990). The genus Brucella comprises nine closely
related species with sequence homology >90%, from which three
3.6.3. Value of molecular diagnostic approaches in the diagnostic are further differentiated in biovars (Verger et al., 1985) (Table 3).
4.2. Disease
The GeneXpert system is broadly used for tuberculosis screen-
ing in Africa (Scott et al., 2014; Wekesa et al., 2014), suggesting Brucellosis is worldwide endemic (Godfroid et al., 2005) and
an increasing importance of molecular procedures for the routine with 500.000 human cases per year a frequent (Pappas et al., 2006)
diagnosis of mycobacteria worldwide. NGS could replace older typ- zoonotic disease which, is particularly common in the Mediter-
ing approaches like spoligotyping or mycobacterial interspersed ranean, on the Arabic peninsula, in the Middle East, in Africa

N.G. Schwarz et al. / Acta Tropica 165 (2017) 40–65 45

Table 3
Brucella spp. and their typical hosts (Alaeddini, 2012; Cloeckaert et al., 2001; Foster et al., 2007; Jahans et al., 1997; Scholz et al., 2008; Verger et al., 1985)

Species Biovars Predominant hosts
Quantitatively most important Brucella spp. isolated from infected humans
B. melitensis 1-3 Goats, sheep
B. abortus 1-6, 9 Cattle, other Bovidae
B. suis 1-3 Pigs
4 Reindeer
B. canis 5 Rodents
– Dogs

Brucella spp. which are only sporadically isolated from infected humans – Sheep
B. ovis – Desert rats
B. neotomae – Whales
B. ceti – Crawl, sea lions, walruses
B. pinnipedalis – Common field mouse (Microtus arvalis)
B. microti

Table 4
Primers of the AMOS-PCR targeting the non-coding insertion sequence IS711 of B. abortus (biovars 1, 2, 4), B. melitensis (biovars 1-3), B. ovis, and B. suis (biovar 1).

Primer type Specificity to IS711 of Fragment length Primer sequence
B. abortus, biovars 1, 2, 4 498 bp 5 -GAC-GAA-CGG-AAT-TTT-TCC-AAT-CCC-3
Reverse B. melitensis 731 bp 5 -AAA-TCG-CGT-CCT-TGC-TGG-TCT-GA-3
B. ovis 976 bp 5 -CGG-GTT-CTG-GCA-CCA-TCG-TCG-3
B. suis, biovar 1 285 bp 5 -GCG-CGG-TTT-TCT-GAA-GGT-TCA-GG-3
Above listed species/biovars Variable/see above! 5 -TGC-CGA-TCA-CTT-AAG-GGC-CTT-CAT-3

and in Middle and South America (Young, 1995b). Sources of be shipped cooled or frozen (Moreno and Moriyon, 2011); (Young,
infection comprise the consumption of unpasteurised milk prod- 1995a).
ucts like cheese from sheep or goats and raw meat, contact with
infected animals or their excrements, aerosols during meat pro- 4.4. Microscopic approaches
cessing, veterinary-obstetrics and processing of clinical samples in
the laboratory. Human-to-human transmission, e.g. due to sexual 4.4.1. Procedures
intercourse or transplantation, is rare. Due to the low infectious If any, Gram-staining (Claus, 1992) might be considered.
dose of only 10 colony forming units, brucellosis is the most fre-
quent laboratory infection worldwide (Young, 1995a). After an 4.4.2. Reliability and critical interpretation of test results
incubation period between 14 days and several months, acute bru- Brucella spp. are visible as small (400 × 800 nm), Gram-negative,
cellosis manifests as febrile systemic disease with unspecific flulike
symptoms, undulating fever, fatigue, tiredness, headache and body coccoid to rod-shaped bacteria that are poorly stainable according
aches. Chronicity is frequent with night sweat, weight loss, hep- to Gram (Meyer and Shaw, 1920).
atosplenomegaly, lymphadenitis as well as organ manifestations
like spondylitis, sakroileitis, neurobrucellosis and endocarditis. 4.4.3. Value of microscopy in the diagnostic process
Case fatality rate is about 2-5% with 80% of deaths due to endocardi- Microscopy is no method of choice for the diagnosis of brucel-
tis with heart insufficiency (Young, 1995a). In spite of long-lasting
combined antibiotic therapy, primary therapy failure and recur- losis.
rence are frequent due to persistence and replication of bacteria
in phagosomes of macrophages (Kohler et al., 2002; Kohler et al., 4.5. Cultural approaches with and without subsequent
2003). Previous vaccine trials for humans failed (Live, 1958; matrix-assisted laser desorption ionisation time-of-flight mass
Vershilova, 1961). Postexposure prophylaxis with 100 mg doxycy- spectrometry (MALDI-TOF-MS)
cline twice daily and 600 mg rifampicin (rifampin) four times daily
for 3 weeks is poorly evaluated, but may be considered in case of 4.5.1. Procedures
relevant exposition (Yagupsky and Baron, 2005). Brucella spp. show optimal cultural growth at 35–37 ◦C and

4.3. Diagnostic procedures 5–10% CO2. Tiny colonies are visible not earlier than after 48-
72 hours of incubation on blood-containing agars. Co-occurrence
4.3.1. Sampling and pre-analytic considerations of young smooth (S) and older rough (R) colony variants may
If brucellosis is suspected, blood cultures should be taken resemble a mixed culture. Centrifugation-lysis techniques and
mechanised blood-culture systems like BACTEC (Becton Dickin-
repeatedly. Isolation of the pathogen from bone marrow is son, East Rutherford, New Jersey, USA) or BAC/ALERT (bioMérieux,
more often successful, sometimes also from tissue samples, cere- Marcy-l’Étoile, France) can increase isolation rates. Incubation time
brospinal fluid, synovial fluid, abscess material, spleen and liver should be prolonged to 3 weeks (Kolman et al., 1991; Ruiz et al.,
tissue or from consumables like milk, cheese or raw meat. Sample 1997; Yagupsky, 1994). For isolation from primary unsterile sites,
transport has to be rapid and cultural growth should start within in-house selective agars containing amphotericin B, bacitracin,
1-2 hours after sample taking. For longer transport times, clinical cycloheximide/natamycin, D-cycloserine, nalixidic acid, polymyxin
samples should be cooled to 2-8 ◦C, tissue samples have to stay B, and vancomycin are suitable for most strains (Alon et al., 1988).
moist during transport (Moreno and Moriyon, 2011). Samples for
cultural growth under BSL-3 conditions should be taken prior to Brucella spp. are catalase, and, with few exemptions, also urease
the onset of antibiotic therapy (Gotuzzo et al., 1986). Serum should and oxidase positive. Biochemical reactions include need for CO2
for growth, H2S-production, splitting of urea, sensitivity against the
stains fuchsine and thionine, sensitivity against phages, and agglu-
tination with polyvalent as well as specific antisera (Al Dahouk

46 N.G. Schwarz et al. / Acta Tropica 165 (2017) 40–65

et al., 2003a; Alon et al., 1988; Jahans et al., 1997; Scholz et al., antibodies against the s(smooth)-LPS of B. abortus, B. melitensis,
2008). Older colonies (R-variants) may autoagglutinate. and B. suis with Brucella-LPS in phenol-inactivated bacteria of
the reference strain B. abortus 1119-3 USDA in tube-agglutinaton
Identification of Brucella spp. by MALDI-TOF-MS from colony assays and in faster micro-agglutination assays (Al Dahouk et al.,
material is possible (Karger et al., 2013). 2003a). Next to SAT, complement binding reaction (CBR) (Diaz and
Moriyon, 1989) and ELISA techniques (Caravano et al., 1987) are
4.5.2. Reliability and critical interpretation of test results established.
Sensitivity of cultural growth is about 50-70% in case of acute
4.7.2. Reliability and critical interpretation of test results
brucellosis but time-to-positivity is long. Automated biochemical High SAT-titres (≥1:160 in non-endemic settings) or a titre
identification systems like API 20NE (bioMérieux) frequently fail to
identify Brucella spp. (Barham et al., 1993). MALDI-TOF-MS is the increase by factor 4 are demanded for the diagnosis of active bru-
method of choice for identification from colony material (Karger cellosis (Lucero and Bolpe, 1998). Low antibody titres close to the
et al., 2013). detection limit are typical in early stages and for chronic courses
of the disease (Ariza et al., 1992) and serology can fail (Naha et al.,
4.5.3. Value of cultural isolation in the diagnostic process 2012). In contrast, healthy farmers, butchers or veterinarians can
Cultural isolation of Brucella spp. is considered as the diagnostic show increased anti-Brucella-IgG titres in case of repeated exposi-
tion to the antigen in absence of an active infection (Karimi et al.,
gold standard. 2003).

4.6. Molecular diagnostic approaches IgM-antibodies against LPS dominate the first week after infec-
tion, followed by an IgG-titre increase in the second week and a
4.6.1. Procedures peak of both 1 month after infection. Although IgM-titres slowly
Genus-specific identification of Brucella-DNA has been decrease with antibiotic therapy, IgM may persist for more than a
year. Newly observed increases of IgM-titre indicate therapy fail-
described for various targets, e.g. omp2, omp2b, bcsp31 ure or recurrence. Increased IgG-titres can persist for years in spite
(e.g. targeting a 223-bp-long amplicon) (primers B4: 5 - of successful therapy (Smits et al., 1999).
CTT-TCA-GGT-CTG-3 ), IS711 (IS6501), 16S-23S rRNA spacer Of note, SAT testing based on B. abortus 1119-3 USDA antigens
region and the 16S rRNA gene (Al Dahouk et al., 2003b; Bricker, does not indicate antibodies against R-variant-LPS of B. canis (Al
2002), also as real-time approaches (Al Dahouk et al., 2004; Dahouk et al., 2003a). In case of very high Brucella titres, inhibition
Probert et al., 2004; Redkar et al., 2001). The so-called AMOS-PCR of agglutination (high-dose hook effect) in SAT has to be overcome
can differentiate frequent Brucella spp. and biovars (Bricker and by serial dilutions (Lucero and Bolpe, 1998).
Halling, 1994) targeting the non-coding insertion sequence IS711
of B. abortus (biovars 1, 2, 4), B. melitensis (biovars 1-3), B. ovis, and CBR for IgG1 is used for confirmation testing due to its higher
B. suis (biovar 1) (Table 4), next to other procedures (Garcia-Yoldi specificity and is even more reliable than SAT during the incuba-
et al., 2006). tion period of infection and in case of chronic brucellosis (Diaz and
Moriyon, 1989). ELISA systems facilitate differentiated adjustment
Fluorescence in situ hybridisation (FISH) can identify Brucella to a specific state of the disease by identifying very small differ-
spp. from blood and agar cultures (Wellinghausen et al., 2006). ences of concentrations of different antibody classes (Caravano
et al., 1987).
Restriction fragment length polymorphism (RFLP) after PCR
allows for further differentiation on species or strain level (Al Decreasing the pH-value can increase specificity of SAT (Lucero
Dahouk et al., 2005; Vizcaino et al., 2000), variable-number tandem and Bolpe, 1998). For whole-cell-antigen tests, cross-reacting
repeat (VNTR), RNA mismatch cleavage (Bricker, 1999) and Fourier antibodies against Afipia clevelandensis, E. coli O:157, Francisella
transform infrared spectroscopy (Miguel Gomez et al., 2003) on tularensis, Salmonella spp., Stenotrophomonas maltophilia, Vibrio
strain level (Bricker et al., 2003; Le Fleche et al., 2006). cholerae, and Yersinia enterocolitica O:9 have to be considered
(Drancourt et al., 1997; Lucero and Bolpe, 1998).
4.6.2. Reliability and critical interpretation of test results
Brucella spp. DNA can be detected in clinical samples earliest 10 4.7.3. Value of serological approaches in the diagnostic process
If cultural or molecular proof of Brucella spp. fails, high titres or
days after infection with a higher sensitivity than blood culture and
a theoretical detection limit of as low as 10 genome equivalents (Al titre-increases by factor 4 indicate acute infection in case of suspi-
Dahouk et al., 2003b; Bricker, 2002). Sequences of amplicons from cious symptoms (Centers for Disease and Prevention, 1994; Young,
traditional PCR approaches should be confirmed. Rapid FISH-based 1995a). Brucella serology also helps monitoring the course of the
genus-identification has not been standardised beyond a proof- disease (Smits et al., 1999).
of-principle evaluation (Wellinghausen et al., 2006). Sequencing
of the 16S rRNA gene is unsuitable for differentiation below the 5. Echinococcosis
genus level due to perfect sequence conservation among Brucella
spp. (Gee et al., 2004). 5.1. Pathogen

4.6.3. Value of PCR methods in the diagnostic process Tapeworms (cestodes) of the genus Echinococcus comprise six
PCR is rapid with better sensitivity in comparison to culture but species of which E. granulosus and E. multilocularis are of major med-
ical importance. E. granulosus occurs worldwide, E. multilocularis
cannot replace culture as diagnostic gold standard allowing antimi- mainly in the northern hemisphere (Eckert and Deplazes, 2004;
crobial sensitivity testing (Navarro et al., 2004; Queipo-Ortuno Jenkins et al., 2005; Romig et al., 2006).
et al., 2005; Redkar et al., 2001).
5.2. Disease
4.7. Serological approaches
Adult or larval stages of E. granulosus and E. multilocularis cause
4.7.1. Procedures cystic (CE) and alveolar echinococcosis (AE), respectively (Jiang
Many approaches rely on the binding of Brucella-LPS et al., 2005; Khuroo, 2002; McManus et al., 2003; Raether and

(lipopolysaccharide)-antibodies to whole-cell-antigens. Serum
agglutination tests (SAT) demonstrate the binding of agglutinative

N.G. Schwarz et al. / Acta Tropica 165 (2017) 40–65 47

Hanel, 2003; Thompson et al., 1995). The definite hosts of E. gran- should be used (Pawłowski et al., 2001). Neurocysticercosis needs
ulosus are carnivores, which shed eggs and proglottides in their to be ruled out by specific ELISA or immunoblotting (Ito et al., 2003).
faeces. Ingestion by humans or ungulate intermediate hosts leads
to the development of hydrated cysts, predominantly in liver (70%) 5.5.2. Reliability and critical interpretation of test results
and lung (20%) (Zhang and McManus, 2006). To reduce the disease Em2, Em10 and Em18 antigens are used for the detection of
burden in developing countries control programmes have to incor-
porate veterinary transmission chains in carnivores and ungulates. E. multilocularis antibodies. Em2-ELISA had sensitivities ranging
between 77 and 92% (Zhang and McManus, 2006). Em2-ELISA
E. multilocularis transmission is maintained in a sylvatic cycle detected antibodies do not always correlate with disease activ-
involving wild carnivores, small mammals, but also domestic dogs ity, since some patients with inactive lesions remain seropositive
as definite hosts (Craig et al., 2000; Kern et al., 2004; Wang et al., (Rausch et al., 1987). The Em2plus ELISA, a combination of Em2-
2006). Human AE is a chronic, often fatal disease with infiltrative antigen and the recombinant protein II/3–10, increased sensitivity
growing liver cysts, leading to destruction of liver parenchyma, bile for detection of AE to 97% (Gottstein et al., 1993). An ELISA detect-
ducts and blood vessels (Zhang and McManus, 2006). AE infections ing antibodies against Em10 antigen revealed a sensitivity of 93.2%
become only symptomatic in advanced stage, which often delays and a specificity of 96.8% (Helbig et al., 1993).
the diagnosis for many years (Moro et al., 2008)
The 18 kDa antigen (Em18) proved to be highly species-specific
5.3. Diagnostic procedures (96.8%) and sensitive (97%), and correlated with the presence of
active lesions whereas a loss or decrease correlated with inac-
5.3.1. Sampling and pre-analytic considerations tive infection, resection or long-term antiparasitic chemotherapy.
An interdisciplinary team of clinicians, radiologists and microbi- In patients with progressive AE, Western blot bands at 18 kDa
increased (Ito et al., 1995; Ito et al., 1993; Tappe et al., 2008). Em18
ologists needs to differentiate echinococcal cysts from benign cysts, is a good candidate for differentiation between CE and AE, in regions
cavitary tuberculosis, mycoses, abscesses, and benign or malig- where both infections are prevalent. However, Em18-ELISA and
nant neoplasms. Radiologic imaging is used as a screening tool Em18-Western blots revealed minor cross-reactivity with neuro-
to diagnose and evaluate liver lesions (Group, 2003; Junghanss cysticercosis and schistosomiasis (Ito et al., 2002).
et al., 2008), which are further confirmed by antibody assays (Moro
and Schantz, 2009). Direct detection of the pathogen requires Serological antibody detection for CE is less reliable than for
ultrasound-guided fine-needle aspiration cytology (FNAC) of cyst AE (McManus et al., 2003). E. granulosus hydatid cyst fluid (EgHF)
fluid or biopsies of liver. is used for antibody detection (Carmena et al., 2006), with sen-
sitivities ranging from 75 to 95% (Zhang et al., 2003). The test
5.4. Microscopic approaches specificity (50-82%) (Moro et al., 2005) is reduced due to frequent
cross-reactions in patients with tumours, liver cirrhosis, nematode,
5.4.1. Procedures trematode and other cestode infections (Craig et al., 2007; Dottorini
Smears of aspirated fluid from suspected E. granulosus cysts can et al., 1985; Eckert and Deplazes, 2004; Ito et al., 1995; Ito et al.,
1993; Khuroo, 2002; Maddison et al., 1989; Ortona et al., 2000;
be examined microscopically for acid-fast protoscolex hooklets as Poretti et al., 1999; Shepherd and McManus, 1987; Siracusano et al.,
described before (Handa et al., 2001; Hira et al., 1988). In pulmonary 1991; Wen and Craig, 1994; Zhang and McManus, 2006). Antibod-
CE infections, protoscolices might be found in sputum or bronchial ies remain detectable following surgical removal or effective drug
washings (Moro and Schantz, 2009). treatment of cysts. Other patients with CE do not demonstrate a
detectable immune response (Dottorini et al., 1985; Force et al.,
5.4.2. Reliability and critical interpretation of test results 1992; Lorenzo et al., 2005). Serodiagnostic performance seems to
Microscopy of cystic fluid proves infection and cyst viability depend on factors, such as cyst location, cyst stage, and cyst size
(Brunetti et al., 2011). .
(Brunetti and Junghanss, 2009). However, some cysts are sterile
(acephalocysts) as their brood capsules are absent or not devel- Use of purified antigens improves specificity of serological
oped. A negative microscopy result can therefore not rule out CE assays but may subvert sensitivity (Gadea et al., 2000; Zhang et al.,
infection. 2003). Therefore the crude EgHF-ELISA has been recommended for
genus specific screening (Pawłowski et al., 2001). Subsequently,
5.4.3. Value of microscopic approaches in the diagnostic process false positive results can be detected with more specific confirma-
FNAC, followed by microscopy of aspirated fluid can confirm the tory tests. Antibodies against antigen B (AgB) or antigen 5 (Ag5)
have shown a specificity between 77 and 91%, but commonly cross-
diagnosis (Hira et al., 1988; Pawłowski et al., 2001). react with E. multilocularis. In regions, where CE and AE co-circulate,
highly specific E. multilocularis antigens are required (Craig et al.,
5.5. Serological approaches 2000; Gottstein et al., 1993; Helbig et al., 1993; Ito et al., 1999; Ito
et al., 1993; Ito et al., 2002; Jiang et al., 2001; Liance et al., 2000;
5.5.1. Procedures Sarciron et al., 1997; Siles-Lucas and Gottstein, 2001). The distinc-
A range of insensitive and non-specific tests has been replaced tion between active or progressive and cured CE patients is not yet
standardised (Zhang and McManus, 2006).
by indirect haemagglutination assays, ELISAs and immunoblots
(Auer et al., 2009; Zhang et al., 2003). Serological screening based The Echinococcus Western Blot IgG kit (Echinococcus West-
upon crude antigens of E. granulosus or E. multilocularis, can lead ern Blot IgG; LDBIO Diagnostics, Lyon, France) differentiated well
to non-specific reactions, due to high genetic homology of E. mul- between AE and CE patients (Ito et al., 1999; Jiang et al., 2001).
tilocularis and E. granulosus antigens (Ito et al., 2003; Jiang et al.,
2001). After screening for CE in endemic areas using crude antigens 5.5.3. Value of serological approaches in the diagnostic process
(hydatid cyst fluid) subsequent confirmatory testing with specific Despite standardisation and cross-reactivity problems, the
E. granulosus antigens (e.g. antigen B) is required. For AE diagnos-
tics in endemic areas, both primary screening and confirmation combination of screening and confirmatory serological tests in con-
should be performed with Em2-, Em2plus-, Em10-, Em18-ELISA or junction with radiologic imaging represents the method of choice
Em18-Western blots. In areas, where both AE and CE are endemic, a for the diagnosis of Echinococcus spp. infections.
combination of very sensitive genus-specific screening and species-
specific confirmation tests (e.g. Echinococcus IgG Western Blot)

48 N.G. Schwarz et al. / Acta Tropica 165 (2017) 40–65

5.6. Molecular diagnostic approaches most active should be chosen for skin-scrapings and biopsy (CDC,
PCR is mainly used for seronegative patients, when imaging is
suggestive for AE or CE. E. multilocularis and E. granulosus specific Skin for biopsy is prepared by injection of local anaesthetics (e.g.
sequences can be amplified from tissue biopsies and FNAC (Brunetti 1% lidocaine preferentially with epinephrine 1:100,000) through
and Junghanss, 2009; Diebold-Berger et al., 1997; Georges et al., intact skin cleansed with 70% alcohol into the dermis underlying
2004; Kern et al., 1995). PCR methods are not able to assess viability the area that will be sampled (Dar and Khurshid, 2005).
and a negative PCR cannot rule out disease (Brunetti et al., 2010).
Skin scraping with a blade (Gazozai et al., 2010) from the mar-
6. Leishmaniasis gins of the ulcer is applied as a smear on multiple slides for
microscopic evaluation thus reaching sensitivities up to 80%.
6.1. Pathogen
Sterile full-thickness, punch-biopsy specimens (4 or 6 mm in
Leishmaniasis is a vector-borne disease caused by obligate intra- diameter) should be taken at the active border of the lesion and
cellular protozoa of the genus Leishmania and spread by the bite of divided into 3 portions, one for impression smears, one for cultural
infected female phlebotomine sandflies. Species-depending tissue growth (leishmania, bacteria, mycobacteria, and fungi) and one for
specificities cause differing clinical manifestations of the various histologic examination of tissue sections (fixed in 10% formalin;
forms of leishmaniasis (Boecken et al., 2011). embedded in paraffin) stained with H&E, Giemsa, and other special
stains to exclude mycobacterial, fungal, and other infectious aeti-
Human infection is caused by 21 of 30 species that infect mam- ologies. The portion placed in leishmania culture medium can also
mals. These include the L. donovani complex with 2 species (L. be used for PCR (CDC, 2014).
donovani, L. infantum) [also known as L. chagasi in the New World];
the L. mexicana complex with 3 main species (L. mexicana, L. ama- For tissue impression smears (touch preparations), bioptic spec-
zonensis, and L. venezuelensis); L. tropica; L. major; L. aethiopica; imens are carefully grasped with forceps and excess blood removed.
and the subgenus Viannia with 4 main species (L. (V.) brazilien- The specimen is gently pressed - with a rolling or circular motion -
sis, L. (V.) guyanensis, L. (V.) panamensis, and L. (V.) peruviana). The onto several glass slides for microscopy, air-dried, fixed and stained
different species are morphologically indistinguishable, but can be according to Giemsa (CDC, 2014).
differentiated by isoenzyme analysis, molecular methods, or with
monoclonal antibodies (Cupolillo et al., 2000; Fraga et al., 2010; Visceral Leishmaniasis (kalar azar). Amastigote parasites
Garcia et al., 2005; Schonian et al., 2010). can be demonstrated in spleen or bone marrow, lymph nodes, liver
biopsy, aspirates or tissue specimens or even in the buffy coat of
6.2. Disease peripheral blood. Amastigote density can be logarithmically scored
from 0 (no parasite per 1000× magnified oil immersion fields) to 6
The disease can present as cutaneous, mucocutaneous, or vis- (>100 parasites per field). For imprint cytology from spleen, liver,
ceral leishmaniasis (Jacobs et al., 2005). Endemic areas are found or lymph node, flat tissue cuts are pressed on microscopic slides,
in Mexico, Central America, and South America - from northern fixed with absolute alcohol, and stained according to Giemsa. Hos-
Argentina to Texas (not in Uruguay, Chile, or Canada) -, south- pital assessment of tissue specimens comprises Giemsa or standard
ern Europe, Asia (not Southeast Asia), but especially in the Middle hematoxylin-eosin staining (Arechavala et al., 2010).
East, and in Africa particularly in East and North Africa (Haouas
et al., 2005; Jamjoom et al., 2004; Magill, 2005; Mahdi et al., 2005; 6.4. Microscopical approaches
Mendonca et al., 2004; Silveira et al., 2004).
6.4.1. Procedures
The cutaneous form presents with skin ulcers, the mucocuta- Amastigotes can be visualised by both Giemsa and hematoxylin-
neous form with purulent ulcerations in the nasal area and parts
of the oral cavity (Antinori et al., 2005). Visceral leishmaniasis eosin (HE) staining (Arechavala et al., 2010), differential staining
affects bone marrow, liver and spleen and presents with undulating of DNA and RNA with acridine orange (Arechavala et al., 2010;
fever, hepatosplenomegaly and pancytopenia. Untreated visceral Gazozai et al., 2010; Mbati et al., 1995; Meymandi et al., 2010;
leishmaniasis leads to death (Mondal et al., 2010), in particular in Perez-Cordero et al., 2011), and immunohistochemical staining
co-existence with HIV (Harms and Feldmeier, 2005). Leishmania (Ismail et al., 1997). It is important to see the nucleus and
are known co-factors in the pathogenesis of HIV-1 infection, medi- the rod-shaped kinetoplast, a mitochondrial structure contain-
ate HIV-1 activation by cytokine secretion and cellular-signalling, ing extra-nuclear DNA, to diagnose leishmaniasis. The kinetoplast
and up-regulate HIV-1 replication, both in monocytoid and lym- differentiates Leishmania from other small organisms such as Histo-
phoid cells in vitro and in co- infected individuals (Olivier et al., plasma (Arechavala et al., 2010; Gazozai et al., 2010; Mbati et al.,
2003). Over 90% of visceral leishmaniasis cases occur in six 1995; Meymandi et al., 2010).
countries: Bangladesh, Brazil, Ethiopia, India, South Sudan, and
Sudan. The majority of cutaneous leishmaniasis cases occur in 6.4.2. Reliability and critical interpretation of test results
Afghanistan, Algeria, Brazil, Colombia, Iran, Pakistan, Peru, Saudi Sensitivity of microscopy from bone marrow smears is about 60
Arabia, and Syria. Almost 90% of mucocutaneous leishmaniasis
cases are observed in Bolivia, Brazil, and Peru (Haouas et al., 2005; to 85%, from spleen aspirates more than 95% (Sundar and Rai, 2002).
Jamjoom et al., 2004; Magill, 2005; Mahdi et al., 2005; Mendonca Immunohistochemistry can increase detection rate of leishmania in
et al., 2004; Silveira et al., 2004). tissue from 20% after H&E staining to 88% (Ismail et al., 1997).

6.4.3. Value of microscopic methods in the diagnostic process
Microscopy is still the most commonly performed procedure for

initial screening in many laboratories.

6.3. Diagnostic procedures 6.5. Cultural approaches

6.3.1. Sampling and pre-analytic considerations 6.5.1. Procedures Cutaneous and mucocutaneous Leishmaniasis. Several tech- Leishmania strains can be maintained as promastigotes in artifi-
niques should be applied simultaneously and several specimens
per technique should be taken. Lesions that appear youngest and cial culture medium, e.g. monophasic (Schneider’s insect medium,
M199, or Grace’s medium) or biphasic media (Novy-McNeal Nicolle

N.G. Schwarz et al. / Acta Tropica 165 (2017) 40–65 49

medium and Tobies medium) (Sundar et al., 2001). Culture tubes need, even decades after the sample has been taken (Volpini et al.,
are inoculated with 1 to 2 drops of bone marrow or spleen aspirate 2006).
and incubated at 22-28 ◦C. Aseptically collected blood (1 to 2 ml) is
diluted with 10 ml of citrated saline; after centrifugation the cellu- 6.6.3. Value of molecular diagnostic approaches in the diagnostic
lar deposit obtained is inoculated in culture media. Contamination process
of the culture media by bacteria or yeast species or other fungi
can be avoided by sterile techniques and by the addition of antibi- As microscopic skills get lost in non-endemic settings, molecular
otics and antifungal drugs (Schur and Jacobson, 1988; Sundar et al., procedures become more important. For new world leishmania-
2001). sis they provide tools for species diagnosis to differentiate species
responsible for cutaneous from species responsible for mucocuta-
In laboratory animals (such as hamsters, mice or guinea pigs), neous forms (Graca et al., 2012).
the parasite can be inoculated via mucous membranes or the
intraperitoneal and intrasplenic routes with infected specimen, fol- Modern typing methods such as multilocus enzyme typ-
lowed by the microscopical demonstration in bioptic specimens ing (MLEE), multilocus sequence typing (MLST), and multilocus
(Mbati et al., 1995). microsatellite typing (MLMT) (Schonian et al., 2011) widely
replaced laborious and time-consuming ISH for typing.
After cultural isolation, parasites can be characterised to the
species-complex and sometimes to the species level using isoen-
zyme analysis (Jamjoom et al., 2004).

6.5.2. Value of cultural approaches in the diagnostic process 6.7. Serological approaches
Cultures are useful for obtaining a sufficient number of organ-
6.7.1. Procedures
isms to use as an antigen for immunologic diagnosis and typing, for Traditional methods for anti-Leishmania-antibody detection
obtaining parasites to be used to inoculate susceptible experimen-
tal animals, for in vitro screening of drug sensitivity, and finally as include gel diffusion, complement fixation testing, indirect
an additional possibility for diagnosing the infection. haemagglutination testing, immunofluorescence assays, and
counter-current immunoelectrophoresis. Specificity depends upon
6.6. Molecular diagnostic approaches the antigen or epitope used (Saha et al., 2005) with recombinant
antigens improving the specificity (Braz et al., 2002).
6.6.1. Procedures
Leishmania spp. diagnosis by various nucleic acid amplifica- Sensitivity and specificity of ELISA in diagnosing visceral leish-
maniasis could be increased by using soluble antigens derived from
tion techniques (NAT) comprise DNA hybridisation, real-time PCR promastigotes cultivated in a protein-free medium (Rajasekariah
with and without quantification, high-resolution melt analysis, et al., 2001). Specific antibodies can further be detected by
PCR-RFLP (restriction fragment length polymorphism analysis), immunoblotting (Brito et al., 2000; Kaur and Kaur, 2013). To ease
sequencing, and DNA microarrays (Antinori et al., 2009; Castilho sample acquisition in a non-invasive way, ELISAs from urine have
et al., 2008; Disch et al., 2005; el Tai et al., 2000; Foulet et al., 2007; been introduced as well (Islam et al., 2008).
Meymandi et al., 2010; Paiva et al., 2004; Rodriguez et al., 1994;
Roelfsema et al., 2011; Salotra et al., 2001; Talmi-Frank et al., 2010; Detection of visceral leishmaniasis antigens in urine by direct
Tavares et al., 2003; van der Meide et al., 2008; Volpini et al., 2006; agglutination (Attar et al., 2001) and dipstick testing (Bern et al.,
Zelazny et al., 2005). A frequent target for molecular approaches, 2000) have shown to be affordable and easy-to-apply rapid tests
which also allows for the discrimination of New World leishmania for resource-limited countries in several studies (Chappuis et al.,
is hsp70 (da Silva et al., 2010; Fraga et al., 2010; Fraga et al., 2012; 2006a; Chappuis et al., 2006b; Rijal et al., 2004).
Graca et al., 2012; Montalvo et al., 2012; Requena et al., 2012).
6.7.2. Reliability and critical interpretation of test results
Molecular in situ hybridisation (ISH) using kinetoplast DNA Antibody detection is useful for visceral leishmaniasis but of
probes is used to identify Leishmania in tissue slides (Barker, 1987,
1989; Kennedy, 1984; Schoone et al., 1991; van Eys et al., 1987). limited value for cutaneous disease, in which measurable antibody
Genus-specific fluorescence in situ hybridisation (FISH) targeting responses are rare. Serology in patients who are HIV-positive at the
the 18S rRNA gene (Frickmann et al., 2012) and ISH (Dinhopl et al., time of infection can remain negative and should not be used to
2011) targeting Leishmania-specific 5.8S RNA use a short labeled rule out visceral leishmaniasis. In patients infected first with leish-
genus-specific DNA-oligonucleotide probe and even work in diffi- mania and afterwards with HIV, even low titres have diagnostic
cult sample materials such as paraffin-embedded tissue specimens. value when combined with the clinical case definition. Western
blots can be used for predicting cases at highest risk of developing
Discrimination beyond the species level of Leishmania spp. is symptomatic leishmaniasis (Cota et al., 2012; Deniau et al., 2003).
nowadays possible by modern typing methods such as multilocus
enzyme typing (MLEE), multilocus sequence typing (MLST), and Cross reactivity can occur with Trypanosoma cruzi (Boecken
multilocus microsatellite typing (MLMT) (Schonian et al., 2011). et al., 2011). Chimeric antigens to serologically test for visceral
If such technologies are not available, traditional ISH based on leishmaniasis have a specificity of about 99% (Boarino et al., 2005).
hybridisation of kinetoplast DNA probes (Barker, 1989; Kennedy,
1984; Schoone et al., 1991; van Eys et al., 1987) can be used. Latex agglutination of visceral leishmaniasis antigens showed
93-100% specificity and 68-100% sensitivity (Attar et al., 2001). For
6.6.2. Reliability and critical interpretation of test results dipstick testing, sensitivity and specificity of 100% (Bern et al., 2000)
PCR is the most sensitive routine diagnostic approach for leish- were reported with animal specimens (Toz et al., 2004).

maniasis (Faber et al., 2003; Gangneux et al., 2003; Oliveira et al., 6.7.3. Value of serological approaches in the diagnostic process
2005; Romero et al., 2001). In HIV positive patients, PCR is the first Serology is restricted to suspected visceral disease, direct proof
indicator of relapse (Pizzuto et al., 2001). Positive Leishmania PCR
results can be obtained even from poor-quality sample materials of the pathogen is preferable. Rapid antigen tests are important
like Giemsa-stained slides (Pandey et al., 2010), in case of forensic for early diagnosis and management of visceral leishmaniasis in
resource-limited settings.

50 N.G. Schwarz et al. / Acta Tropica 165 (2017) 40–65

7. Rabies used post-mortem (Rudd et al., 2005). If applied to formalin-fixed
tissue, proteinase K pre-treatment dissociates chemical bonds and
7.1. Pathogen exposes rabies virus epitopes (Whitfield et al., 2001). As an alterna-
tive to DFA, direct rapid immunohistochemical testing (DRIT) with
Rabies is caused by varies species of the genus Lyssavirus (fam- biotinylated monoclonal antibodies against rabies nucleo-capsid
ily Rhabdoviridae, enveloped viruses of 100-430 nm x 45-100 nm in proteins can be used (Lembo et al., 2006). Negri bodies act as sites of
size), to which also belong species like Aravan virus, Australian bat viral transcription and replication. They are inclusions in neurons
lyssavirus, Duvenhage virus, European bat lyssavirus 1 & 2, Irkut between 1 and 7 ␮m in diameter containing viral nucleocapsids.
virus, Khujand virus, Lagos bat virus, and Mokola virus, respec- Traditional histological assessment by Seller’s staining allows for
tively (Kuzmin et al., 2005; Warrell and Warrell, 2004). The bullet the detection of Negri bodies (De Brito et al., 1973; Lahaye et al.,
shaped rabies virus contains a single-stranded non-segmented RNA 2009).
genome of negative polarity, which codes for 5 proteins (Fishbein
and Robinson, 1993). 7.4.2. Reliability and critical interpretation of test results
DFA testing lasts 3-4 h. Sensitivity and specificity are close to
7.2. Disease
100%. DFA is positive in about 50% of samples during the first
Lyssaviruses cause fatal encephalitis, leading to coma, respira- week with symptoms if multiple samples are stained. Positivity
tory paralysis and finally death (Hankins and Rosekrans, 2004). increases in later stages of the disease (Rudd et al., 2005). Sensi-
Rabies virus is transmitted by saliva of infected mammals, in par- tivity and specificity of DRIT are comparable to DFA (Lembo et al.,
ticular dogs. Neurons of the central nervous system are infected 2006). Negri bodies are only detectable in a small percentage of
by retrograde axonal transport with a speed of 8-20 mm / day infected cells (Jackson and Fenton, 2001; Jackson et al., 2001).
(Hemachudha et al., 2002). Neural cell adhesion protein (CD56) Identification of Negri bodies in rabid animals shows a sensitiv-
may confer neuronal attachment (Thoulouze et al., 1998). Mean ity of only 50-80% even if correctly performed (Perl and Good,
incubation time is 1-3 months but can vary from a few days to sev- 1991).
eral weeks (Boland et al., 2014; Hemachudha et al., 2002). Disease
starts with an unspecific prodromal state comprising fever, nausea, 7.4.3. Value of microscopic approaches in the diagnostic process
headache, and paraesthesia at the inoculation site. Later, a paralytic DFA or DRIT (Lembo et al., 2006; Rudd et al., 2005) are still in
(about 20%) or an encephalitic (about 80%) course develops.
use. DFA from postmortem brain tissue is the standard, by which
The encephalitic course is characterised by hydrophobia, pha- other diagnostic approaches are evaluated (Orciari and Rupprecht,
ryngeal spasms, fever, agitation, and hyper-salivation; the paralytic 2007).
course by increasing flaccid paresis, similar to Guillan Barré syn-
drome (Fishbein and Robinson, 1993; Hattwick, 1972; Hattwick 7.5. Cultural approaches
et al., 1972; Hemachudha et al., 2002; Rupprecht et al., 2002). Once
rabies manifests, it is virtually 100% fatal. 7.5.1. Procedures
Lyssaviruses can be cultured under BSL-3 conditions (Orciari
Lyssaviruses occur worldwide, only the Australian continent
is declared to be free. According to WHO estimations, approxi- and Rupprecht, 2007) in mouse neuroblastoma cells (cell lines
mately 60,000 individuals die from rabies every year (WHO, 2013). NA C 1300 or CCL 131). After inoculation of sample material
Prae- and postexposure prophylaxis exist (Manning et al., 2008; and incubation for 2-4 days, viral growth can be visualised using
Rupprecht et al., 2002). immunofluorescence-labelled anti-lyssavirus-antibodies (Webster
and Casey, 1996). Viral growth in mice is possible as well (Valleca
7.3. Diagnostic procedures and Forester, 1981). A laboratory infection via the airborne route
has been described (Winkler et al., 1973).

7.3.1. Sampling and pre-analytic considerations 7.5.2. Reliability and critical interpretation of test results
Sampling and pre-analytic aspects for suspected rabies are sum- Culture unambiguously demonstrates the presence of active

marised in the WHO Expert Consultation on Rabies: second report lyssavirus and allows for an enrichment for subsequent assess-
(WHO, 2013). ments, e.g. sequencing for epidemiological typing (Bourhy et al.,
2005; Kuzmin et al., 2004; Kuzmin et al., 2003; Nadin-Davis et al.,
Diagnostic materials from vital patients comprise bioptic mate- 1994; Nadin-Davis et al., 2006; Smits et al., 1999).
rials (5 -6 ␮m) from the nuchal skin with hair follicles (Dacheux
et al., 2008) and secretions like saliva, cerebrospinal fluid, tears 7.5.3. Value of cultural approaches in the diagnostic process
or serum. Due to intermitting virus shedding, 3 saliva samples in Diagnostic culture is rarely performed. Inoculation of mice has
intervals of 3-6 hours should be taken into a sterile container, and
for transport and storage cooled or frozen, preferably on dry ice. been abandoned because it is not superior to cell culture (Valleca
Long-term storage should be avoided. For transport at ambient and Forester, 1981).
temperature, samples should be stabilised in 50% glycerine-saline.
Material for post-mortem rabies diagnostics comprises brain biop- 7.6. Molecular diagnostic approaches
sies, e.g. from the brainstem, hippocampus, thalamus, cerebellum,
or medulla oblongata. 7.6.1. Procedures
Reverse transcriptase PCR (RT-PCR) and LAMP (loop mediated
7.4. Microscopic approaches
amplification) approaches show high sensitivity for the detection of
7.4.1. Procedures lyssavirus infections already ante mortem and allow for discrimina-
Microscopic assessment of suspected rabies comprises direct tion on molecular level from tissue, saliva and cerebrospinal fluids
(Coertse et al., 2010; Crepin et al., 1998; Fooks et al., 2009; Mani and
fluorescence antibody (DFA) testing (Rudd et al., 2005) with mon- Madhusudana, 2013; Wunner and Briggs, 2010). A high diversity
oclonal or polyclonal antibodies from at least 20 biopsies of vital among lyssavirus variants result in a lack of non-degenerate uni-
patients from the nuchal skin with hair follicles. Brain tissue, in versal primers, so multiple sets of degenerate primers have to be
particular cross-sections of brain stem and cerebellum, should be used if RT-PCR is used. E.g. degenerated primers for the detection of

N.G. Schwarz et al. / Acta Tropica 165 (2017) 40–65 51

the N gene of rabies virus strains found in the United States (Orciari as definite hosts. It principally occurs worldwide, but is most
and Rupprecht, 2007) were described, comprising six forward prevalent in countries where pork is frequently eaten if swine pop-
primers (21 g: 5 -ATG-TAA-CAC-CTC- TAC-AAT-G-3 , 10 g: 5 -CTA- ulations are infected.
CAA-CGG-ATG-CCG-AC- 3 , 1066 deg: 5 -GA(AG)-AGA-AGA-TTC-
TTC-AG(AG)-GA-3 , 1087i: 5 -GAG-AA(AG)-GAA- CTT-CA(AG)-GAi- Humans get infected by eating meat containing at least one larva
TA-3 , 1087s deg: 5 -GAG-AA(AG)-GAA-CTT-CA(AG)-GA-3 , 1004s: – the Cysticercus cellulosae. After entering the small intestine, a Cys-
5 - TCA-TGA-TGA-ATG-GAG-GT-3 ) and one reverse primer (304: ticercus releases the scolex and anchors itself to the mucosa, grows
5 -TTG-ACG-AAG-ATC-TTG-CTC-AT- 3 ). Sequence based typing within 12 weeks to an adult worm, reproduces by self-fertilisation,
makes use of N-gene-sequences (Kuzmin et al., 2004; Nadin-Davis and releases proglottides containing eggs to the faeces.
et al., 1994; Smith et al., 1995), supplemented by sequencing of
the G, P, and L-genes (Bourhy et al., 2005; Kuzmin et al., 2004; Swine ingest the embryonated eggs with food contaminated
Nadin-Davis et al., 2006) with human excreta. In the porcine intestine, eggs hatch and
unleash motile oncospheres that enter lymphatic and blood vessels.
7.6.2. Reliability and critical interpretation of test results Oncospheres migrate to striated muscles, liver, brain, and other tis-
Diagnostic reliability of PCR largely depends on sample material, sues. They settle in these organs and form cysts called cysticerci
(Peters and Pasvol, 2007).
mode of nucleic acid extraction, and the applied PCR protocol. Best
sensitivity can be achieved from brain tissue (Fooks et al., 2009; 8.2. Disease
Mani and Madhusudana, 2013; Wunner and Briggs, 2010). RT-PCR
allows determination of geographic origin of the virus and host Humans accidentally ingesting embryonated eggs may also
species origin regarding the biological reservoir (Arai et al., 1997; become incidental intermediate hosts. After formation of mus-
Nadin-Davis, 1998). Successful diagnosis from decomposed brain cular cysts, clinical symptoms of cysticercosis like eosinophilia,
material is also possible with RT-PCR (Whitby et al., 1997). fever, myositis, muscular pseudohypertrophy followed by atro-
phy and fibrosis develop. If cysts localise in the brain, so-called
7.6.3. Value of molecular diagnostic approaches in the diagnostic neurocysticercosis manifests with headaches, depression, brain-
process stem dysfunction, cerebellar ataxia, sensory deficits, involuntary
movements, stroke-like symptoms, extrapyramidal signs, demen-
If region specific primers (Orciari and Rupprecht, 2007) are tia, Bruns syndrome, Kluver-Bucy syndrome, cortical blindness,
available, PCR is the method of choice for rabies diagnostics and has and, most notably, epileptic seizures (Carpio, 2002). Approximately
widely replaced alternative approaches (Fooks et al., 2009; Mani 29% (about 14.5 million) of epilepsy cases in endemic countries
and Madhusudana, 2013; Wunner and Briggs, 2010). are associated with neurocysticercosis making it the most frequent
preventable cause of epilepsy in the developing world, therefore
7.7. Serological approaches it was included in the WHO NZD list (WHO, 2014). The mortality
rate is up to 50% when cysticercotic meningitis with hydrocephalus
7.7.1. Procedures occurs (Cardenas et al., 2010; Sotelo and Marin, 1987). However,
Neutralising antibodies against the G protein of rabies can be a high percentage of neurocysticercosis patients remain asymp-
tomatic (Carpio, 2002).
assessed by RFFIT/FAVN (rapid fluorescent focus inhibition testing
/ fluorescence antibody virus neutralisation) (Briggs et al., 1998; 8.2.1. Sampling and pre-analytic considerations
Cliquet et al., 1998; Smith et al., 1973). Robust neutralisation assays The clinical diagnosis of neurocysticercosis can be difficult
for serosurveillance have been described (Wright et al., 2009).
because of the non-specificity of clinical manifestations and neu-
IgG and IgM antibodies from blood and cerebrospinal fluid can roimaging findings (Garcia et al., 2002). A set of graded criteria
be detected using immunofluorescence assays (IFA). A stepwise including clinical, imaging, immunological, and epidemiological
dilution of the material is applied in thin layers on slides carrying data is used to classify suspected cases as definitive or probable
infected cell culture material and, e.g. neutralising antibodies like (Del Brutto, 2012; Del Brutto et al., 2001; Del Brutto et al., 1996;
protein G specific antibodies are assessed (Orciari and Rupprecht, Yancey et al., 2005).
2007). Standardised in-house or commercially available ELISA sys-
tems for the detection of anti-rabies virus antibodies have been The laboratory diagnosis of taeniasis and cysticercosis is based
introduced (Welch et al., 2009). on (i.) microscopic detection of proglottides in stool and taeniid
cysts in muscle or cerebral biopsies, on (ii.) detection of T. solium-
7.7.2. Reliability and critical interpretation of test result specific DNA in stool samples and muscle or cerebral biopsies, as
RFFIT/FAVN testing is used for the assessment of vaccination well as (iii.) detection of T. solium-specific antibodies in serum and
cerebrospinal fluid.
state (Moore and Hanlon, 2010). Few infected patients have neu-
tralising antibodies by day six of illness, 50% by day 8 and 100% by 8.3. Microscopical approaches
day 15. Any antibody titres in cerebrospinal fluid are diagnostically
relevant and specific oligoclonal bands in cerebrospinal fluid con- 8.3.1. Procedures
firm infection of the central nervous system (Alvarez et al., 1994). Microscopic diagnoses of T. solium associated diseases splits

7.7.3. Value of serological approaches in the diagnostic process into diagnosing taeniasis and diagnosing cysticercosis. While the
Serological assessment is of very limited value for the diagnosis diagnosis of taeniasis is based on morphologic identification of
proglottides in stool samples stained with India ink (Peters and
of acute rabies due to the late occurence of detectable antibodies Pasvol, 2007; Soulsby, 1982), the diagnosis of cysticercosis relies on
(Alvarez et al., 1994). microscopic examination of cysts that were dissected from muscle
tissue, subcutaneous tissue, and in case of neurocysticercosis from
8. Taenia solium: Neuro-/Cysticercosis and Taeniasis cerebral material. The taeniid cysts are exposed to 10% bile in order
to examine the evaginated rostellum.
8.1. Pathogen

Taenia solium, the pork tapeworm, is an intestinal parasite,
which passes through swine, as intermediate hosts, into humans

52 N.G. Schwarz et al. / Acta Tropica 165 (2017) 40–65

8.3.2. Reliability and critical interpretation of test results secretory antigens collected from in vitro cultured T. solium tape-
T. solium and T. saginata eggs cannot be differentiated by worms (Wilkins et al., 1999). Tsang and co-workers developed on
the basis of lentil-lectin, affinity-purified glycoprotein antigens an
microscopy, and should be discriminated using PCR (Jimenez et al., enzyme-linked immunoelectrotransfer blot (EITB) that can be used
2010). The assessment of the rostellum is relatively simple and to test serum as well as cerebrospinal fluid (CSF) samples (Tsang
unambiguous. More than 15 lateral uterine branches in the proglot- et al., 1989).
tid are characteristic for T. saginata, while less than 12 to 15
branches are typical for T. solium (Peters and Pasvol, 2007; Soulsby, 8.5.1. Reliability and critical interpretation of test results
1982). Sensitivity of the Wilkins’s immunoblot assay was 95% (69 of

8.3.3. Value of microscopy in the diagnostic process 73) using sera of patients with confirmed T. solium tapeworm infec-
Histological/microscopic demonstration of the parasite from tions. Testing of 193 sera of patients with other parasitic infections,
including T. saginata tapeworm infection, indicated a specificity of
biopsy of a brain or spinal cord lesion is one of the unambiguous 100% (Wilkins et al., 1999). The EITB was 98% sensitive and 100%
criteria for the diagnosis of cysticercosis. The method is limited specific (Tsang et al., 1989).
because cerebral material can often not be gained before autopsy.

8.4. Molecular diagnostic approaches 8.5.2. Value of serological approaches in the diagnostic process
Detection of anticysticercal antibodies with the serum EITB is
8.4.1. Procedures
Different PCR protocols are available for the diagnosis of tae- one of the major criteria and a positive result in a CSF ELISA
for detection of cysticercal antigens or anticysticercal antibodies
niasis and cysticercosis for simplex PCR (Gonzalez et al., 2000; belong to the minor criteria (Del Brutto, 2012; Del Brutto et al.,
Sreedevi et al., 2012), nested PCR (Mayta et al., 2007), multiplex 2001; Del Brutto et al., 1996; Yancey et al., 2005) for reliable diag-
PCR (Yamasaki et al., 2004a), and real-time quantitative PCR (Praet nosis (Table 6).
et al., 2013).
9. Trypanosomiasis
Most established PCR assays are designed for simultaneous
detection and differentiation of T. solium and T. saginata. Primarily 9.1. Pathogen
ubiquitous target sequences present in both Taenia species were
chosen, e.g. the oncosphere-specific protein gene tso31 (Mayta Human pathogenic trypanosomes (protozoan haemoflagellates)
et al., 2007), the cytochrome c oxidase subunit 1 gene cox1 are Trypanosoma cruzi, causal agent of the American trypanoso-
(Yamasaki et al., 2004b), the internal transcribed spacer 1 and miasis (Chagas disease), transmitted mainly in the Americas, and
spacer 2 (ITS-1 and ITS-2) (Garcia et al., 2002; Praet et al., 2013), and T. brucei, which causes human African trypanosomiasis (sleep-
the non-coding DNA fragments HDP1 and HDP2 (Gonzalez et al., ing sickness) in Sub-Saharan Africa. Only sleeping sickness due to
2000). For three well-established PCR assays, please see Table 5. the subspecies T. brucei rhodesiense in southern and East Africa
is considered a zoonosis, involving cattle and other mammals as
8.4.2. Reliability and critical interpretation of test results reservoirs. Humans are the main reservoir of T. brucei gambiense,
Compared to other diagnostic procedures including imaging, which is found in central and West Africa and accounts for more
than 90% of sleeping sickness (Barrett et al., 2003; Brun et al., 2010;
clinical, immunological, and microscopic methods, PCR ensures Franco et al., 2014; Mitashi et al., 2012). T. brucei is directly trans-
sufficient specificity and differentiates between T. solium and T. mitted by the bite of the tsetse fly, whereas T. cruzi is transmitted
saginata. indirectly by contaminated faeces of triatomine bugs. Infection
with trypanosomes can also occur via needle-stick injuries and
8.4.3. Value of molecular diagnostic approaches in the diagnostic blood transfusions (Barrett et al., 2003; Herrera, 2014; Urdaneta-
process Morales, 2014).

In the process of taeniasis and cysticercosis diagnosis, especially 9.2. Disease
nested-PCR (Mayta et al., 2008) but also simplex PCR (Garcia et al.,
2002) and qPCR (Praet et al., 2013) from stool samples are the tools 9.2.1. Chagas disease (ChD)
of choice to increase sensitivity and specificity in comparison to The acute phase mostly remains asymptomatic, but few patients
microscopic observation of eggs. Multiplex PCR from proglottid
samples or taeniid cysts dissected from muscle tissue of mammals experience mild symptoms within 7-14 days after vector-borne
are the tools of choice for the species-specific diagnosis of T. asi- transmission, including fever, malaise, oedema, and chagoma or
atica, T. saginata, and T. solium and for the differentiation of the Roman˜ a sign at the parasite’s entry site. Patients infrequently
two T. solium genotypes (Yamasaki et al., 2004b). In developing develop lethal complications, but usually symptoms resolve within
countries, in particular in tropical regions, these assays should be 2 months without treatment, until 10-30 years later approximately
transformed into loop-mediated isothermal amplification (LAMP) 40% of patients are diagnosed with chronically persistent manifes-
assays. Portable integrated systems with high reproducibility ful- tations, such as megaviscera and dilated cardiomyopathy. Mortality
filling high quality standards are required. LAMP systems that are varies depending on transmission route, phase, clinical presenta-
already successfully used in the tropics, are the Genie® system tion and progression of ChD (Nunes et al., 2013; Rassi et al., 2010).
(Amplex Biosystems GmbH, Gießen, Germany) or the Realtime Tur-
bidimeter LA-200 (Eiken Chemical Co. LTD., Tokyo, Japan). 9.2.2. Human African trypanosomiasis (HAT)
The acute haemo–lymphatic stage presents few days to weeks
8.5. Serological approaches
after infection with unspecific symptoms such as fever, headache,
Several T. solium tapeworm antigen preparations have been malaise, myalgia, lymphadenopathy, or a trypanosomal chancre.
developed, allowing identification of tapeworm carriers (Ferrer The second meningo–encephalitic stage is characterised by neu-
et al., 2007; Hancock et al., 2004; Hancock et al., 2006; Levine ropsychiatric symptoms and abnormal sleeping patterns due to
et al., 2004; Wilkins et al., 1999). The earliest serologic test, an parasitic penetration into the central nervous system. Without
immunoblot assay developed by Wilkins and co-workers, used treatment, patients usually die within 6 months, when infected

N.G. Schwarz et al. / Acta Tropica 165 (2017) 40–65 53
Cycleno. Annealing
Table 5 25 55 ◦C for 30 s
PCR approaches for Taenia. 40 60 ◦C for 30 s
50 60 ◦C for 30 s
Purpose of the PCR Primer name Oligonucleotide sequence Amplicon size
5 →3 in bp 35 60 ◦C for 30 s
T. solium nested PCR Mayta et al. (2008)

nested PCR F589


copro-PCR (qPCR) for the detection of T. solium and T. saginata DNA Praet et al. (2013)






Multiplex PCR the detection of T. asiatica, T. saginata and for American/African and Asian genotypes of T. solium Yamasaki et al. (2004a)

cytochrome coxidasesubunit 1(cox1) Tsag TTGATTCCTTCGATGGCTTTTCTTTTG 827





Abbreviations: BHQ: black hole quencher; CT: Q7: Quasar705 fluorescence dye, bp: base pairs, no.: numbers.

Table 6
Diagnostic criteria for neurocysticercosis and categories of diagnostic certainty (revised version, Del Brutto, 2012 (Del Brutto, 2012)).

Category Diagnostic criteria
• Histological demonstration of the parasite from biopsy of a brain or spinal cord lesion
• Evidence of cystic lesions showing the scolex on neuroimaging studies
• Direct visualisation of subretinal parasites by fundoscopic examination

Major • Evidence of lesions highly suggestive of neurocysticercosis on neuroimaging studies
• Positive serum immunoblot for the detection of anticysticercal antibodies
• Resolution of intracranial cystic lesions after therapy with albendazole or praziquantel
• Spontaneous resolution of small single enhancing lesions

Minor • Evidence of lesions compatible with neurocysticercosis on neuroimaging studies
• Presence of clinical manifestations suggestive of neurocysticercosis
• Positive CSF ELISA for detection of anticysticercal antibodies or cysticercal antigens
• Evidence of cysticercosis outside the central nervous system

Epidemiologic • Individuals coming from or living in an area where cysticercosis is endemic
• History of travel to disease-endemic areas
• Evidence of a household contact with T. solium infection

Legend: definitive diagnosis of neurocysticercosis: (a.) Presence of one absolute criterion; (b.) Presence of two major criteria plus one minor and one epidemiological criterion;
Probable diagnosis of neurocysticercosis: (a.) presence of one major criterion plus two minor criteria; (b.) presence of one major plus one minor and one epidemiological
criterion; (c.) presence of three minor criteria plus one epidemiological criterion.

with T. brucei rhodesiense, or within 3 years, if disease is caused by films or concentration methods, comprising mini haematocrit
T. brucei gambiense (Brun et al., 2010; Chappuis et al., 2005; Franco centrifugation technique (mHCT), quantitative buffy coat (QBC),
et al., 2014). and miniature anion-exchange centrifugation technique (mAECT).
Strout’s method (Diez et al., 2007) is a serum precipitate concen-
9.3. Diagnostic procedures tration technique used for the diagnosis of ChD. For patients at
risk for HAT, fresh aspirates from suspicious cervical lymph nodes
9.3.1. Sampling and pre-analytic considerations are obtained (Mitashi et al., 2012). For the second HAT stage CSF
Blood, lymph node aspirates and cerebral spine fluid (CSF) is centrifuged for microscopic examination (Chappuis et al., 2005;
Mitashi et al., 2012; Njiru et al., 2008).
for microscopy have to be processed immediately, or within few
hours after sampling if kept cool and dark, to prevent lysis of try- 9.4.2. Reliability and critical interpretation of test results
panosomes (Brun et al., 2010; Chappuis et al., 2005). For serology, High T. brucei rhodesiense parasitaemia in the acute stage allows
whole blood, serum or plasma is required. For molecular diagnos-
tics whole blood, buffy coat (BC), lymph node aspirates, tissues diagnosis by wet or Giemsa stained thin and thick blood films. For
(Diez et al., 2007), or CSF (Mitashi et al., 2012) can be used. microscopic detection of T. brucei gambiense, blood concentration
methods may be required due to short and low parasitaemia (Brun
9.4. Microscopic approaches et al., 2010; Chappuis et al., 2005; Matovu et al., 2012). Sensitiv-
ity for microscopic detection of T. brucei gambiense in blood varies
9.4.1. Procedures from 4-54% for wet or thin blood films and 26-100% for thick blood
Detection of trypanosomes by traditional microscopy requires films, but can be increased using concentration techniques (Mitashi
et al., 2012). Less than 50 trypanosomes/ml may be detected by
400x magnification. Capillary or anticoagulated venous blood is mAECT as the most sensitive method, which can even be improved
used for wet blood films, Giemsa-stained thick and thin blood

54 N.G. Schwarz et al. / Acta Tropica 165 (2017) 40–65

if used with buffy coats (mAECT-BC) and combined with lymph (Agranoff et al., 2005; Papadopoulos et al., 2004; Ricciardi and Ndao,
node aspiration (Camara et al., 2010; Mumba Ngoyi et al., 2014). 2015).
For lymph node aspiration sensitivity is reported to be only 19-64%
(Mitashi et al., 2012), with higher sensitivity in the acute stage (Brun 9.7. Molecular diagnostic approaches
et al., 2010). Modified single centrifugation of CSF allows detection
of less than 2 trypanosomes/ml in less than 15 minutes in second 9.7.1. Procedures
stage HAT (Buscher et al., 2009). In the acute phase of ChD, T. cruzi For HAT, conventional polymerase chain reaction (PCR), real-
can be detected in blood smears, however, only with a low sen-
sitivity (WHO, 2012). Strout’s method or mHCT are considered as time PCR and oligochromatography-PCR (OC-PCR) were described
diagnostic reference methods for ChD (Lescure et al., 2010). and evaluated. For DNA detection on subgenus level (T. brucei
genome), satellite, kinetoplast or ribosomal DNA, ITS1 or ESAG 6/7
9.4.3. Value of microscopic procedures in the diagnostic process are targeted by PCR. In contrast, loop-mediated isothermal amplifi-
Apart from chronic phase of ChD (WHO, 2012), detection of try- cation (LAMP) assays mainly target the PFRA gene or the repetitive
insertion mobile element (RIME), allowing amplification of DNA
panosomes in blood, and cerebral spine fluid (CSF), lymphatic or sequences under isothermal conditions (Table 7). To discriminate
chancre fluid for HAT, remains the gold standard for the confir- T. brucei gambiense and T. brucei rhodesiense, PCRs and LAMPs, tar-
mation of trypanosomiasis (Ricciardi and Ndao, 2015). Staging of geting the TgsGP and SRA genes, are available. Nucleic acid-based
HAT, based on white blood cell count and presence of trypanosomes amplification (NASBA) provides a technique for RNA amplification
determined in CSF, is essential for the choice of treatment (Matovu under isothermal conditions. Furthermore, OC represents a dipstick
et al., 2012; Mumba Ngoyi et al., 2013). solution in HAT diagnostics for the detection of previously ampli-
fied DNA or RNA by PCR or NASBA (Deborggraeve and Buscher,
9.5. Cultural approaches 2012; Mitashi et al., 2012). For the diagnosis of ChD an OC-PCR,
the T. cruzi OligoC-TesT (Deborggraeve et al., 2009), targeting the
9.5.1. Procedures satellite DNA sequence, is commercially available, whilst many
Trypanosomes can be detected by mouse inoculation (Holland other PCR methods still require standardisation and optimisation
(Lescure et al., 2010; Ricciardi and Ndao, 2015).
et al., 2001; Rickman and Robson, 1970) or by haemoculture in HAT
(Chappuis et al., 2005) and in the acute phase of ChD (Santamaria 9.7.2. Reliability and critical interpretation of test results
et al., 2014). However, these procedures require specialised per- PCR methods for HAT have a sensitivity between 70–100% and a
sonnel, laboratory bio-safety 2 or 3 (in case of ChD suspicion)
conditions and about one month of time (Diez et al., 2007). specificity of 92–100%, with reduced performance for OC-PCR. The
most sensitive PCRs target the satellite and ribosomal DNA, pro-
9.5.2. Reliability and critical interpretation of test results viding a detection limit per reaction (LOD) below 1 parasite, but
Sensitivity of blood culture for the diagnosis of ChD is lim- in comparison, RIME LAMP (Table 7) is the most sensitive method
for DNA detection (LOD < 0.001 parasites), followed by RNA detec-
ited (Bhattacharyya et al., 2014) and varies when used for HAT tion by NASBA (0005 parasites). If subspecies differentiation by
(Chappuis et al., 2005). LAMP is desired, LOD decreases to an acceptable level (1-10 par-
asites), compared to PCR (LOD ≈ 100 parasites). For ChD and the
9.5.3. Value of cultural approaches in the diagnostic process T. cruzi OligoC-TesT, sensitivity of 94% and specificity of 100% were
Whilst the need of a fast-established diagnosis disqualifies blood reported (Deborggraeve and Buscher, 2012; Matovu et al., 2012;
Mitashi et al., 2012).
culture as diagnostic tool for HAT in the field (Chappuis et al., 2005),
it is still described as an option for diagnosis of ChD (Bhattacharyya 9.7.3. Value of molecular diagnostic approaches in the diagnostic
et al., 2014; Lescure et al., 2010). process

9.6. Matrix-assisted laser desorption ionisation time-of-flight For HAT, LAMP and NASBA require less equipment and provide
mass spectrometry operational benefits, while showing equal or improved sensitivity
compared to PCR, which remains a helpful technique for scien-
9.6.1. Procedures tific or epidemiological use and travel medicine. LAMP is the only
Surface-enhanced laser-desorption-ionisation time-of-flight molecular technique, which might qualify for diagnosis in the field,
whereas by targeting RNA, the field use of NASBA is limited by its
mass spectrometry (SELDI-TOF MS) has been successfully com- high vulnerability for degradation, although it might be favourable
bined with data-mining tools for the detection of specific for therapy control (Deborggraeve and Buscher, 2012; Matovu
proteins in serum, expressed in HAT (Papadopoulos et al., et al., 2012; Mitashi et al., 2012). For ChD, PCR could provide a
2004). Furthermore, biomarkers strongly associated with ChD, sensitive method for congenital screening, follow up, and reacti-
have been identified in sera by the use of SELDI-TOF and vation monitoring in immunocompromised patients (Lescure et al.,
matrix-assisted laser-desorption-ionisation time-of-flight mass 2010; Ricciardi and Ndao, 2015). The commercially available T. cruzi
spectrometry (MALDI-TOF MS) (Ndao et al., 2010; Santamaria et al., OligoC-TesT can be used in laboratories with midlevel equipment
2014). and its simplicity might even be improved by implementing NASBA
or LAMP methods (Deborggraeve et al., 2009).
9.6.2. Reliability and critical interpretation of test results
SELDI-TOF MS showed 100% sensitivity and 98.6% specificity 9.8. Serological approaches

for the use in patients with HAT (Papadopoulos et al., 2004). 9.8.1. Procedures
Comparable accuracy for identified ChD biomarkers via SELDI and Antibodies against T. brucei gambiense can be detected by the
MALDI-TOF has been reported (Santamaria et al., 2014).
card agglutination test for trypanosomiasis (CATT) in blood, plasma
9.6.3. Value of time-of-flight (TOF) mass spectrometry or serum after 5 min incubation or by lateral flow rapid diagnos-
approaches in the diagnostic process tic tests, detecting low antibody concentrations from finger prick
blood within 15 minutes (Sternberg et al., 2014; Sullivan et al.,
MALDI-TOF, and its refined variation SELDI-TOF work with
small amounts of (even haemolytic) serum, but require accurate
and time-consuming protocols and technical expensive equipment

N.G. Schwarz et al. / Acta Tropica 165 (2017) 40–65 55

Table 7
Primers of the highly sensitive RIME LAMP for the amplification of trypanozoon-specific DNA sequences (T. brucei genome). F3 and B3 primers are not required for LAMP
reaction, but used for initial strand displacement (Njiru et al., 2008).

Primer Type Sequence Length Target

2014). Further methods comprise indirect agglutination latex test- Due to the pathological potential of the causative agents, labo-
ing, indirect immunofluorescence (IIF), enzyme-linked immuno ratory safety has to be taken into account. We therefore discuss
sorbent assays (ELISA), and immune trypanolysis test (TL) (Brun such laboratory safety considerations in the absence of a BSL-3
et al., 2010; Mitashi et al., 2012; Sullivan et al., 2013). Antibod- laboratory (Discussion I).
ies against T. cruzi are detected by IIF, indirect haemagglutination,
ELISA and immunoblot. A mix-ELISA, combining recombinant anti- When considering the consequences of false diagnoses of such
gens, has been described for potential use (Lescure et al., 2010; hp-NZDs for the individual, one has to consider the harmful effects
Ricciardi and Ndao, 2015). of the disease in case of a false negative diagnosis. In contrast, there
might be severe side effects of treatment in case of a false positive
9.8.2. Reliability and critical interpretation of test results diagnosis. Treatment will be covered by other reviews in this Acta
CATT is reported to be 87-98% sensitive and 95% specific (Brun Tropica special edition. However, we want to remind the reader
that some drugs used, e.g. for treating leishmaniasis or trypanoso-
et al., 2010; Sullivan et al., 2014). Lateral flow RDTs were 82-88% miasis, are quite toxic. To sensitise the reader for the need of careful
sensitive and 94-97% specific (Sternberg et al., 2014). In compari- interpretation of test results, we close this review on the diagno-
son, TL might have a sensitivity of 97-100% and specificity of 100%, sis of hp-NZDs with some general remarks on the interpretation
however, TL use is limited by equipment requirements and high of test results and the diagnosis as sequel of multiple subsequent
infection risk for laboratory personnel (Mitashi et al., 2012). For tests (Discussion II).
detection of antibodies against T. cruzi, IIF and ELISA are the most
sensitive ones among the standardised tests, although use of the Most hp-NZDs are diagnosed in poor countries, where the lack
mix-ELISA might offer advantages (Ricciardi and Ndao, 2015). of diagnostic laboratory facilities will not allow for carrying out
some of the above-mentioned tests. Activities to improve health
9.8.3. Value of serological approaches in the diagnostic process care in poor countries often address prevention and treatment
Simple and fast screening for infection with T. brucei gambiense of infectious diseases, considering the reliable diagnosis as given.
If health care improvements are not only measured as “number
is performed by CATT or thermostable CATT-D10 (Hasker et al., of patients treated”, but also take the success of treatment into
2010) and may be simplified cost effectively by the use of lateral account, it becomes clear that reliable diagnosis is a prerequisite for
flow RDTs, whilst screening for infection with T. brucei rhodesiense proper treatment. Diagnostic capacities are essential for an opera-
is limited to history of exposure and clinical signs of individuals tive health system that serves the population (Petti et al., 2006).
(Brun et al., 2010; Matovu et al., 2012; Mitashi et al., 2012; Njiru
et al., 2008; Sternberg et al., 2014). The TL could be a useful tool for 10.1. Discussion I: diagnostic management in the absence of a
HAT surveillance (Jamonneau et al., 2010). ChD serology plays an BSL-3 laboratory
important role for screening of blood donors at risk and diagnosis of
chronic stages, for which positive results by at least 2 different sero- If no BSL-3 laboratory facilities are available, any diagnostic
logical methods, determined with the same sample, are required procedure with vital organisms of BSL-3 category, in particularly
(Lescure et al., 2010; Ricciardi and Ndao, 2015). cultural growth or enrichment, should be discouraged. However,
inadvertent cultural growth in case of lacking clinical suspicion
9.8.4. Diagnostic issues even of pathogens like Bacillus anthracis under BSL-2 conditions
Realisation of large-scale HAT screening programmes, allowing occurs. Such undesirable situations do not necessarily lead to lab-
oratory infections, if good laboratory practice is obeyed, direct
early diagnosis before the meningo–encephalitic stage, which is contacts are minimised, and appropriate hand hygiene agents are
lethal if left untreated, is limited by lack of human and material applied (CDC, 1999; Weber et al., 2003).
resources (Brun et al., 2010; Matovu et al., 2012; Mitashi et al., 2012;
Njiru et al., 2008; Sternberg et al., 2014). In America diagnostics for If abstinence from cultural approaches is required for safety rea-
ChD are available and implemented in urban settings, however, sons, serology and traditional microscopy or molecular approaches
accuracy is limited and improved tests are needed for diagnosis have to be considered. Preferably, they should be performed with
and prognosis as for blood bank screening to reduce transfusion inactivated samples if the diagnostic procedures are validated for
associated transmission. Furthermore, in rural areas serological and such materials.
molecular methods, allowing diagnosis in the chronic stage, are
usually not provided (Ndao et al., 2010; Porras et al., 2015). Diagnostic procedures with inactivated biological material are
preferable whenever possible without loss of diagnostic qual-
10. Discussion and conclusion ity. Validated procedures for inactivation depend on the kind of
pathogen. Spores and mycolic acid containing cell walls of, e.g.
We described laboratory diagnostic tests for 8 highly pathogenic mycobacteria, are particularly resistant in the environment and
neglected zoonotic diseases (hp-NZDs), anthrax, bovine tuber- to inactivating procedures (Logan et al., 2007; Mitscherlich and
culosis, brucellosis, echinococcosis, leishmaniasis, rabies, Taenia Marth, 1984; Weber et al., 2003). Heat >65 ◦C, direct UV radia-
solium-associated diseases (neuro-/cysticercosis & taeniasis) and tion, ethylene oxide, formaldehyde vapour, chlorine compounds,
trypanosomiasis and also addressed challenges around the diag- 70% ethanol (in non-protein-containing materials), 2% alkaline glu-
nostical methods that we tried to concisely summarise in Table 8. taraldehyde, peracetic acid, iodophors depending on the presence

56 N.G. Schwarz et al. / Acta Tropica 165 (2017) 40–65

Table 8 Anthrax Bovine tuberculosis Brucellosis Echinococcosis Leishmaniasis
Issues in diagnosing NZDs.
Identification of B. Reliable identification Reliable identification Diagnosis requires Discrimination of
Is there a diagnostic issue? anthracis by traditional requires sequencing of Brucella spp. beyond clinicians, radiologists Leishmania spp. to
bio-chemical approaches. Culture the genus level by and micro-biologists. identify pathogens
Is the diagnostic issue approaches is hardly allowing drug traditional biochemical No reliable associated with
essential for the possible. Sophisticated sensitivity testing is approaches is standardised mucocutaneous
management of the cases approaches like important, but requires impossible. serological assay for disease requires
and/or the control of the MALDI-TOF-MS or PCR a BSL-3 laboratory. Sophisticated the detection of cystic sophisticated
disease? are required. approaches like echinococcosis, with molecular approaches.
No. With the MALDI-TOF-MS or PCR clinical correlates
What methods are Yes. Antimicrobial exemption of are obligatory. available No. If mucocutaneous
convenient for treatment of anthrax pyrazinamide- Yes. Reliable Yes. The absence of disease is not excluded,
tropical/developing differs from treatment resistance of M. bovis identification is reliable markers for treatment regimens
countries or in the field? of other infectious spp. bovis, the clinical important for both cystic echinococcosis will usually comprise
diseases with similar management is similar laboratory safety and delays treatment and this option.
clinical symptoms. to M. tuberculosis treatment. precludes judgment of
infections. treatment success. Microscopy
Automated Microscopy still widely Automated
point-of-care PCR used point-of-care PCR No reliable & simple to HAT
use laboratory test
available Yes. Diagnosis relies on
micro-scopy (low
Is there a diagnostic issue? Rabies T. solium/cysticercosis Chagas disease sensitivity). Serological
screening only
Immunofluorescence or A complex algorithm Yes. Early stage available for T. brucei
immunohistochemistry comprising imaging, diagnosis relies on gambiense. Current
requires considerable symp-toms and microscopy (low points of care tests still
experience. PCR serology is required for sensitivity). Tests with require certain
approaches are hampered a reliable diag-nosis of improved accuracy for infrastructure for
by a high degree of genetic neuro-cysti-cercosis. diagnosis, prognosis molecular biology.
diversity. and blood bank Yes. HAT is lethal, if left
screening are needed. untreated, but severe
side effects due to
Is the diagnostic issue essential No. Rabies is basically a Incorrect diagnosis Yes. Early diagnosis is toxicity of the
for the management of the clinical diagnosis. may lead to delayed or crucial for most treatment demand for
cases and/or the control of incorrect treatment. effective treatment in reliable diagnosis.
the disease? acute stage and
No reliable & simple to prevention from Microscopy, serology
What methods are convenient Immunofluorescence/ use laboratory test irreversible chronic (CATT/Lateral Flow
for tropical/developing immunohistochemistry available manifestations. RDTs)
countries or in the field? Microscopy,
serology/OC-PCR (T.
HAT = human african trypanosomiasis. cruzi, OligoC-TesT)

of organic matter, and stabilised hydrogen peroxide can be used for direct pathogen detection have limitations. The interpretation of
the inactivation of Mycobacterium spp. (Best et al., 1990). each diagnostic test should generally consider information on the
diagnostic procedure and its performance (sensitivity, specificity,
Reliable inactivation of anthrax spores demands 5% formalde- lower detection limit, positive and negative predictive value), on
hyde or glutaraldehyde solution, a pH 7-adjusted 1:10 dilution limitations, possible errors, disturbances, interference, and cross
of household bleach, or an aqueous solution of 500 mg/L chlo- reactions, availability of quality control procedures as well as
rine dioxide (Logan et al., 2007). Testing of reliable inactivation known reference values, and sample quality. In case of rare infec-
should comprise titration of the least harsh, but securely inacti- tious disease, even reliable respective information may be scarce.
vating approach and the measurement of time-inactivation-curves Of course, the above-mentioned general rule also applies to culture-
as recently demonstrated for rickettsiae (Frickmann and Dobler, dependent approaches.
Some procedures, e.g. the measurement of antimicrobial resis-
Harsh inactivation procedures can affect the quality of sub- tant patterns and several typing approaches as described in the
sequent diagnostic approaches. E.g. release of large quantities of pathogen-specific chapters, make cultural approaches yet desir-
human DNA from the sample or of heme from lyzed erythro- able. Therefore, reference laboratories with BSL-3 safety standards
cytes may inhibit PCR reactions (Alaeddini, 2012). Accordingly, are needed even in resource-limited settings.
diagnostic procedures always have to be evaluated together with
the complete pre-analytic procedure to ensure reliable and repro- For resource-limited settings without highly developed labo-
ducible results. Evaluated pre-analytic procedures have to be ratory infrastructure, point-of-care molecular diagnostic systems
maintained in the routine diagnostic setting. are usually the method of choice. Examples for such easy-to-use
point-of-care solutions comprise, e.g. the PCR-based GenXpert sys-
If no vital pathogen can be isolated and subjected to exten- tem (Cepheid, Sunnyvale, CA, USA), the FilmArray system (BioFire
sive further analyses to ensure its identity, alternative, usually Diagnostics, LLC, Salt Lake City, UT, USA), the cobas Liat system
non-cultural diagnostic approaches have to be chosen. As shown (Roche, Basel, Switzerland), and the loop-mediated amplification
in the pathogen-specific chapters, various alternatives to cultural (LAMP)-based Genie II device (amplex, Gießen, Germany). The
diagnostic approaches exist but all non-cultural approaches of

N.G. Schwarz et al. / Acta Tropica 165 (2017) 40–65 57

extent of quality control procedures, which should usually com- racy of each single sub-test and of course the “human factor”. An
prise extraction and inhibition control reactions for molecular accurate test result is not only important for the care seeker. Pro-
diagnostic procedures is usually defined by the manufacturer for viding reliable test accuracy information is important for reliable
fully automatic molecular diagnostic devices. It is an issue of cur- estimations of prevalence (Gart and Buck, 1966; Rogan and Gladen,
rent debate whether deviations from the strict regulations for 1978) or treatment effects (Lachenbruch, 1998) in epidemiological
molecular labs can be accepted for such fully-automated, closed studies and in clinical trials. Therefore, every diagnostic procedure
systems to save effort and money. has to be evaluated for its accuracy if it is planned to conduct it in
scientific context.
10.2. Discussion II: some general remarks on diagnosis as sequel
of multiple subsequent tests

We provided an overview of diagnostic tests for NZDs. For the 10.3. Discussion III: simple rapid diagnostic tests are needed, but
interpretation of a test result and its consequence, for example should not preclude developing countries from state of the art
regarding treatment, the clinician also has to keep in mind that diagnostics
all diagnostic tests have limitations. Ideally sensitivity and speci-
ficity of a test are 100%, however, hardly any diagnostic test reaches In our review, we included some very sophisticated diagnostic
these values. A false negative result leads to disadvantages for the techniques from the frontline of scientific development, such as
patient, who will not receive a required treatment, while a false pos- MALDI-TOF or next generation sequencing. Yet it is unlikely that
itive result may burden the patient with side effects of subsequent such approaches may substantially impact disease control mea-
treatment measures. sures for NTDs or NZDs on the short term, as they will hardly
be available on a large scale for the affected. Large impact can
When assessing the risk for false (positive or negative) test rather be expected from cheap and easy to use tests that are usu-
results, one has to consider the sensitivity and specificity of a test ally called “rapid diagnostic tests” (RDTs). Usually RDTs are strip
and the relative frequency of the disease in the population, to which tests containing antibodies to detect the diagnostic agent, i.e., a
the tested individual belongs to. Following this concept, individuals pathogen specific antigen in urine or serum. Nucleic acid sequence
presenting with disease typical symptoms belong to the “popula- specific molecular rapid tests as mentioned above are alternatives
tion of disease typical symptoms”. Such people have a much higher of increasing importance even in the tropics (Scott et al., 2014;
relative frequency of the real disease than the general population. Wekesa et al., 2014). Before such protein- or nucleic acid-based
The positive predictive value of the test will consequently be higher, tests can be used in disease diagnostics and control programmes,
and the negative predictive value lower. In this concept, “preselect- their performance (sensitivity and specificity) has to be assessed
ing through assessing disease typical symptoms” is a test to select and demonstrated to be sufficient.
for the population of individuals to be tested.
Test assessments are usually made against an established
Each individual laboratory diagnostic test is therefore only one method considered to be a standard. To assess the real value of
test in a sequel of multiple subsequent tests. Every test step has its a new test, one need to be able to compare positive and negative
own test accuracy. The first step is to define, which cases have to be test results against truly-infected versus non-infected (true positive
selected for a laboratory diagnostic test. If it is not a screening test, and true negative) samples.
this selection commonly depends on a couple of symptoms. Proving
each symptom is a sequel of multiple subsequent tests, too. Every In the absence of a gold standard, comparative test methods in
of these “symptom tests” has a sensitivity and specificity. A set of combination could be used as a composite reference standard to
initial symptoms can thus be regarded as requirement for each lab- classify a sample as being “true positive” or “true negative”. This
oratory test. The laboratory test is a combination of several steps is part of a TDR/WHO recommended strategy for evaluating new
(e.g. sampling of material for the test, accurate labelling, transport diagnostic methods in the absence of a gold standard (WHO-TDR-
under appropriate conditions, etc.). Diagnostics-Evaluation-Expert-Panel et al., 2010).

Not the test accuracy and performance of a lab test, which is Designing reliable composite reference panels is crucial in the
reflected by the sensitivity and specificity given in the accompany- planning phase of studies assessing the performance of a new test.
ing information of the laboratory test assay, matter to the clinician. As part of such a composite reference standard the abovemen-
Instead, reliable results of the “combined test” are of importance, tioned sophisticated technologies can be precious instruments for
consisting of the sequel of multiple subsequent tests including the development of new RDTs.
symptom assessment, clinical examination, the actual laboratory
test and the steps during the test preparations, where handling Beyond population-based infectious disease screenings by epi-
errors can additionally influence the test performance. demiologists, reliable test results for the individual patient are
the most important goal for the physician in charge. Sensitivities
The system of the sequel of multiple subsequent (clinical and and specificities that are sufficient for epidemiological assessments
laboratory) tests can be described as follows: might be deleterious for the individual by leading to wrong ther-
apy or – even worse – cohort isolation with truly infected patients
The clinical requirement to confirm the need of a laboratory with highly contagious diseases and resulting nosocomial infec-
diagnostic test is a combination of k tests that prove clinical symp- tions. Validity is more important than rapidity for diagnostic test
toms. Each k test has its own sensitivity and specificity. The overall results and incorrect test results can lead to worse consequences
sensitivity and specificity of the clinical symptom combination is for the patient than no results at all because they mislead the
therefore a function of its k sub-sensitivities and k specificities. The differential-diagnostic thinking of the physician.
same is true for the laboratory diagnostic approach. If l tests are
conducted and each of them has its own sensitivity and specificity, Therefore one should keep in mind that RDTs with limited
the overall accuracy for the laboratory-based diagnosis is a function sensitivity and specificity can only be bridging technologies for
of l sensitivities and l specificities. Finally, the diagnostic accuracy of resource-limited settings as long as better alternatives are not
the whole diagnostic procedure is a function of the functions of the available. However, the implementation of state-of-the-art tech-
test accuracy for the clinical and laboratory diagnostic elements. nologies in all parts of the world has to be aspired on the long term
(Petti et al., 2006).
Sensitivity and specificity of a diagnostic procedure is influenced
by the complexity of the diagnostic procedure itself, the test accu-

58 N.G. Schwarz et al. / Acta Tropica 165 (2017) 40–65

References lineage-specific serology for chronic Chagas disease: geographical and clinical
distribution of epitope recognition. PLoS Negl. Trop. Dis. 8, e2892.
Abshire, T.G., Brown, J.E., Ezzell, J.W., 2005. Production and validation of the use of Boarino, A., Scalone, A., Gradoni, L., Ferroglio, E., Vitale, F., Zanatta, R., Giuffrida,
gamma phage for identification of Bacillus anthracis. J. Clin. Microbiol. 43, M.G., Rosati, S., 2005. Development of recombinant chimeric antigen
4780–4788. expressing immunodominant B epitopes of Leishmania infantum for
serodiagnosis of visceral leishmaniasis. Clin. Diagn. Lab. Immunol. 12,
Adekambi, T., Colson, P., Drancourt, M., 2003. rpoB-based identification of 647–653.
nonpigmented and late-pigmenting rapidly growing mycobacteria. J. Clin. Boecken, G., Sunderkotter, C., Bogdan, C., Weitzel, T., Fischer, M., Muller, A.,
Microbiol. 41, 5699–5708. Lobermann, M., Anders, G., von Stebut, E., Schunk, M., Burchard, G., Grobusch,
M., Bialek, R., Harms-Zwingenberger, G., Fleischer, B., Pietras, M., Faulde, M.,
Agranoff, D., Stich, A., Abel, P., Krishna, S., 2005. Proteomic fingerprinting for the Erkens, K., Germany Society of, D., German Society of Tropical, M., German
diagnosis of human African trypanosomiasis. Trends Parasitol. 21, 154–157. Society of, C., 2011. [Diagnosis and therapy of cutaneous and mucocutaneous
Leishmaniasis in Germany]. Journal der Deutschen Dermatologischen
Agren, J., Hamidjaja, R.A., Hansen, T., Ruuls, R., Thierry, S., Vigre, H., Janse, I., Gesellschaft 9 (Suppl. 8), 1–51.
Sundstrom, A., Segerman, B., Koene, M., Lofstrom, C., Van Rotterdam, B., Boland, T.A., McGuone, D., Jindal, J., Rocha, M., Cumming, M., Rupprecht, C.E.,
Derzelle, S., 2013. In silico and in vitro evaluation of PCR-based assays for the Barbosa, T.F., de Novaes Oliveira, R., Chu, C.J., Cole, A.J., Kotait, I., Kuzmina, N.A.,
detection of Bacillus anthracis chromosomal signature sequences. Virulence 4, Yager, P.A., Kuzmin, I.V., Hedley-Whyte, E.T., Brown, C.M., Rosenthal, E.S., 2014.
671–685. Phylogenetic and epidemiologic evidence of multiyear incubation in human
rabies. Ann. Neurol. 75, 155–160.
Al Dahouk, S., Tomaso, H., Nockler, K., Neubauer, H., 2004. The detection of Brucella Bourhy, H., Cowley, J.A., Larrous, F., Holmes, E.C., Walker, P.J., 2005. Phylogenetic
spp. using PCR-ELISA and real-time PCR assays. Clin. Lab. 50, 387–394. relationships among rhabdoviruses inferred using the L polymerase gene. J.
Gen. Virol. 86, 2849–2858.
Al Dahouk, S., Tomaso, H., Nockler, K., Neubauer, H., Frangoulidis, D., 2003a. Braz, R.F., Nascimento, E.T., Martins, D.R., Wilson, M.E., Pearson, R.D., Reed, S.G.,
Laboratory-based diagnosis of brucellosis–a review of the literature part I: Jeronimo, S.M., 2002. The sensitivity and specificity of Leishmania chagasi
techniques for direct detection and identification of Brucella spp. Clin. Lab. 49, recombinant K39 antigen in the diagnosis of American visceral leishmaniasis
487–505. and in differentiating active from subclinical infection. Am. J. Trop. Med. Hyg.
67, 344–348.
Al Dahouk, S., Tomaso, H., Nockler, K., Neubauer, H., Frangoulidis, D., 2003b. Bricker, B.J., 1999. Differentiation of hard-to-type bacterial strains by RNA
Laboratory-based diagnosis of brucellosis–a review of the literature part II: mismatch cleavage. BioTechniques 27, 321–324, 326.
serological tests for brucellosis. Clin. Lab. 49, 577–589. Bricker, B.J., 2002. PCR as a diagnostic tool for brucellosis. Vet. Microbiol. 90,
Al Dahouk, S., Tomaso, H., Prenger-Berninghoff, E., Splettstoesser, W.D., Scholz, Bricker, B.J., Ewalt, D.R., Halling, S.M., 2003. Brucella ‘HOOF-Prints’: strain typing by
H.C., Neubauer, H., 2005. Identification of brucella species and biotypes using multi-locus analysis of variable number tandem repeats (VNTRs). BMC
polymerase chain reaction- restriction fragment length polymorphism Microbiol. 3, 15.
(PCR-RFLP). Crit. Rev. Microbiol. 31, 191–196. Bricker, B.J., Halling, S.M., 1994. Differentiation of Brucella abortus bv. 1, 2, and 4,
Brucella melitensis, Brucella ovis, and Brucella suis bv. 1 by PCR. J. Clin. Microbiol.
Alaeddini, R., 2012. Forensic implications of PCR inhibition–a review: forensic 32, 2660–2666.
science international. Genetics 6, 297–305. Briggs, D.J., Smith, J.S., Mueller, F.L., Schwenke, J., Davis, R.D., Gordon, C.R.,
Schweitzer, K., Orciari, L.A., Yager, P.A., Rupprecht, C.E., 1998. A comparison of
Alon G.G., Jones L.M., Angus R.D., Verger J.M., 1988. Serological methods. two serological methods for detecting the immune response after rabies
Techniques for the Brucellosis laboratory. Institut National de la Recherche vaccination in dogs and cats being exported to rabies-free areas. Biologicals 26,
Agronomique. 347–355.
Brito, M.E., Mendonca, M.G., Gomes, Y.M., Jardim, M.L., Abath, F.G., 2000.
Alvarez, L., Fajardo, R., Lopez, E., Pedroza, R., Hemachudha, T., Kamolvarin, N., Identification of potentially diagnostic Leishmania braziliensis antigens in
Cortes, G., Baer, G.M., 1994. Partial recovery from rabies in a nine-year-old boy. human cutaneous leishmaniasis by immunoblot analysis. Clin. Diagn. Lab.
Pediatr. Infect. Dis. J. 13, 1154–1155. Immunol. 7, 318–321.
Brun, R., Blum, J., Chappuis, F., Burri, C., 2010. Human African trypanosomiasis.
Antinori, S., Calattini, S., Piolini, R., Longhi, E., Bestetti, G., Cascio, A., Parravicini, C., Lancet 375, 148–159.
Corbellino, M., 2009. Is real- time polymerase chain reaction (PCR) more useful Brunetti, E., Garcia, H.H., Junghanss, T., International Ce Workshop in Lima, P.,
than a conventional PCR for the clinical management of leishmaniasis? Am. J. 2011. Cystic echinococcosis: chronic, complex, and still neglected. PLoS Negl.
Trop. Med. Hyg. 81, 46–51. Trop. Dis. 5, e1146.
Brunetti, E., Junghanss, T., 2009. Update on cystic hydatid disease. Curr. Opin.
Antinori, S., Gianelli, E., Calattini, S., Longhi, E., Gramiccia, M., Corbellino, M., 2005. Infect. Dis. 22, 497–502.
Cutaneous leishmaniasis: an increasing threat for travellers. Clin. Microbiol. Brunetti, E., Kern, P., Vuitton, D.A., Writing, P., anel for, the, W.-I., 2010. Expert
Infect. 11, 343–346. consensus for the diagnosis and treatment of cystic and alveolar
echinococcosis in humans. Acta Trop. 114, 1–16.
Antwerpen, M.H., Zimmermann, P., Bewley, K., Frangoulidis, D., Meyer, H., 2008. Burans, J., Keleher, A., O’Brien, T., Hager, J., Plummer, A., Morgan, C., 1996. Rapid
Real-time PCR system targeting a chromosomal marker specific for Bacillus method for the diagnosis of Bacillus anthracis infection in clinical samples
anthracis. Mol. Cell. Probes 22, 313–315. using a hand-held assay. Salisbury Med. Bull. 87, 36–37.
Buscher, P., Mumba Ngoyi, D., Kabore, J., Lejon, V., Robays, J., Jamonneau, V.,
Arai, Y.T., Yamada, K., Kameoka, Y., Horimoto, T., Yamamoto, K., Yabe, S., Bebronne, N., Van der Veken, W., Bieler, S., 2009. Improved models of mini
Nakayama, M., Tashiro, M., 1997. Nucleoprotein gene analysis of fixed and anion exchange centrifugation technique (mAECT) and modified single
street rabies virus variants using RT- PCR. Arch. Virol. 142, 1787–1796. centrifugation (MSC) for sleeping sickness diagnosis and staging. PLoS Negl.
Trop. Dis. 3, e471.
Arechavala, A.I., Bianchi, M.H., Santiso, G.M., Lehmann, E.A., Walker, L., Negroni, R., Camara, M., Camara, O., Ilboudo, H., Sakande, H., Kabore, J., N’Dri, L., Jamonneau, V.,
2010. [Giemsa stain in the differential diagnosis of infectious diseases with Bucheton, B., 2010. Sleeping sickness diagnosis: use of buffy coats improves
cutaneous involvement]. Rev. Argent. Microbiol. 42, 316. the sensitivity of the mini anion exchange centrifugation test. Trop. Med. Int.
Health 15, 796–799.
Ariza, J., Pellicer, T., Pallares, R., Foz, A., Gudiol, F., 1992. Specific antibody profile in Caravano, R., Chabaud, F., Oberti, J., 1987. Application of immunoenzymatic
human brucellosis. Clin. Infect. Dis. 14, 131–140. techniques for epidemiological surveys on brucellosis among human
populations. Annales de l’Institut Pasteur. Microbiology 138, 79–84.
Attar, Z.J., Chance, M.L., el-Safi, S., Carney, J., Azazy, A., El-Hadi, M., Dourado, C., Cardenas, G., Jung, H., Rios, C., Fleury, A., Soto-Hernandez, J.L., 2010. Severe
Hommel, M., 2001. Latex agglutination test for the detection of urinary cysticercal meningitis: clinical and imaging characteristics. Am. J. Trop. Med.
antigens in visceral leishmaniasis. Acta Trop. 78, 11–16. Hyg. 82, 121–125.
Carmena, D., Benito, A., Eraso, E., 2006. Antigens for the immunodiagnosis of
Auer, H., Stockl, C., Suhendra, S., Schneider, R., 2009. [Sensitivity and specificity of Echinococcus granulosus infection: an update. Acta Trop. 98, 74–86.
new commercial tests for the detection of specific Echinococcus antibodies]. Carpio, A., 2002. Neurocysticercosis: an update. Lancet Infect. Dis. 2, 751–762.
Wien. Klin. Wochenschr. 121 (Suppl. 3), 37–41. Cassidy, J.P., 2006. The pathogenesis and pathology of bovine tuberculosis with
insights from studies of tuberculosis in humans and laboratory animal models.
Ayele, W.Y., Neill, S.D., Zinsstag, J., Weiss, M.G., Pavlik, I., 2004. Bovine tuberculosis: Vet. Microbiol. 112, 151–161.
an old disease but a new threat to Africa. Int. J. Tuberc. Lung Dis. 8, 924–937. Castilho, T.M., Camargo, L.M., McMahon-Pratt, D., Shaw, J.J., Floeter-Winter, L.M.,
2008. A real-time polymerase chain reaction assay for the identification and
Barham, W.B., Church, P., Brown, J.E., Paparello, S., 1993. Misidentification of quantification of American Leishmania species on the basis of
Brucella species with use of rapid bacterial identification systems. Clin. Infect. glucose-6-phosphate dehydrogenase. Am. J. Trop. Med. Hyg. 78, 122–132.
Dis. 17, 1068–1069.

Barker, D.C., 1987. DNA diagnosis of human leishmaniasis. Parasitol. Today 3,

Barker, D.C., 1989. Molecular approaches to DNA diagnosis. Parasitology 99
(Suppl), S125–S146.

Barrett, M.P., Burchmore, R.J., Stich, A., Lazzari, J.O., Frasch, A.C., Cazzulo, J.J.,
Krishna, S., 2003. The trypanosomiases. Lancet 362, 1469–1480.

Bern, C., Jha, S.N., Joshi, A.B., Thakur, G.D., Bista, M.B., 2000. Use of the recombinant
K39 dipstick test and the direct agglutination test in a setting endemic for
visceral leishmaniasis in Nepal. Am. J. Trop. Med. Hyg. 63, 153–157.

Best, M., Sattar, S.A., Springthorpe, V.S., Kennedy, M.E., 1990. Efficacies of selected
disinfectants against Mycobacterium tuberculosis. J. Clin. Microbiol. 28,

Beyer, W., 2004. Vaccination strategies for anthrax prevention. Berl Münch
Tierärztl Wochenschr 117, 508–524.

Beyer, W., Bartling, C., Neubauer, H., 2003. Zum Stand der Nachweisverfahren für
Bacillus anthracis in klinischen und Umweltproben. Tierärztl Umschau 58,

Bhattacharyya, T., Falconar, A.K., Luquetti, A.O., Costales, J.A., Grijalva, M.J., Lewis,
M.D., Messenger, L.A., Tran, T.T., Ramirez, J.D., Guhl, F., Carrasco, H.J., Diosque,
P., Garcia, L., Litvinov, S.V., Miles, M.A., 2014. Development of peptide-based

N.G. Schwarz et al. / Acta Tropica 165 (2017) 40–65 59

CDC, 1999. Centers for Disease Control and Prevention and National Institutes of Diaz, R., Moriyon, I., 1989. Laboratory techniques in the diagnosis of human
Health. Biosafety in Microbiological Laboratories, 4th ed. U.S. Government brucellosis. In: Young, E.J., Corbel, M.J. (Eds.), Brucellosis Clinical and
Printing Office, Washington, D.C. Laboratory Aspects. CRC Press, Florida, pp. 73–83.

CDC, 2014. Center of diseases control. Practical Guide for Specimen Collection and Diebold-Berger, S., Khan, H., Gottstein, B., Puget, E., Frossard, J.L., Remadi, S., 1997.
Reference Diagnosis of Leishmaniasis, May. Cytologic diagnosis of isolated pancreatic alveolar hydatid disease with
immunologic and PCR analyses. A case report. Acta Cytol. 41, 1381–1386.
Centers for Disease, C., Prevention, 1994. Brucellosis outbreak at a pork processing
plant–North Carolina, 1992. MMWR. Morbidity and mortality weekly report Diel, R., Goletti, D., Ferrara, G., Bothamley, G., Cirillo, D., Kampmann, B., Lange, C.,
43, 113-116. Losi, M., Markova, R., Migliori, G.B., Nienhaus, A., Ruhwald, M., Wagner, D.,
Zellweger, J.P., Huitric, E., Sandgren, A., Manissero, D., 2011. Interferon- gamma
Chappuis, F., Loutan, L., Simarro, P., Lejon, V., Buscher, P., 2005. Options for field release assays for the diagnosis of latent Mycobacterium tuberculosis infection:
diagnosis of human african trypanosomiasis. Clin. Microbiol. Rev. 18, 133–146. a systematic review and meta-analysis. Eur. Respir. J. 37, 88–99.

Chappuis, F., Rijal, S., Jha, U.K., Desjeux, P., Karki, B.M., Koirala, S., Loutan, L., Diel, R., Loddenkemper, R., Meywald-Walter, K., Gottschalk, R., Nienhaus, A., 2009.
Boelaert, M., 2006a. Field validity, reproducibility and feasibility of diagnostic Comparative performance of tuberculin skin test, QuantiFERON-TB-Gold In
tests for visceral leishmaniasis in rural Nepal. Trop. Med. Int. Health 11, 31–40. Tube assay, and T-Spot.TB test in contact investigations for tuberculosis. Chest
135, 1010–1018.
Chappuis, F., Rijal, S., Soto, A., Menten, J., Boelaert, M., 2006b. A meta- analysis of
the diagnostic performance of the direct agglutination test and rK39 dipstick Diel, R., Loddenkemper, R., Nienhaus, A., 2010. Evidence- based comparison of
for visceral leishmaniasis. BMJ 333, 723. commercial interferon-gamma release assays for detecting active TB: a
metaanalysis. Chest 137, 952–968.
Claus, D., 1992. A standardized gram staining procedure. World J. Microbiol.
Biotechnol. 8, 451–452. Diez, M., Favaloro, L., Bertolotti, A., Burgos, J.M., Vigliano, C., Lastra, M.P., Levin, M.J.,
Arnedo, A., Nagel, C., Schijman, A.G., Favaloro, R.R., 2007. Usefulness of PCR
Cliquet, F., Aubert, M., Sagne, L., 1998. Development of a fluorescent antibody virus strategies for early diagnosis of Chagas’ disease reactivation and treatment
neutralisation test (FAVN test) for the quantitation of rabies-neutralising follow-up in heart transplantation. Am. J. Transplant. 7, 1633–1640.
antibody. J. Immunol. Methods 212, 79–87.
Dinhopl, N., Mostegl, M.M., Richter, B., Nedorost, N., Maderner, A., Fragner, K.,
Cloeckaert, A., Verger, J.M., Grayon, M., Paquet, J.Y., Garin-Bastuji, B., Foster, G., Weissenbock, H., 2011. In situ hybridisation for the detection of Leishmania
Godfroid, J., 2001. Classification of Brucella spp. isolated from marine mammals species in paraffin wax-embedded canine tissues using a digoxigenin-labelled
by DNA polymorphism at the omp2 locus. Microbes Infect./Institut Pasteur 3, oligonucleotide probe. Vet. Rec. 169, 525.
Dinnes, J., Deeks, J., Kunst, H., Gibson, A., Cummins, E., Waugh, N., Drobniewski, F.,
CLSI, 2008. Clinical and Laboratory Standards Institute. Laboratory Detection and Lalvani, A., 2007. A systematic review of rapid diagnostic tests for the detection
Identification of Mycobacteria; Approved Guideline. CLSI document M48-A. of tuberculosis infection. Health Technol. Assess. 11, 1–196.
Wayne–Clinical and Laboratory Standards Institute.
Disch, J., Pedras, M.J., Orsini, M., Pirmez, C., de Oliveira, M.C., Castro, M., Rabello, A.,
Coertse, J., Weyer, J., Nel, L.H., Markotter, W., 2010. Improved PCR methods for 2005. Leishmania (Viannia) subgenus kDNA amplification for the diagnosis of
detection of African rabies and rabies-related lyssaviruses. J. Clin. Microbiol. mucosal leishmaniasis. Diagnostic microbiology and infectious disease 51,
48, 3949–3955. 185–190.

Cosivi, O., Grange, J.M., Daborn, C.J., Raviglione, M.C., Fujikura, T., Cousins, D., Doganay, M., Aydin, N., 1991. Antimicrobial susceptibility of Bacillus anthracis.
Robinson, R.A., Huchzermeyer, H.F., de Kantor, I., Meslin, F.X., 1998. Zoonotic Scand. J. Infect. Dis. 23, 333–335.
tuberculosis due to Mycobacterium bovis in developing countries. Emerg.
Infect. Dis. 4, 59–70. Dottorini, S., Sparvoli, M., Bellucci, C., Magnini, M., 1985. Echinococcus granulosus:
diagnosis of hydatid disease in man. Ann. Trop. Med. Parasitol. 79, 43–49.
Cota, G.F., de Sousa, M.R., Demarqui, F.N., Rabello, A., 2012. The diagnostic accuracy
of serologic and molecular methods for detecting visceral leishmaniasis in HIV Drancourt, M., Brouqui, P., Raoult, D., 1997. Afipia clevelandensis antibodies and
infected patients: meta-analysis. PLoS Negl. Trop. Dis. 6, e1665. cross-reactivity with Brucella spp. and Yersinia enterocolitica O:9. Clin. Diagn.
Lab. Immunol. 4, 748–752.
Craig, P.S., Giraudoux, P., Shi, D., Bartholomot, B., Barnish, G., Delattre, P., Quere,
J.P., Harraga, S., Bao, G., Wang, Y., Lu, F., Ito, A., Vuitton, D.A., 2000. An Dybwad, M., van der Laaken, A.L., Blatny, J.M., Paauw, A., 2013. Rapid identification
epidemiological and ecological study of human alveolar echinococcosis of Bacillus anthracis spores in suspicious powder samples by using
transmission in south Gansu, China. Acta Trop. 77, 167–177. matrix-assisted laser desorption ionization-time of flight mass spectrometry
(MALDI-TOF MS). Appl. Environ. Microbiol. 79, 5372–5383.
Craig, P.S., McManus, D.P., Lightowlers, M.W., Chabalgoity, J.A., Garcia, H.H.,
Gavidia, C.M., Gilman, R.H., Gonzalez, A.E., Lorca, M., Naquira, C., Nieto, A., Easterday, W.R., Van Ert, M.N., Zanecki, S., Keim, P., 2005. Specific detection of
Schantz, P.M., 2007. Prevention and control of cystic echinococcosis. Lancet bacillus anthracis using a TaqMan mismatch amplification mutation assay.
Infect. Dis. 7, 385–394. BioTechniques 38, 731–735.

Crepin, P., Audry, L., Rotivel, Y., Gacoin, A., Caroff, C., Bourhy, H., 1998. Intravitam Eckert, J., Deplazes, P., 2004. Biological, epidemiological, and clinical aspects of
diagnosis of human rabies by PCR using saliva and cerebrospinal fluid. J. Clin. echinococcosis, a zoonosis of increasing concern. Clin. Microbiol. Rev. 17,
Microbiol. 36, 1117–1121. 107–135.

Cupolillo, E., Medina-Acosta, E., Noyes, H., Momen, H., Grimaldi Jr, G., 2000. A el Tai, N.O., Osman, O.F., el Fari, M., Presber, W., Schonian, G., 2000. Genetic
revised classification for Leishmania and Endotrypanum. Parasitol. Today 16, heterogeneity of ribosomal internal transcribed spacer in clinical samples of
142–144. Leishmania donovani spotted on filter paper as revealed by single-strand
conformation polymorphisms and sequencing. Trans. R. Soc. Trop. Med. Hyg.
da Silva, L.A., de Sousa Cdos, S., da Graca, G.C., Porrozzi, R., Cupolillo, E., 2010. 94, 575–579.
Sequence analysis and PCR-RFLP profiling of the hsp70 gene as a valuable tool
for identifying Leishmania species associated with human leishmaniasis in Ellerbrok, H., Nattermann, H., Ozel, M., Beutin, L., Appel, B., Pauli, G., 2002. Rapid
Brazil. Infect. Genet. Evol. 10, 77–83. and sensitive identification of pathogenic and apathogenic Bacillus anthracis by
real-time PCR. FEMS Microbiol. Lett. 214, 51–59.
Dacheux, L., Reynes, J.M., Buchy, P., Sivuth, O., Diop, B.M., Rousset, D., Rathat, C.,
Jolly, N., Dufourcq, J.B., Nareth, C., Diop, S., Iehle, C., Rajerison, R., Sadorge, C., Evans, J.T., Smith, E.G., Banerjee, A., Smith, R.M., Dale, J., Innes, J.A., Hunt, D.,
Bourhy, H., 2008. A reliable diagnosis of human rabies based on analysis of skin Tweddell, A., Wood, A., Anderson, C., Hewinson, R.G., Smith, N.H., Hawkey,
biopsy specimens. Clin. Infect. Dis. 47, 1410–1417. P.M., Sonnenberg, P., 2007. Cluster of human tuberculosis caused by
Mycobacterium bovis: evidence for person-to-person transmission in the UK.
Dar, N.R., Khurshid, T., 2005. Comparison of skin smears and biopsy specimens for Lancet 369, 1270–1276.
demonstration of Leishmania tropica bodies in cutaneous leishmaniasis. J.
College Phys. Surg. Pak. 15, 765–767. Faber, W.R., Oskam, L., van Gool, T., Kroon, N.C., Knegt-Junk, K.J., Hofwegen, H., van
der Wal, A.C., Kager, P.A., 2003. Value of diagnostic techniques for cutaneous
De Brito, T., Araujo, M.D., Tiriba, A., 1973. Ultrastructure of the Negri body in leishmaniasis. J. Am. Acad. Dermatol. 49, 70–74.
human rabies. J. Neurol. Sci. 20, 363–372.
Fandinho, F.C., Orsi-Souza, A.T., Salem, J.I., 1990. A comparison of the Ziehl-Neelsen
de Jong, B.C., Onipede, A., Pym, A.S., Gagneux, S., Aga, R.S., DeRiemer, K., Small, P.M., and Kinyoun methods in staining smears from leprosy patients. Int. J. Leprosy
2005. Does resistance to pyrazinamide accurately indicate the presence of Other Mycobacterial Dis. 58, 389–391.
Mycobacterium bovis? J. Clin. Microbiol. 43, 3530–3532.
Felber, A., Graninger, W., 2013. Weakly positive tests and chronologic variation of
Deborggraeve, S., Buscher, P., 2012. Recent progress in molecular diagnosis of the QuantiFERON assay: a retrospective appraisal of usefulness. Tuberculosis
sleeping sickness. Exp. Rev. Mol. Diagn. 12, 719–730. 93, 647–653.

Deborggraeve, S., Coronado, X., Solari, A., Zulantay, I., Apt, W., Mertens, P., Laurent, Ferrara, G., Losi, M., D’Amico, R., Roversi, P., Piro, R., Meacci, M., Meccugni, B., Dori,
T., Leclipteux, T., Stessens, T., Dujardin, J.C., Herdewijn, P., Buscher, P., 2009. T. I.M., Andreani, A., Bergamini, B.M., Mussini, C., Rumpianesi, F., Fabbri, L.M.,
cruzi OligoC-TesT a simplified and standardized polymerase chain reaction Richeldi, L., 2006. Use in routine clinical practice of two commercial blood tests
format for diagnosis of Chagas disease. PLoS Negl. Trop Dis. 3, e450. for diagnosis of infection with Mycobacterium tuberculosis: a prospective study.
Lancet 367, 1328–1334.
Del Brutto, O.H., 2012. Diagnostic criteria for neurocysticercosis, revisited. Pathog.
Global Health 106, 299–304. Ferrer, E., Bonay, P., Foster-Cuevas, M., Gonzalez, L.M., Davila, I., Cortez, M.M.,
Harrison, L.J., Parkhouse, R.M., Garate, T., 2007. Molecular cloning and
Del Brutto, O.H., Rajshekhar, V., White, A.C.J., r, Tsang, V.C., Nash, T.E., Takayanagui, characterisation of Ts8B1, Ts8B2 and Ts8B3, three new members of the Taenia
O.M., Schantz, P.M., Evans, C.A., Flisser, A., Correa, D., Botero, D., Allan, J.C., Sarti, solium metacestode 8 kDa diagnostic antigen family. Mol. Biochem. Parasitol.
E., Gonzalez, A.E., Gilman, R.H., Garcia, H.H., 2001. Proposed diagnostic criteria 152, 90–100.
for neurocysticercosis. Neurology 57, 177–183.
Fishbein, D.B., Robinson, L.E., 1993. Rabies. N. Engl. J. Med. 329, 1632–1638.
Del Brutto, O.H., Wadia, N.H., Dumas, M., Cruz, M., Tsang, V.C., Schantz, P.M., 1996. Fooks, A.R., Johnson, N., Freuling, C.M., Wakeley, P.R., Banyard, A.C., McElhinney,
Proposal of diagnostic criteria for human cysticercosis and neurocysticercosis.
J. Neurol. Sci. 142, 1–6. L.M., Marston, D.A., Dastjerdi, A., Wright, E., Weiss, R.A., Muller, T., 2009.

Deniau, M., Canavate, C., Faraut-Gambarelli, F., Marty, P., 2003. The biological
diagnosis of leishmaniasis in HIV-infected patients. Ann. Trop. Med. Parasitol.
97 (Suppl. 1), 115–133.

60 N.G. Schwarz et al. / Acta Tropica 165 (2017) 40–65

Emerging technologies for the detection of rabies virus: challenges and hopes population with high prevalence of tuberculosis infection. Sarcoidosis
in the 21st century. PLoS Negl. Trop. Dis. 3, e530. Vasculitis Diffuse Lung Dis. 28, 95–101.
Force, L., Torres, J.M., Carrillo, A., Busca, J., 1992. Evaluation of eight serological Hamouda, T., Shih, A.Y., Baker Jr., J.R.J., 2002. A rapid staining technique for the
tests in the diagnosis of human echinococcosis and follow-up. Clin. Infect. Dis. detection of the initiation of germination of bacterial spores. Lett. Appl.
15, 473–480. Microbiol. 34, 86–90.
Foster, G., Osterman, B.S., Godfroid, J., Jacques, I., Cloeckaert, A., 2007. Brucella ceti Hancock, K., Pattabhi, S., Greene, R.M., Yushak, M.L., Williams, F., Khan, A., Priest,
sp. nov. and Brucella pinnipedialis sp. nov. for Brucella strains with cetaceans J.W., Levine, M.Z., Tsang, V.C., 2004. Characterization and cloning of GP50, a
and seals as their preferred hosts. Int. J. Syst. Evol. Microbiol. 57, 2688–2693. Taenia solium antigen diagnostic for cysticercosis. Mol. Biochem. Parasitol.
Foulet, F., Botterel, F., Buffet, P., Morizot, G., Rivollet, D., Deniau, M., Pratlong, F., 133, 115–124.
Costa, J.M., Bretagne, S., 2007. Detection and identification of Leishmania Hancock, K., Pattabhi, S., Whitfield, F.W., Yushak, M.L., Lane, W.S., Garcia, H.H.,
species from clinical specimens by using a real-time PCR assay and sequencing Gonzalez, A.E., Gilman, R.H., Tsang, V.C., 2006. Characterization and cloning of
of the cytochrome B gene. J. Clin. Microbiol. 45, 2110–2115. T24, a Taenia solium antigen diagnostic for cysticercosis. Mol. Biochem.
Fraga, J., Montalvo, A.M., De Doncker, S., Dujardin, J.C., Van der Auwera, G., 2010. Parasitol. 147, 109–117.
Phylogeny of Leishmania species based on the heat-shock protein 70 gene. Handa, U., Mohan, H., Ahal, S., Mukherjee, K.K., Dabra, A., Lehl, S.S., Yadav, T.D.,
Infect. Genet. Evol. 10, 238–245. 2001. Cytodiagnosis of hydatid disease presenting with Horner’s syndrome: a
Fraga, J., Veland, N., Montalvo, A.M., Praet, N., Boggild, A.K., Valencia, B.M., Arevalo, case report. Acta Cytol. 45, 784–788.
J., Llanos-Cuentas, A., Dujardin, J.C., Van der Auwera, G., 2012. Accurate and Hankins, D.G., Rosekrans, J.A., 2004. Overview, prevention, and treatment of rabies.
rapid species typing from cutaneous and mucocutaneous Leishmaniasis lesions Mayo Clin. Proc. 79, 671–676.
of the New World. Diagn. Microbiol. Infect. Dis. 74, 142–150. Haouas, N., Chargui, N., Chaker, E., Ben Said, M., Babba, H., Belhadj, S., Kallel, K.,
Franco, J.R., Simarro, P.P., Diarra, A., Jannin, J.G., 2014. Epidemiology of human Pratlong, F., Dedet, J.P., Mezhoud, H., Azaiez, R., 2005. Anthroponotic cutaneous
African trypanosomiasis. Clin. Epidemiol. 6, 257–275. Leishmaniasis in Tunisia: presence of Leishmania killicki outside its original
Frickmann, H., Alnamar, Y., Essig, A., Clos, J., Racz, P., Barth, T.F., Hagen, R.M., focus of Tataouine. Trans. R. Soc. Trop. Med. Hyg. 99, 499–501.
Fischer, M., Poppert, S., 2012. Rapid identification of Leishmania spp. in Harms, G., Feldmeier, H., 2005. The impact of HIV infection on tropical diseases.
formalin-fixed, paraffin-embedded tissue samples by fluorescence in situ Infect. Dis. Clin. N. Am. 19, 121–135, ix.
hybridization. Trop. Med. Int. Health 17, 1117–1126. Hasker, E., Mitashi, P., Baelmans, R., Lutumba, P., Jacquet, D., Lejon, V., Kande, V.,
Frickmann, H., Dobler, G., 2013. Inactivation of rickettsiae. Eur. J. Microbiol. Declercq, J., Van der Veken, W., Boelaert, M., 2010. A new format of the CATT
Immunol. 3, 188–193. test for the detection of human African Trypanosomiasis, designed for use in
Gadea, I., Ayala, G., Diago, M.T., Cunat, A., Garcia de Lomas, J., 2000. Immunological peripheral health facilities. Trop. Med. Int. Health 15, 263–267.
diagnosis of human hydatid cyst relapse: utility of the enzyme-linked Hattwick, M.A., 1972. Reactions to rabies. N Engl. J. Med. 287, 1204.
immunoelectrotransfer blot and discriminant analysis. Clin. Diagn. Lab. Hattwick, M.A., Weis, T.T., Stechschulte, C.J., Baer, G.M., Gregg, M.B., 1972. Recovery
Immunol. 7, 549–552. from rabies. A case report. Ann. Internal Med. 76, 931–942.
Gangneux, J.P., Menotti, J., Lorenzo, F., Sarfati, C., Blanche, H., Bui, H., Pratlong, F., Helbig, M., Frosch, P., Kern, P., Frosch, M., 1993. Serological differentiation between
Garin, Y.J., Derouin, F., 2003. Prospective value of PCR amplification and cystic and alveolar echinococcosis by use of recombinant larval antigens. J.
sequencing for diagnosis and typing of old world Leishmania infections in an Clin. Microbiol. 31, 3211–3215.
area of nonendemicity. J. Clin. Microbiol. 41, 1419–1422. Helgason, E., Okstad, O.A., Caugant, D.A., Johansen, H.A., Fouet, A., Mock, M., Hegna,
Garcia, A.L., Kindt, A., Quispe-Tintaya, K.W., Bermudez, H., Llanos, A., Arevalo, J., I., Kolsto, A.B., 2000. Bacillus anthracis, Bacillus cereus, and Bacillus
Banuls, A.L., De Doncker, S., Le Ray, D., Dujardin, J.C., 2005. American thuringiensis-one species on the basis of genetic evidence. Appl. Environ.
tegumentary leishmaniasis: antigen-gene polymorphism, taxonomy and Microbiol. 66, 2627–2630.
clinical pleomorphism. Infect. Genet. Evol. 5, 109–116. Hemachudha, T., Laothamatas, J., Rupprecht, C.E., 2002. Human rabies: a disease of
Garcia, H.H., Evans, C.A.W., Nash, T.E., Takayanagui, O.M., White, A.C., Botero, D., complex neuropathogenetic mechanisms and diagnostic challenges. Lancet
Rajshekhar, V., Tsang, V.C.W., Schantz, P.M., Allan, J.C., Flisser, A., Correa, D., Neurol. 1, 101–109.
Sarti, E., Friedland, J.S., Martinez, S.M., Gonzalez, A.E., Gilman, R.H., Del Brutto, Herrera, L., 2014. Trypanosoma cruzi, the causal agent of chagas disease:
O.H., 2002. Current consensus guidelines for treatment of neurocysticercosis. boundaries between wild and domestic cycles in Venezuela. Front. Public
Clin. Microbiol. Rev. 15, 747–756. Health 2 (259).
Garcia-Yoldi, D., Marin, C.M., de Miguel, M.J., Munoz, P.M., Vizmanos, J.L., Hira, P.R., Shweiki, H., Lindberg, L.G., Shaheen, Y., Francis, I., Leven, H., Behbehani,
Lopez-Goni, I., 2006. Multiplex PCR assay for the identification and K., 1988. Diagnosis of cystic hydatid disease: role of aspiration cytology. Lancet
differentiation of all Brucella species and the vaccine strains Brucella abortus 2, 655–657.
S19 and RB51 and Brucella melitensis Rev1. Clin. Chem. 52, 779–781. Hlavsa, M.C., Moonan, P.K., Cowan, L.S., Navin, T.R., Kammerer, J.S., Morlock, G.P.,
Gart, J.J., Buck, A.A., 1966. Comparison of a screening test and a reference test in Crawford, J.T., Lobue, P.A., 2008. Human tuberculosis due to Mycobacterium
epidemiologic studies. II. A probabilistic model for the comparison of bovis in the United States, 1995–2005. Clin. Infect. Dis. 47, 168–175.
diagnostic tests. Am. J. Epidemiol. 83, 593–602. Hoffmaster, A.R., Meyer, R.F., Bowen, M.D., Marston, C.K., Weyant, R.S., Thurman,
Gazozai, S., Iqbal, J., Bukhari, I., Bashir, S., 2010. Comparison of diagnostic methods K., Messenger, S.L., Minor, E.E., Winchell, J.M., Rassmussen, M.V., Newton, B.R.,
in cutaneous Leishmaniasis (histopathology compared to skin smears). Pak. J. Parker, J.T., Morrill, W.E., McKinney, N., Barnett, G.A., Sejvar, J.J., Jernigan, J.A.,
Pharm. Sci. 23, 363–366. Perkins, B.A., Popovic, T., 2002. Evaluation and validation of a real-time
Gee, J.E., De, B.K., Levett, P.N., Whitney, A.M., Novak, R.T., Popovic, T., 2004. Use of polymerase chain reaction assay for rapid identification of Bacillus anthracis.
16S rRNA gene sequencing for rapid confirmatory identification of Brucella Emerg. Infect. Dis. 8, 1178–1182.
isolates. J. Clin. Microbiol. 42, 3649–3654. Holani, A.G., Ganvir, S.M., Shah, N.N., Bansode, S.C., Shende, I., Jawade, R., Bijjargi,
Gelderblom, H., 1991. Negative staining in diagnostic virology. Micron Microscop S.C., 2014. Demonstration of mycobacterium tuberculosis in sputum and saliva
Acta 22, 435–447. smears of tuberculosis patients using ziehl neelsen and flurochrome staining-
Georges, S., Villard, O., Filisetti, D., Mathis, A., Marcellin, L., Hansmann, Y., Candolfi, a comparative study. J. Clin. Diagn. Res. 8, ZC42–ZC45.
E., 2004. Usefulness of PCR analysis for diagnosis of alveolar echinococcosis Holland, W.G., Claes, F., My, L.N., Thanh, N.G., Tam, P.T., Verloo, D., Buscher, P.,
with unusual localizations: two case studies. J. Clin. Microbiol. 42, 5954–5956. Goddeeris, B., Vercruysse, J., 2001. A comparative evaluation of parasitological
Godfroid, J., Cloeckaert, A., Liautard, J.P., Kohler, S., Fretin, D., Walravens, K., tests and a PCR for Trypanosoma evansi diagnosis in experimentally infected
Garin-Bastuji, B., Letesson, J.J., 2005. From the discovery of the Malta fever’s water buffaloes. Vet. Parasitol. 97, 23–33.
agent to the discovery of a marine mammal reservoir, brucellosis has Irenge, L.M., Gala, J.L., 2012. Rapid detection methods for Bacillus anthracis in
continuously been a re-emerging zoonosis. Vet. Res. 36, 313–326. environmental samples: a review. Appl. Microbiol. Biotechnol. 93, 1411–1422.
Gonzalez, L.M., Montero, E., Harrison, L.J., Parkhouse, R.M., Garate, T., 2000. Isenberg, H.D., 2004/2007. Clinical Microbiology Procedures Handbook.
Differential diagnosis of Taenia saginata and Taenia solium infection by PCR. J. Washington, D.C.
Clin. Microbiol. 38, 737–744. Islam, M.Z., Itoh, M., Takagi, H., Islam, A.U., Ekram, A.R., Rahman, A., Takesue, A.,
Gottstein, B., Jacquier, P., Bresson-Hadni, S., Eckert, J., 1993. Improved primary Hashiguchi, Y., Kimura, E., 2008. Enzyme-linked immunosorbent assay to
immunodiagnosis of alveolar echinococcosis in humans by an enzyme-linked detect urinary antibody against recombinant rKRP42 antigen made from
immunosorbent assay using the Em2plus antigen. J. Clin. Microbiol. 31, Leishmania donovani for the diagnosis of visceral leishmaniasis. Am. J. Trop.
373–376. Med. Hyg. 79, 599–604.
Gotuzzo, E., Carrillo, C., Guerra, J., Llosa, L., 1986. An evaluation of diagnostic Ismail, A., Kharazmi, A., Permin, H., el Hassan, A.M., 1997. Detection and
methods for brucellosis–the value of bone marrow culture. J. Infect. Dis. 153, characterization of Leishmania in tissues of patients with post kala-azar
122–125. dermal leishmaniasis using a specific monoclonal antibody. Trans. R. Soc. Trop.
Graca, G.C., Volpini, A.C., Romero, G.A., Oliveira Neto, M.P., Hueb, M., Porrozzi, R., Med. Hyg. 91, 283–285.
Boite, M.C., Cupolillo, E., 2012. Development and validation of PCR-based Ito, A., Ma, L., Schantz, P.M., Gottstein, B., Liu, Y.H., Chai, J.J., Abdel-Hafez, S.K.,
assays for diagnosis of American cutaneous Leishmaniasis and identification of Altintas, N., Joshi, D.D., Lightowlers, M.W., Pawlowski, Z.S., 1999. Differential
the parasite species. Memorias do Instituto Oswaldo Cruz 107, 664–674. serodiagnosis for cystic and alveolar echinococcosis using fractions of
Group, W.H.O.I.W., 2003. International classification of ultrasound images in cystic Echinococcus granulosus cyst fluid (antigen B) and E. multilocularis protoscolex
echinococcosis for application in clinical and field epidemiological settings. (EM18). Am. J. Trop. Med. Hyg. 60, 188–192.
Acta Trop. 85, 253–261. Ito, A., Sako, Y., Yamasaki, H., Mamuti, W., Nakaya, K., Nakao, M., Ishikawa, Y., 2003.
Gupta, D., Kumar, S., Aggarwal, A.N., Verma, I., Agarwal, R., 2011. Interferon gamma Development of Em18-immunoblot and Em18-ELISA for specific diagnosis of
release assay (QuantiFERON-TB Gold In Tube) in patients of sarcoidosis from a alveolar echinococcosis. Acta Trop. 85, 173–182.

N.G. Schwarz et al. / Acta Tropica 165 (2017) 40–65 61

Ito, A., Schantz, P.M., Wilson, J.F., 1995. Em18, a new serodiagnostic marker for Kohler, S., Foulongne, V., Ouahrani-Bettache, S., Bourg, G., Teyssier, J., Ramuz, M.,
differentiation of active and inactive cases of alveolar hydatid disease. Am. J. Liautard, J.P., 2002. The analysis of the intramacrophagic virulome of Brucella
Trop. Med. Hyg. 52, 41–44. suis deciphers the environment encountered by the pathogen inside the
macrophage host cell. Proc. Natl. Acad. Sci. U. S. A. 99, 15711–15716.
Ito, A., Wang, X.G., Liu, Y.H., 1993. Differential serodiagnosis of alveolar and cystic
hydatid disease in the People’s Republic of China. Am. J. Trop. Med. Hyg. 49, Kohler, S., Michaux-Charachon, S., Porte, F., Ramuz, M., Liautard, J.P., 2003. What is
208–213. the nature of the replicative niche of a stealthy bug named Brucella? Trends
Microbiol. 11, 215–219.
Ito, A., Xiao, N., Liance, M., Sato, M.O., Sako, Y., Mamuti, W., Ishikawa, Y., Nakao, M.,
Yamasaki, H., Nakaya, K., Bardonnet, K., Bresson-Hadni, S., Vuitton, D.A., 2002. Kolman, S., Maayan, M.C., Gotesman, G., Rozenszajn, L.A., Wolach, B., Lang, R., 1991.
Evaluation of an enzyme-linked immunosorbent assay (ELISA) with Comparison of the Bactec and lysis concentration methods for recovery of
affinity-purified Em18 and an ELISA with recombinant Em18 for differential Brucella species from clinical specimens. Eur. J. Clin. Microbiol. Infect. Dis. 10,
diagnosis of alveolar echinococcosis: results of a blind test. J. Clin. Microbiol. 647–648.
40, 4161–4165.
Kuzmin, I.V., Botvinkin, A.D., McElhinney, L.M., Smith, J.S., Orciari, L.A., Hughes, G.J.,
Jackson, A.C., Fenton, M.B., 2001. Human rabies and bat bites. Lancet 357, 1714. Fooks, A.R., Rupprecht, C.E., 2004. Molecular epidemiology of terrestrial rabies
Jackson, A.C., Ye, H., Ridaura-Sanz, C., Lopez-Corella, E., 2001. Quantitative study of in the former Soviet Union. J. Wildlife Dis. 40, 617–631.

the infection in brain neurons in human rabies. J. Med. Virol. 65, 614–618. Kuzmin, I.V., Hughes, G.J., Botvinkin, A.D., Orciari, L.A., Rupprecht, C.E., 2005.
Jacobs, T., Andra, J., Gaworski, I., Graefe, S., Mellenthin, K., Kromer, M., Halter, R., Phylogenetic relationships of Irkut and West Caucasian bat viruses within the
Lyssavirus genus and suggested quantitative criteria based on the N gene
Borlak, J., Clos, J., 2005. Complement C3 is required for the progression of sequence for lyssavirus genotype definition. Virus Res. 111, 28–43.
cutaneous lesions and neutrophil attraction in Leishmania major infection.
Med. Microbiol. Immunol. 194, 143–149. Kuzmin, I.V., Orciari, L.A., Arai, Y.T., Smith, J.S., Hanlon, C.A., Kameoka, Y.,
Jahans, K.L., Foster, G., Broughton, E.S., 1997. The characterisation of Brucella Rupprecht, C.E., 2003. Bat lyssaviruses (Aravan and Khujand) from Central
strains isolated from marine mammals. Vet. microbiol. 57, 373–382. Asia: phylogenetic relationships according to N, P and G gene sequences. Virus
Jamjoom, M.B., Ashford, R.W., Bates, P.A., Chance, M.L., Kemp, S.J., Watts, P.C., Res. 97, 65–79.
Noyes, H.A., 2004. Leishmania donovani is the only cause of visceral
leishmaniasis in East Africa; previous descriptions of L. infantum and L. Lachenbruch, P.A., 1998. Sensitivity, specificity, and vaccine efficacy. Control. Clin.
archibaldi from this region are a consequence of convergent evolution in the Trials 19, 569–574.
isoenzyme data. Parasitology 129, 399–409.
Jamonneau, V., Bucheton, B., Kabore, J., Ilboudo, H., Camara, O., Courtin, F., Solano, Lahaye, X., Vidy, A., Pomier, C., Obiang, L., Harper, F., Gaudin, Y., Blondel, D., 2009.
P., Kaba, D., Kambire, R., Lingue, K., Camara, M., Baelmans, R., Lejon, V., Buscher, Functional characterization of Negri bodies (NBs) in rabies virus-infected cells:
P., 2010. Revisiting the immune trypanolysis test to optimise epidemiological evidence that NBs are sites of viral transcription and replication. J. Virol. 83,
surveillance and control of sleeping sickness in West Africa. PLoS Negl. Trop. 7948–7958.
Dis. 4, e917.
Jenkins, D.J., Romig, T., Thompson, R.C., 2005. Emergence/re-emergence of Lasch, P., Beyer, W., Nattermann, H., Stammler, M., Siegbrecht, E., Grunow, R.,
Echinococcus spp.–a global update. Int. J. Parasitol. 35, 1205–1219. Naumann, D., 2009. Identification of Bacillus anthracis by using
Jiang, C.P., Don, M., Jones, M., 2005. Liver alveolar echinococcosis in China: clinical matrix-assisted laser desorption ionization-time of flight mass spectrometry
aspect with relative basic research. World J. Gastroenterol. 11, 4611–4617. and artificial neural networks. Appl. Environ. Microbiol. 75, 7229–7242.
Jiang, L., Wen, H., Ito, A., 2001. Immunodiagnostic differentiation of alveolar and
cystic echinococcosis using ELISA test with 18-kDa antigen extracted from Le Fleche, P., Jacques, I., Grayon, M., Al Dahouk, S., Bouchon, P., Denoeud, F., Nockler,
Echinococcus protoscoleces. Trans. R. Soc. Trop. Med. Hyg. 95, 285–288. K., Neubauer, H., Guilloteau, L.A., Vergnaud, G., 2006. Evaluation and selection
Jimenez, J.A., Rodriguez, S., Moyano, L.M., Castillo, Y., García, H.H., 2010. of tandem repeat loci for a Brucella MLVA typing assay. BMC Microbiol. 6, 9.
Differentiating Taenia eggs found in human stools: does Ziehl-Neelsen staining
help? Trop. Med. Int. Health. Lembo, T., Niezgoda, M., Velasco-Villa, A., Cleaveland, S., Ernest, E., Rupprecht, C.E.,
Jones, A.L., Brown, J.M., Mishra, V., Perry, J.D., Steigerwalt, A.G., Goodfellow, M., 2006. Evaluation of a direct, rapid immunohistochemical test for rabies
2004. Rhodococcus gordoniae sp. nov., an actinomycete isolated from clinical diagnosis. Emerging infectious diseases 12, 310–313.
material and phenol-contaminated soil. Int. J. Syst. Evol. Microbiol. 54,
407–411. Lescure, F.X., Le Loup, G., Freilij, H., Develoux, M., Paris, L., Brutus, L., Pialoux, G.,
Junghanss, T., da Silva, A.M., Horton, J., Chiodini, P.L., Brunetti, E., 2008. Clinical 2010. Chagas disease: changes in knowledge and management. Lancet Infect.
management of cystic echinococcosis: state of the art, problems, and Dis. 10, 556–570.
perspectives. Am. J. Trop. Med. Hyg. 79, 301–311.
Kamerbeek, J., Schouls, L., Kolk, A., van Agterveld, M., van Soolingen, D., Kuijper, S., Levine, M.Z., Calderon, J.C., Wilkins, P.P., Lane, W.S., Asara, J.M., Hancock, K.,
Bunschoten, A., Molhuizen, H., Shaw, R., Goyal, M., van Embden, J., 1997. Gonzalez, A.E., Garcia, H.H., Gilman, R.H., Tsang, V.C., 2004. Characterization,
Simultaneous detection and strain differentiation of Mycobacterium cloning, and expression of two diagnostic antigens for Taenia solium
tuberculosis for diagnosis and epidemiology. J. Clin. Microbiol. 35, 907–914. tapeworm infection. J. Parasitol. 90, 631–638.
Karesh, W.B., Dobson, A., Lloyd-Smith, J.O., Lubroth, J., Dixon, M.A., Bennett, M.,
Aldrich, S., Harrington, T., Formenty, P., Loh, E.H., Machalaba, C.C., Thomas, M.J., Liance, M., Janin, V., Bresson-Hadni, S., Vuitton, D.A., Houin, R., Piarroux, R., 2000.
Heymann, D.L., 2012. Ecology of zoonoses: natural and unnatural histories. Immunodiagnosis of Echinococcus infections: confirmatory testing and species
Lancet 380, 1936–1945. differentiation by a new commercial Western Blot. J. Clin. Microbiol. 38,
Karger, A., Melzer, F., Timke, M., Bettin, B., Kostrzewa, M., Nockler, K., Hohmann, A., 3718–3721.
Tomaso, H., Neubauer, H., Al Dahouk, S., 2013. Interlaboratory comparison of
intact-cell matrix-assisted laser desorption ionization- time of flight mass Ling, D.I., Flores, L.L., Riley, L.W., Pai, M., 2008. Commercial nucleic-acid
spectrometry results for identification and differentiation of Brucella spp. J. amplification tests for diagnosis of pulmonary tuberculosis in respiratory
Clin. Microbiol. 51, 3123–3126. specimens: meta- analysis and meta-regression. PLoS One 3, e1536.
Karimi, A., Alborzi, A., Rasooli, M., Kadivar, M.R., Nateghian, A.R., 2003. Prevalence
of antibody to Brucella species in butchers: slaughterers and others. East. Lista, F., Faggioni, G., Valjevac, S., Ciammaruconi, A., Vaissaire, J., le Doujet, C.,
Mediterranean Health J. 9, 178–184. Gorge, O., De Santis, R., Carattoli, A., Ciervo, A., Fasanella, A., Orsini, F.,
Kasai, H., Ezaki, T., Harayama, S., 2000. Differentiation of phylogenetically related D’Amelio, R., Pourcel, C., Cassone, A., Vergnaud, G., 2006. Genotyping of
slowly growing mycobacteria by their gyrB sequences. J. Clin. Microbiol. 38, Bacillus anthracis strains based on automated capillary 25-loci multiple locus
301–308. variable-number tandem repeats analysis. BMC Microbiol. 6, 33.
Kaur, J., Kaur, S., 2013. ELISA and western blotting for the detection of Hsp70 and
Hsp83 antigens of Leishmania donovani. J. Parasitic Dis. 37, 68–73. Live, I., 1958. Immunization studies on human volunteers with ether-killer
Kennedy, W.P., 1984. Novel identification of differences in the kinetoplast DNA of Brucella abortus: preliminary report. Bull. World Health Organ. 19, 197–199.
Leishmania isolates by recombinant DNA techniques and in situ hybridisation.
Mol. Biochem. Parasitol. 12, 313–325. Logan, N.A., Popovic, T., Hoffmaster, A., 2007. Gram- positive rods. Bacillus and
Kern, P., Ammon, A., Kron, M., Sinn, G., Sander, S., Petersen, L.R., Gaus, W., Kern, P., other aerobic endospore-forming bacteria. In: Murray, P.R., Baron, E.J.,
2004. Risk factors for alveolar echinococcosis in humans. Emerg. Infect. Dis. 10, Jorgensen, J.H., Landry, M.L., Pfaller, M.A. (Eds.), Manual Clin. Microbiol, 9th ed.
2088–2093. ASM Press), Washington, D.C.
Kern, P., Frosch, P., Helbig, M., Wechsler, J.G., Usadel, S., Beckh, K., Kunz, R., Lucius,
R., Frosch, M., 1995. Diagnosis of Echinococcus multilocularis infection by Lorenzo, C., Ferreira, H.B., Monteiro, K.M., Rosenzvit, M., Kamenetzky, L., Garcia,
reverse-transcription polymerase chain reaction. Gastroenterology 109, H.H., Vasquez, Y., Naquira, C., Sanchez, E., Lorca, M., Contreras, M., Last, J.A.,
596–600. Gonzalez-Sapienza, G.G., 2005. Comparative analysis of the diagnostic
Khuroo, M.S., 2002. Hydatid disease: current status and recent advances. Ann. performance of six major Echinococcus granulosus antigens assessed in a
Saudi Med. 22, 56–64. double-blind, randomized multicenter study. J. Clin. Microbiol. 43, 2764–2770.
Kirschner, P., Springer, B., Vogel, U., Meier, A., Wrede, A., Kiekenbeck, M., Bange,
F.C., Bottger, E.C., 1993. Genotypic identification of mycobacteria by nucleic Lucero, N.E., Bolpe, J.E., 1998. Buffered plate antigen test as a screening test for
acid sequence determination: report of a 2-year experience in a clinical diagnosis of human brucellosis. J. Clin. Microbiol. 36, 1425–1427.
laboratory. J. Clin. Microbiol. 31, 2882–2889.
Lumb, R., Bastian, I., Gilpin, C., Jelfs, P., Keehner, T., Sievers, A., Australian
Mycobacterium Reference Laboratory, N., 2007. Tuberculosis in Australia:
bacteriologically confirmed cases and drug resistance, 2005. Communicable
diseases intelligence quarterly report 31, 80-86.

M’Fadyean, J., 1903. A further note with regard to the staining reaction of anthrax
blood with methylene blue. J. Comp. Pathol. 16, 360–361.

Mack, U., Migliori, G.B., Sester, M., Rieder, H.L., Ehlers, S., Goletti, D., Bossink, A.,
Magdorf, K., Holscher, C., Kampmann, B., Arend, S.M., Detjen, A., Bothamley, G.,
Zellweger, J.P., Milburn, H., Diel, R., Ravn, P., Cobelens, F., Cardona, P.J., Kan, B.,
Solovic, I., Duarte, R., Cirillo, D.M., Lange, C., Tbnet, 2009. LTBI: latent
tuberculosis infection or lasting immune responses to M. tuberculosis? A TBNET
consensus statement. Eur. Respir. J. 33, 956–973.

Maddison, S.E., Slemenda, S.B., Schantz, P.M., Fried, J.A., Wilson, M., Tsang, V.C.,
1989. A specific diagnostic antigen of Echinococcus granulosus with an
apparent molecular weight of 8 kDA. Am. J. Trop. Med. Hyg. 40, 377–383.

62 N.G. Schwarz et al. / Acta Tropica 165 (2017) 40–65

Magill, A.J., 2005. Cutaneous leishmaniasis in the returning traveler. Infect. Dis. Muller, B., Durr, S., Alonso, S., Hattendorf, J., Laisse, C.J., Parsons, S.D., van Helden,
Clin. N. Am. 19, 241–266, x- xi. P.D., Zinsstag, J., 2013. Zoonotic Mycobacterium bovis-induced tuberculosis in
humans. Emerg. Infect. Dis. 19, 899–908.
Mahdi, M., Elamin, E.M., Melville, S.E., Musa, A.M., Blackwell, J.M., Mukhtar, M.M.,
Elhassan, A.M., Ibrahim, M.E., 2005. Sudanese mucosal leishmaniasis: isolation Mumba Ngoyi, D., Ali Ekangu, R., Mumvemba Kodi, M.F., Pyana, P.P., Balharbi, F.,
of a parasite within the Leishmania donovani complex that differs genotypically Decq, M., Kande Betu, V., Van der Veken, W., Sese, C., Menten, J., Buscher, P.,
from L. donovani causing classical visceral leishmaniasis. Infect. Genet. Evol. 5, Lejon, V., 2014. Performance of parasitological and molecular techniques for
29–33. the diagnosis and surveillance of gambiense sleeping sickness. PLoS Negl. Trop.
Dis. 8, e2954.
Mani, R.S., Madhusudana, S.N., 2013. Laboratory diagnosis of human rabies: recent
advances. ScientificWorldJournal 2013 (569712). Mumba Ngoyi, D., Menten, J., Pyana, P.P., Buscher, P., Lejon, V., 2013. Stage
determination in sleeping sickness: comparison of two cell counting and two
Manning S.E., Rupprecht C.E., Fishbein D., Hanlon C.A., Lumlertdacha B., Guerra M., parasite detection techniques. Trop. Med. Int. Health 18, 778–782.
Meltzer M.I., Dhankhar P., Vaidya S.A., Jenkins S.R., Sun B., Hull H.F., Advisory
Committee on Immunization Practices Centers for Disease, C., Prevention, Nadin-Davis, S.A., 1998. Polymerase chain reaction protocols for rabies virus
2008. Human rabies prevention–United States, 2008: recommendations of the discrimination. J. Virol. Methods 75, 1–8.
Advisory Committee on Immunization Practices. MMWR. Recommendations
and reports: Morbidity and mortality weekly report. Recommendations and Nadin-Davis, S.A., Casey, G.A., Wandeler, A.I., 1994. A molecular epidemiological
reports / Centers for Disease Control 57, 1-28. study of rabies virus in central Ontario and western Quebec. J. Gen. Virol. 75 (Pt
10), 2575–2583.
Marston, C.K., Gee, J.E., Popovic, T., Hoffmaster, A.R., 2006. Molecular approaches to
identify and differentiate Bacillus anthracis from phenotypically similar Nadin-Davis, S.A., Muldoon, F., Wandeler, A.I., 2006. A molecular epidemiological
Bacillus species isolates. BMC Microbiol. 6, 22. analysis of the incursion of the raccoon strain of rabies virus into Canada.
Epidemiol. Infect. 134, 534–547.
Matovu, E., Kazibwe, A.J., Mugasa, C.M., Ndungu, J.M., Njiru, Z.K., 2012. Towards
Point-of-Care Diagnostic and Staging Tools for Human African Naha, K., Dasari, S., Pandit, V., Seshadri, S., 2012. A rare case of seronegative
Trypanosomiaisis. Journal of tropical medicine 2012 (340538). culture–proven infection with Brucella suis. Aust. Med. J. 5, 340–343.

Mayta, H., Gilman, R.H., Prendergast, E., Castillo, J.P., Tinoco, Y.O., Garcia, H.H., Navarro, E., Casao, M.A., Solera, J., 2004. Diagnosis of human brucellosis using PCR.
Gonzalez, A.E., Sterling, C.R., 2007. Nested PCR for Specific Diagnosis of Taenia Exp. Rev. Mol. Diagn. 4, 115–123.
solium Taeniasis. Journal of clinical microbiology 46, 286–289.
Ndao, M., Spithill, T.W., Caffrey, R., Li, H., Podust, V.N., Perichon, R., Santamaria, C.,
Mayta, H., Gilman, R.H., Prendergast, E., Castillo, J.P., Tinoco, Y.O., Garcia, H.H., Ache, A., Duncan, M., Powell, M.R., Ward, B.J., 2010. Identification of novel
Gonzalez, A.E., Sterling, C.R., Cysticercosis Working Group in, P., 2008. Nested diagnostic serum biomarkers for Chagas’ disease in asymptomatic subjects by
PCR for specific diagnosis of Taenia solium taeniasis. Journal of clinical mass spectrometric profiling. J. Clin. Microbiol. 48, 1139–1149.
microbiology 46, 286–289.
Niemann, S., Harmsen, D., Rusch-Gerdes, S., Richter, E., 2000. Differentiation of
Mbati, P.A., Abok, K., Anjili, C.O., Orago, A.S., Kagai, J.M., Githure, J.I., Koech, D.K., clinical Mycobacterium tuberculosis complex isolates by gyrB DNA sequence
1995. Comparison of Giemsa and acridine orange stains for the diagnosis of polymorphism analysis. J. Clin. Microbiol. 38, 3231–3234.
Leishmania donovani in biopsies of infected hamsters. African journal of health
sciences 2, 228–231. Nienhaus, A., Costa, J.T., 2013. Screening for tuberculosis and the use of a
borderline zone for the interpretation of the interferon-gamma release assay
McManus, D.P., Zhang, W., Li, J., Bartley, P.B., 2003. Echinococcosis. Lancet 362, (IGRA) in Portuguese healthcare workers. J. Occup. Med. Toxicol. 8, 1.
Njiru, Z.K., Mikosza, A.S., Matovu, E., Enyaru, J.C., Ouma, J.O., Kibona, S.N.,
Meissner, G., Schroder, K., 1969. [The so- called African mycobacteria strains from Thompson, R.C., Ndung’u, J.M., 2008. African trypanosomiasis: sensitive and
the tropical West Africa]. Zentralblatt fur Bakteriologie Parasitenkunde. rapid detection of the sub-genus Trypanozoon by loop-mediated isothermal
Infektionskrankheiten und Hygiene. 1. Abt. Medizinisch- hygienische amplification (LAMP) of parasite DNA. Int. J. Parasitol. 38, 589–599.
Bakteriologie, Virusforschung und Parasitologie. Originale 211, 69–81.
Nunes, M.C., Dones, W., Morillo, C.A., Encina, J.J., Ribeiro, A.L., Council on Chagas
Mendonca, M.G., de Brito, M.E., Rodrigues, E.H., Bandeira, V., Jardim, M.L., Abath, Disease of the Interamerican Society of, C., 2013. Chagas disease: an overview
F.G., 2004. Persistence of leishmania parasites in scars after clinical cure of of clinical and epidemiological aspects. J. Am. College Cardiol. 62, 767–776.
American cutaneous leishmaniasis: is there a sterile cure? The Journal of
infectious diseases 189, 1018–1023. Oliveira, J.G., Novais, F.O., de Oliveira, C.I., da Cruz Junior, A.C., Campos, L.F., da
Rocha, A.V., Boaventura, V., Noronha, A., Costa, J.M., Barral, A., 2005.
Menzies, D., 1999. Interpretation of repeated tuberculin tests. Boosting, Polymerase chain reaction (PCR) is highly sensitive for diagnosis of mucosal
conversion, and reversion. Am. J. Respir. Crit. Care Med. 159, 15–21. leishmaniasis. Acta Trop. 94, 55–59.

Meyer, K.F., Shaw, E.B., 1920. A comparison of the morphologic, culture and Olivier, M., Badaro, R., Medrano, F.J., Moreno, J., 2003. The pathogenesis of
biochemical characteristics of B. abortus and B. melitensis: studies on the genus Leishmania/HIV co-infection: cellular and immunological mechanisms. Ann.
Brucella nov. gen. Int. J. Infect. Dis. 27, 173–184. Trop. Med. Parasitol. 97 (Suppl. 1), 79–98.

Meymandi, S.S., Bahmanyar, M., Dabiri, S., Aflatonian, M.R., Bahmanyar, S., Orciari, L.A., Rupprecht, C.E., 2007. Rabies Virus. In: Murray, P.R., Baron, E.J.,
Meymandi, M.S., 2010. Comparison of cytologic giemsa and real-time Jorgensen, J.H., Landry, M.L., Pfaller, M.A. (Eds.), Manual of Clinical
polymerase chain reaction technique for the diagnosis of cutaneous Microbiology. , 9th ed. ASM Press, Washington, D.C, pp. 1470–1477.
leishmaniasis on scraping smears. Acta Cytol. 54, 539–545.
Ortona, E., Rigano, R., Margutti, P., Notargiacomo, S., Ioppolo, S., Vaccari, S., Barca,
Miguel Gomez, M.A., Bratos Perez, M.A., Martin Gil, F.J., Duenas Diez, A., Martin S., Buttari, B., Profumo, E., Teggi, A., Siracusano, A., 2000. Native and
Rodriguez, J.F., Gutierrez Rodriguez, P., Orduna Domingo, A., Rodriguez Torres, recombinant antigens in the immunodiagnosis of human cystic
A., 2003. Identification of species of Brucella using Fourier transform infrared echinococcosis. Parasite Immunol. 22, 553–559.
spectroscopy. J. Microbiol. Methods 55, 121–131.
Outhred, A.C., Jelfs, P., Suliman, B., Hill-Cawthorne, G.A., Crawford, A.B., Marais, B.J.,
Mitashi, P., Hasker, E., Lejon, V., Kande, V., Muyembe, J.J., Lutumba, P., Boelaert, M., Sintchenko, V., 2014. Added value of whole-genome sequencing for
2012. Human african trypanosomiasis diagnosis in first-line health services of management of highly drug-resistant TB. J. Antimicrob. Chemother.
endemic countries, a systematic review. PLoS Negl. Trop. Dis. 6, e1919.
Pai, M., Zwerling, A., Menzies, D., 2008. Systematic review: T-cell-based assays for
Mitscherlich, E., Marth, E.H., 1984. Microbial survival in the environment. Springer, the diagnosis of latent tuberculosis infection: an update. Ann. Internal Med.
New York, N.Y, pp. 232–266. 149, 177–184.

Mondal, S., Bhattacharya, P., Ali, N., 2010. Current diagnosis and treatment of Paiva, B.R., Passos, L.N., Falqueto, A., Malafronte Rdos, S., Andrade, H.F.J., r, 2004.
visceral leishmaniasis. Exp. Rev. Anti-infective Therapy 8, 919–944. Single step polymerase chain reaction (PCR) for the diagnosis of the
Leishmania (Viannia) subgenus. Revista do Instituto de Medicina Tropical de
Montalvo, A.M., Fraga, J., Maes, I., Dujardin, J.C., Van der Auwera, G., 2012. Three Sao Paulo 46, 335–338.
new sensitive and specific heat-shock protein 70 PCRs for global Leishmania
species identification. Eur. J. Clin. Microbiol. Infect. Dis. 31, 1453–1461. Pandey, K., Pandey, B.D., Mallik, A.K., Kaneko, O., Uemura, H., Kanbara, H., Yanagi,
T., Hirayama, K., 2010. Diagnosis of visceral leishmaniasis by polymerase chain
Moore, S.M., Hanlon, C.A., 2010. Rabies- specific antibodies: measuring surrogates reaction of DNA extracted from Giemsa’s solution-stained slides. Parasitol. Res.
of protection against a fatal disease. PLoS Negl. Trop. Dis. 4, e595. 107, 727–730.

Moreno, E., Moriyon, I., 2011. The genus Brucella. In: Dworkin, M. (Ed.), The Papadopoulos, M.C., Abel, P.M., Agranoff, D., Stich, A., Tarelli, E., Bell, B.A., Planche,
Prokaryotes: An Evolving Electronic Resource for the Microbiological T., Loosemore, A., Saadoun, S., Wilkins, P., Krishna, S., 2004. A novel and
Community. , 3rd ed. Springer-Verlag, New York, N.Y, release 3.7. accurate diagnostic test for human African trypanosomiasis. Lancet 363,
Moreno, E., Stackebrandt, E., Dorsch, M., Wolters, J., Busch, M., Mayer, H., 1990.
Brucella abortus 16S rRNA and lipid A reveal a phylogenetic relationship with Pappas, G., Papadimitriou, P., Akritidis, N., Christou, L., Tsianos, E.V., 2006. The new
members of the alpha-2 subdivision of the class Proteobacteria. J. Bacteriol. global map of human brucellosis. Lancet Infect. Dis. 6, 91–99.
172, 3569–3576.
Paramasivan, C.N., Narayana, A.S., Prabhakar, R., Rajagopal, M.S., Somasundaram,
Moro, P., Schantz, P.M., 2009. Echinococcosis: a review. Int. J. Infect. Dis. 13, P.R., Tripathy, S.P., 1983. Effect of storage of sputum specimens at room
125–133. temperature on smear and culture results. Tubercle 64,
Moro, P.L., Garcia, H.H., Gonzales, A.E., Bonilla, J.J., Verastegui, M., Gilman, R.H.,
2005. Screening for cystic echinococcosis in an endemic region of Peru using Pawłowski, Z.S., Eckert, J., Vuitton, D.A., Ammann, R.W., Kern, P., Craig, P.S., 2001.
portable ultrasonography and the enzyme-linked immunoelectrotransfer blot Echinococcosis in humans: clinical aspects, diagnosis and treatment. In: Eckert
(EITB) assay. Parasitol. Res. 96, 242–246. J, Gemmell MA, Meslin F-X, Pawłowski ZS, editors. WHO/OIE Manual on
Echinococcosis in Humans and Animals: a Public Health Problem of Global
Muddaiah, R.K., James, P.M., Lingegowda, R.K., 2013. Comparative study of smear Concern. WHO/OIE Ma. 2001. p. 20–72.
microscopy rapid slide culture, and Lowenstein–Jensen culture in cases of
pulmonary tuberculosis in a tertiary care hospital. J. Res. Med. Sci. 18, Perez-Cordero, J.J., Sanchez-Suarez, J., Delgado, G., 2011. Use of a fluorescent stain
767–771. for evaluating in vitro infection with Leishmania panamensis. Exp. Parasitol.
129, 31–35.

N.G. Schwarz et al. / Acta Tropica 165 (2017) 40–65 63

Perl, D.P., Good, P.F., 1991. The pathology of rabies in the central nervous system. Roelfsema, J.H., Nozari, N., Herremans, T., Kortbeek, L.M., Pinelli, E., 2011.
In: Baer, G.M. (Ed.), The Natural History of Rabies. , 2nd ed. CRC Press, Boca Evaluation and improvement of two PCR targets in molecular typing of clinical
Raton, Fla, pp. 163–190, 163–190. samples of Leishmania patients. Exp. Parasitol. 127, 36–41.

Peters, W., Pasvol, G., 2007. Atlas of Tropical Medicine and Parasitology, 6th ed. Rogan, W.J., Gladen, B., 1978. Estimating prevalence from the results of a screening
Elsevier Ltd. test. Am. J. Epidemiol. 107, 71–76.

Petti, C.A., Polage, C.R., Quinn, T.C., Ronald, A.R., Sande, M.A., 2006. Laboratory Romero, G.A., Guerra, M.V., Paes, M.G., Cupolillo, E., Bentin Toaldo, C., Macedo, V.O.,
medicine in Africa: a barrier to effective health care. Clin. Infect. Dis. 42, Fernandes, O., 2001. Sensitivity of the polymerase chain reaction for the
377–382. diagnosis of cutaneous leishmaniasis due to Leishmania (Viannia) guyanensis.
Acta Trop. 79, 225–229.
Pfyffer, G.E., 2007. Mycobacterium: general characteristics, laboratory detection,
and staining procedures. In: Murray, P.R., Baron, E.J., Jorgensen, J.H., Landry, Romig, T., Dinkel, A., Mackenstedt, U., 2006. The present situation of
M.L., Pfaller, M.A. (Eds.), Manual of Clinical Microbiology. , 9th ed. Washington, echinococcosis in Europe. Parasitology international 55 (Suppl), S187–S191.
D.C., ASM Press.
Rudd, R.J., Smith, J.S., Yager, P.A., Orciari, L.A., Trimarchi, C.V., 2005. A need for
Phillips, A.P., Martin, K.L., Broster, M.G., 1983. Differentiation between spores of standardized rabies-virus diagnostic procedures: effect of cover-glass
Bacillus anthracis and Bacillus cereus by a quantitative immunofluorescence mountant on the reliability of antigen detection by the fluorescent antibody
technique. J. Clin. Microbiol. 17, 41–47. test. Virus Res. 111, 83–88.

Pizzuto, M., Piazza, M., Senese, D., Scalamogna, C., Calattini, S., Corsico, L., Persico, Ruiz, J., Lorente, I., Perez, J., Simarro, E., Martinez-Campos, L., 1997. Diagnosis of
T., Adriani, B., Magni, C., Guaraldi, G., Gaiera, G., Ludovisi, A., Gramiccia, M., brucellosis by using blood cultures. J. Clin. Microbiol. 35, 2417–2418.
Galli, M., Moroni, M., Corbellino, M., Antinori, S., 2001. Role of PCR in diagnosis
and prognosis of visceral leishmaniasis in patients coinfected with human Rupprecht, C.E., Hanlon, C.A., Hemachudha, T., 2002. Rabies re-examined. Lancet
immunodeficiency virus type 1. J. Clin. Microbiol. 39, 357–361. Infect. Dis. 2, 327–343.

Poretti, D., Felleisen, E., Grimm, F., Pfister, M., Teuscher, F., Zuercher, C., Reichen, J., Rusch-Gerdes, S., Hillemann, D., 2008. [Modern laboratory diagnostics for
Gottstein, B., 1999. Differential immunodiagnosis between cystic hydatid mycobacteria]. Pneumologie 62, 533–540.
disease and other cross-reactive pathologies. Am. J. Trop. Med. Hyg. 60,
193–198. Saha, S., Mazumdar, T., Anam, K., Ravindran, R., Bairagi, B., Saha, B., Goswami, R.,
Pramanik, N., Guha, S.K., Kar, S., Banerjee, D., Ali, N., 2005. Leishmania
Porras, A.I., Yadon, Z.E., Altcheh, J., Britto, C., Chaves, G.C., Flevaud, L., Martins-Filho, promastigote membrane antigen-based enzyme-linked immunosorbent assay
O.A., Ribeiro, I., Schijman, A.G., Shikanai-Yasuda, M.A., Sosa-Estani, S., and immunoblotting for differential diagnosis of Indian post-kala-azar dermal
Stobbaerts, E., Zicker, F., 2015. Target product profile (TPP) for chagas disease leishmaniasis. J. Clin. Microbiol. 43, 1269–1277.
point-of-care diagnosis and assessment of response to treatment. PLoS Negl.
Trop. Dis. 9, e0003697. Salotra, P., Sreenivas, G., Pogue, G.P., Lee, N., Nakhasi, H.L., Ramesh, V., Negi, N.S.,
2001. Development of a species-specific PCR assay for detection of Leishmania
Praet, N., Verweij, J.J., Mwape, K.E., Phiri, I.K., Muma, J.B., Zulu, G., van Lieshout, L., donovani in clinical samples from patients with kala-azar and post-kala-azar
Rodriguez-Hidalgo, R., Benitez-Ortiz, W., Dorny, P., Gabriel, S., 2013. Bayesian dermal leishmaniasis. J. Clin. Microbiol. 39, 849–854.
modelling to estimate the test characteristics of coprology, coproantigen ELISA
and a novel real-time PCR for the diagnosis of taeniasis. Tropical medicine & Sanders, C.A., Nieda, R.R., Desmond, E.P., 2004. Validation of the use of
international health: TM & IH 18, 608–614. Middlebrook 7H10 agar, BACTEC MGIT 960, and BACTEC 460 12B media for
testing the susceptibility of Mycobacterium tuberculosis to levofloxacin. J. Clin.
Probert, W.S., Schrader, K.N., Khuong, N.Y., Bystrom, S.L., Graves, M.H., 2004. Microbiol. 42, 5225–5228.
Real-time multiplex PCR assay for detection of Brucella spp., B. abortus, and B.
melitensis. Journal of clinical microbiology 42, 1290–1293. Santamaria, C., Chatelain, E., Jackson, Y., Miao, Q., Ward, B.J., Chappuis, F., Ndao, M.,
2014. Serum biomarkers predictive of cure in chagas disease patients after
Qi, Y., Patra, G., Liang, X., Williams, L.E., Rose, S., Redkar, R.J., DelVecchio, V.G., 2001. nifurtimox treatment. BMC Infect. Dis. 14, 302.
Utilization of the rpoB gene as a specific chromosomal marker for real-time
PCR detection of Bacillus anthracis. Appl Environ Microbiol 67, 3720–3727. Sarciron, E.M., Bresson-Hadni, S., Mercier, M., Lawton, P., Duranton, C., Lenys, D.,
Petavy, A.F., Vuitton, D.A., 1997. Antibodies against Echinococcus multilocularis
Quammen, D., 2012. Spillover. Animal Infections and the Next Human Pandemic. alkaline phosphatase as markers for the specific diagnosis and the serological
W.W. Norton & Company, New York, USA. monitoring of alveolar echinococcosis. Parasite Immunol. 19, 61–68.

Queipo-Ortuno, M.I., Colmenero, J.D., Baeza, G., Morata, P., 2005. Comparison Sawadogo, T.L., Savadogo, L.G., Diande, S., Ouedraogo, F., Mourfou, A., Gueye, A.,
between lightcycler real-time polymerase chain reaction (PCR) assay with Sawadogo, I., Nebie, B., Sangare, L., Ouattara, A.S., 2012. [Comparison of
serum and PCR-enzyme-linked immunosorbent assay with whole blood Kinyoun, auramine O, and Ziehl-Neelsen staining for diagnosing tuberculosis
samples for the diagnosis of human brucellosis. Clin. Infect. dis. 40, 260–264. at the National Tuberculosis Center in Burkina Faso]. Medecine et sante
tropicales 22, 302–306.
Quinlan, P., Phelan, E., Doyle, M., 2014. Matrix- assisted laser desorption/ionisation
time-of-flight (MALDI-TOF) mass spectrometry (MS) for the identification of Scholz, H.C., Hubalek, Z., Sedlacek, I., Vergnaud, G., Tomaso, H., Al Dahouk, S.,
mycobacteria from MBBacT ALERT 3D liquid cultures and Lowenstein-Jensen Melzer, F., Kampfer, P., Neubauer, H., Cloeckaert, A., Maquart, M., Zygmunt,
(LJ) solid cultures. J. Clin. Pathol. M.S., Whatmore, A.M., Falsen, E., Bahn, P., Gollner, C., Pfeffer, M., Huber, B.,
Busse, H.J., Nockler, K., 2008. Brucella microti sp. nov., isolated from the
Raether, W., Hanel, H., 2003. Epidemiology, clinical manifestations and diagnosis common vole Microtus arvalis. Int. J. Syst. Evol. Microbiol. 58, 375–382.
of zoonotic cestode infections: an update. Parasitol. Res. 91, 412–438.
Schonian, G., Kuhls, K., Mauricio, I.L., 2011. Molecular approaches for a better
Rajasekariah, G.H., Ryan, J.R., Hillier, S.R., Yi, L.P., Stiteler, J.M., Cui, L., Smithyman, understanding of the epidemiology and population genetics of Leishmania.
A.M., Martin, S.K., 2001. Optimisation of an ELISA for the serodiagnosis of Parasitology 138, 405–425.
visceral leishmaniasis using in vitro derived promastigote antigens. J.
Immunol. Methods 252, 105–119. Schonian, G., Mauricio, I., Cupolillo, E., 2010. Is it time to revise the nomenclature
of Leishmania? Trends Parasitol. 26, 466–469.
Ramos, D.F., Tavares, L., da Silva, P.E., Dellagostin, O.A., 2014. Molecular typing of
Mycobacterium bovis isolates: a review. Braz. J. Microbiol. 45, 365–372. Schoone, G.J., van Eys, G.J., Ligthart, G.S., Taub, F.E., Zaal, J., Mebrahtu, Y., Laywer, P.,
1991. Detection and identification of Leishmania parasites by in situ
Rao, S.S., Mohan, K.V., Atreya, C.D., 2010. Detection technologies for Bacillus hybridization with total and recombinant DNA probes. Exp. Parasitol. 73,
anthracis: prospects and challenges. J. Microbiol. Methods 82, 1–10. 345–353.

Rassi Jr., A.J., Rassi, A., Marin-Neto, J.A., 2010. Chagas disease. Lancet 375, Schuch, R., Nelson, D., Fischetti, V.A., 2002. A bacteriolytic agent that detects and
1388–1402. kills Bacillus anthracis. Nature 418, 884–889.

Rausch, R.L., Wilson, J.F., Schantz, P.M., McMahon, B.J., 1987. Spontaneous death of Schur, L.F., Jacobson, R.L., 1988. Parasitological techniques. In: Peters
Echinococcus multilocularis: cases diagnosed serologically (by Em2 ELISA) and Killick-Kendrick, W.R. (Ed.), Leishmaniasis in Biology and Medicine, Vol. 1.
clinical significance. Am. J. Trop. Med. Hyg. 36, 576–585. Academic Press, New York, N.Y, pp. 500–541.

Redkar, R., Rose, S., Bricker, B., DelVecchio, V., 2001. Real-time detection of Brucella Scott, L., Albert, H., Gilpin, C., Alexander, H., DeGruy, K., Stevens, W., 2014.
abortus, Brucella melitensis and Brucella suis. Mol. Cell. Probes 15, 43–52. Multicenter feasibility study to assess external quality assessment panels for
Xpert MTB/RIF assay in South Africa. J. Clin. Microbiol. 52, 2493–2499.
Requena, J.M., Chicharro, C., Garcia, L., Parrado, R., Puerta, C.J., Canavate, C., 2012.
Sequence analysis of the 3’-untranslated region of HSP70 (type I) genes in the Sester, M., Sotgiu, G., Lange, C., Giehl, C., Girardi, E., Migliori, G.B., Bossink, A.,
genus Leishmania: its usefulness as a molecular marker for species Dheda, K., Diel, R., Dominguez, J., Lipman, M., Nemeth, J., Ravn, P., Winkler, S.,
identification. Parasites Vectors 5, 87. Huitric, E., Sandgren, A., Manissero, D., 2011. Interferon- gamma release assays
for the diagnosis of active tuberculosis: a systematic review and meta-analysis.
Ricciardi, A., Ndao, M., 2015. Diagnosis of parasitic infections: what’s going on? J. Eur. Respir. J. 37, 100–111.
Biomol. Screen. 20, 6–21.
Shepherd, J.C., McManus, D.P., 1987. Specific and cross-reactive antigens of
Richter, E., 2009. [Mykobakterien.] In Neumeister B, Geiss HK, Braun RW, Kimming Echinococcus granulosus hydatid cyst fluid. Mol. Biochem. Parasitol. 25,
(ed.). [Mikrobiologische Diagnostik.] Thieme Verlag, Stuttgart. 143–154.

Rickman, L.R., Robson, J., 1970. The testing of proven Trypanosoma brucei and T. Siles-Lucas, M.M., Gottstein, B.B., 2001. Molecular tools for the diagnosis of cystic
rhodesiense strains by the blood incubation infectivity test. Bulletin of the and alveolar echinococcosis. Trop. Med. Int. Health 6, 463–475.
World Health Organization 42, 911–916.
Silveira, F.T., Lainson, R., Corbett, C.E., 2004. Clinical and immunopathological
Rijal, S., Boelaert, M., Regmi, S., Karki, B.M., Jacquet, D., Singh, R., Chance, M.L., spectrum of American cutaneous leishmaniasis with special reference to the
Chappuis, F., Hommel, M., Desjeux, P., Van der Stuyft, P., Le Ray, D., Koirala, S., disease in Amazonian Brazil: a review. Memorias do Instituto Oswaldo Cruz
2004. Evaluation of a urinary antigen-based latex agglutination test in the 99, 239–251.
diagnosis of kala-azar in eastern Nepal. Trop. Med. Int. Health 9, 724–729.
Siracusano, A., Ioppolo, S., Notargiacomo, S., Ortona, E., Rigano, R., Teggi, A., De
Rodriguez, N., Guzman, B., Rodas, A., Takiff, H., Bloom, B.R., Convit, J., 1994. Rosa, F., Vicari, G., 1991. Detection of antibodies against Echinococcus
Diagnosis of cutaneous leishmaniasis and species discrimination of parasites granulosus major antigens and their subunits by immunoblotting. Trans. R.
by PCR and hybridization. J. Clin. Microbiol. 32, 2246–2252. Soc. Trop. Med. Hyg. 85, 239–243.

64 N.G. Schwarz et al. / Acta Tropica 165 (2017) 40–65

Smith, J.S., Orciari, L.A., Yaeger, P.A., 1995. Molecular epidemiology of rabies in the Tsang, V.C., Brand, J.A., Boyer, A.E., 1989. An enzyme- linked
United States. Semin. Virol., 387–400. immunoelectrotransfer blot assay and glycoprotein antigens for diagnosing
human cysticercosis (Taenia solium). J. Infect. Dis. 159, 50–59.
Smith, J.S., Yager, P.A., Baer, G.M., 1973. A rapid reproducible test for determining
rabies neutralizing antibody. Bull. World Health Organ. 48, 535–541. Turnbull, P.C., Hutson, R.A., Ward, M.J., Jones, M.N., Quinn, C.P., Finnie, N.J.,
Duggleby, C.J., Kramer, J.M., Melling, J., 1992. Bacillus anthracis but not always
Smits, H.L., Basahi, M.A., Diaz, R., Marrodan, T., Douglas, J.T., Rocha, A., Veerman, J., anthrax. J. Appl. Bacteriol. 72, 21–28.
Zheludkov, M.M., Witte, O.W., de Jong, J., Gussenhoven, G.C., Goris, M.G., van
Der Hoorn, M.A., 1999. Development and evaluation of a rapid dipstick assay Urdaneta-Morales, S., 2014. Chagas’ disease: an emergent urban zoonosis. The
for serodiagnosis of acute human brucellosis. J. Clin. Microbiol. 37, caracas valley (Venezuela) as an epidemiological model. Front. Public Health 2,
4179–4182. 265.

Sotelo, J., Marin, C., 1987. Hydrocephalus secondary to cysticercotic arachnoiditis. Valleca, V.M., Forester, F.T., 1981. The mouse inoculation test. In: Laboratory
A long-term follow-up review of 92 cases. J. Neurosurg. 66, 686–689. Methods for the Detecting Rabies. U.S. Government Printing Office,
Washington, D.C, pp. 91–100.
Soulsby, E.J.L., 1982. Helminths, Arthropods and Protozoa of Domesticated
Animals, 8 ed. Bailliere and Tyndall, London. van der Meide, W., Guerra, J., Schoone, G., Farenhorst, M., Coelho, L., Faber, W.,
Peekel, I., Schallig, H., 2008. Comparison between quantitative nucleic acid
Springer, B., Stockman, L., Teschner, K., Roberts, G.D., Bottger, E.C., 1996. Two- sequence-based amplification, real-time reverse transcriptase PCR, and
laboratory collaborative study on identification of mycobacteria: molecular real-time PCR for quantification of Leishmania parasites. J. Clin. Microbiol. 46,
versus phenotypic methods. J. Clin. Microbiol. 34, 296–303. 73–78.

Sreedevi, C., Hafeez, M., Kumar, P.A., Rayulu, V.C., Subramanyam, K.V., Sudhakar, K., Van Ert, M.N., Easterday, W.R., Huynh, L.Y., Okinaka, R.T., Hugh-Jones, M.E., Ravel,
2012. PCR test for detecting Taenia solium cysticercosis in pig carcasses. Trop. J., Zanecki, S.R., Pearson, T., Simonson, T.S., U’Ren, J.M., Kachur, S.M.,
Anim. Health Prod. 44, 95–99. Leadem-Dougherty, R.R., Rhoton, S.D., Zinser, G., Farlow, J., Coker, P.R., Smith,
K.L., Wang, B., Kenefic, L.J., Fraser-Liggett, C.M., Wagner, D.M., Keim, P., 2007.
Steingart, K.R., Flores, L.L., Dendukuri, N., Schiller, I., Laal, S., Ramsay, A., Hopewell, Global genetic population structure of Bacillus anthracis. PLoS One 2, e461.
P.C., Pai, M., 2011. Commercial serological tests for the diagnosis of active
pulmonary and extrapulmonary tuberculosis: an updated systematic review van Eys, G.J., Schoone, G.J., Ligthart, G.S., Laarman, J.J., Terpstra, W.J., 1987.
and meta-analysis. PLoS Med. 8, e1001062. Detection of Leishmania parasites by DNA in situ hybridization with
non-radioactive probes. Parasitol. Res. 73, 199–202.
Steingart, K.R., Henry, M., Laal, S., Hopewell, P.C., Ramsay, A., Menzies, D.,
Cunningham, J., Weldingh, K., Pai, M., 2007. A systematic review of commercial Verger, J.M., Grimont, F., Grimont, P.A.D., Grayon, M., 1985. Brucella, a
serological antibody detection tests for the diagnosis of extrapulmonary monospecific genus as shown by deoxyribonucleic acid hybridization. Int. J.
tuberculosis. Postgrad. Med. J. 83, 705–712. Syst. Evol. Bacteriol. 35, 292–295.

Sternberg, J.M., Gierlinski, M., Bieler, S., Ferguson, M.A., Ndung’u, J.M., 2014. Vershilova, P.A., 1961. The use of live vaccine for vaccination of human beings
Evaluation of the diagnostic accuracy of prototype rapid tests for human against brucellosis in the USSR. Bull. World Health Organ. 24, 85–89.
african trypanosomiasis. PLoS Negl. Trop. Dis. 8, e3373.
Vizcaino, N., Cloeckaert, A., Verger, J., Grayon, M., Fernandez-Lago, L., 2000. DNA
Stratilo, C.W., Lewis, C.T., Bryden, L., Mulvey, M.R., Bader, D., 2006. Single- polymorphism in the genus Brucella. Microbes Infect./Institut Pasteur 2,
nucleotide repeat analysis for subtyping Bacillus anthracis isolates. J. Clin. 1089–1100.
Microbiol. 44, 777–782.
Volpini, A.C., Marques, M.J., Lopes dos Santos, S., Machado-Coelho, G.L., Mayrink,
Sullivan, L., Fleming, J., Sastry, L., Mehlert, A., Wall, S.J., Ferguson, M.A., 2014. W., Romanha, A.J., 2006. Leishmania identification by PCR of Giemsa-stained
Identification of sVSG117 as an immunodiagnostic antigen and evaluation of a lesion imprint slides stored for up to 36 years. Clin. Microbiol. Infect. 12,
dual- antigen lateral flow test for the diagnosis of human African 815–818.
trypanosomiasis. PLoS Negl. Trop. Dis. 8, e2976.
Wang, Q., Qiu, J., Yang, W., Schantz, P.M., Raoul, F., Craig, P.S., Giraudoux, P.,
Sullivan, L., Wall, S.J., Carrington, M., Ferguson, M.A., 2013. Proteomic selection of Vuitton, D.A., 2006. Socioeconomic and behavior risk factors of human alveolar
immunodiagnostic antigens for human African trypanosomiasis and echinococcosis in Tibetan communities in Sichuan, People’s Republic of China.
generation of a prototype lateral flow immunodiagnostic device. PLoS Negl. Am. J. Trop. Med. Hyg. 74, 856–862.
Trop. Dis. 7, e2087.
Warrell, M.J., Warrell, D.A., 2004. Rabies and other lyssavirus diseases. Lancet 363,
Sundar, S., Pai, K., Kumar, R., Pathak-Tripathi, K., Gam, A.A., Ray, M., Kenney, R.T., 959–969.
2001. Resistance to treatment in Kala-azar: speciation of isolates from
northeast India. Am. J. Trop. Med. Hyg. 65, 193–196. Weber, D.J., Sickbert-Bennett, E., Gergen, M.F., Rutala, W.A., 2003. Efficacy of
selected hand hygiene agents used to remove Bacillus atrophaeus (a surrogate
Sundar, S., Rai, M., 2002. Laboratory diagnosis of visceral leishmaniasis. Clin. Diagn. of Bacillus anthracis) from contaminated hands. Jama 289, 1274–1277.
Lab. Immunol. 9, 951–958.
Webster, W.A., Casey, G.A., 1996. Virus isolation in neuroblastoma cell culture. In:
Sweeney, D.A., Hicks, C.W., Cui, X., Li, Y., Eichacker, P.Q., 2011. Anthrax infection. Meslin, F.X., Kaplan, M.M., Koprowski, H. (Eds.), Laboratory Techniques in
Am. J. Respir. Crit. Care Med. 184, 1333–1341. Rabies. , 4th ed. World Health Organization, Geneva, Switzerland, pp. 96–104.

Talmi-Frank, D., Nasereddin, A., Schnur, L.F., Schonian, G., Toz, S.O., Jaffe, C.L., Wekesa, C., Kirenga, B.J., Joloba, M.L., Bwanga, F., Katamba, A., Kamya, M.R., 2014.
Baneth, G., 2010. Detection and identification of old world Leishmania by high Chest X-ray vs. Xpert(R) MTB/RIF assay for the diagnosis of sputum
resolution melt analysis. PLoS Negl. Trop. Dis. 4, e581. smear-negative tuberculosis in Uganda. Int. J. Tuberculosis Lung Dis. 18,
Tappe, D., Gruner, B., Kern, P., Frosch, M., 2008. Evaluation of a commercial
Echinococcus Western blot assay for serological follow-up of patients with Welch, R.J., Anderson, B.L., Litwin, C.M., 2009. An evaluation of two commercially
alveolar echinococcosis. Clin. Vaccine Immunol. 15, 1633–1637. available ELISAs and one in-house reference laboratory ELISA for the
determination of human anti-rabies virus antibodies. J. Med. Microbiol. 58,
Tavares, C.A., Fernandes, A.P., Melo, M.N., 2003. Molecular diagnosis of 806–810.
leishmaniasis. Exp. Rev. Mol. Diagn. 3, 657–667.
Wellinghausen, N., Nockler, K., Sigge, A., Bartel, M., Essig, A., Poppert, S., 2006.
Thijsen, S.F., van Rossum, S.V., Arend, S., Koster, B., Machiels, A.M., Bossink, A.W., Rapid detection of Brucella spp. in blood cultures by fluorescence in situ
2008. The value of interferon gamma release assays for diagnosis infection hybridization. J. Clin. Microbiol. 44, 1828–1830.
with Mycobacterium tuberculosis during an annual screening of health care
workers. J. Occup. Environ. Med./Am. College Occup. Environ. Med. 50, Wen, H., Craig, P.S., 1994. Immunoglobulin G subclass responses in human cystic
1207–1208. and alveolar echinococcosis. Am. J. Trop. Med. Hyg. 51, 741–748.

Thompson, R.C., Lymbery, A.J., Constantine, C.C., 1995. Variation in Echinococcus: Whitby, J.E., Johnstone, P., Sillero-Zubiri, C., 1997. Rabies virus in the decomposed
towards a taxonomic revision of the genus. Advances in parasitology 35, brain of an Ethiopian wolf detected by nested reverse
145–176. transcription-polymerase chain reaction. J. Wildlife Dis. 33, 912–915.

Thoulouze, M.I., Lafage, M., Schachner, M., Hartmann, U., Cremer, H., Lafon, M., Whitfield, S.G., Fekadu, M., Shaddock, J.H., Niezgoda, M., Warner, C.K., Messenger,
1998. The neural cell adhesion molecule is a receptor for rabies virus. J. Virol. S.L., Rabies Working, G., 2001. A comparative study of the fluorescent antibody
72, 7181–7190. test for rabies diagnosis in fresh and formalin-fixed brain tissue specimens. J.
Virol. Methods 95, 145–151.
Tittsler, R.P., Sandholzer, L.A., 1936. The use of semi-solid Agar for the detection of
bacterial motility. J. Bacteriol. 31, 575–580. Whittaker, E., Gordon, A., Kampmann, B., 2008. Is IP-10 a better biomarker for
active and latent tuberculosis in children than IFNgamma? PLoS One 3, e3901.
Tomaso, H., Bartling, C., Al Dahouk, S., Hagen, R.M., Scholz, H.C., Beyer, W.,
Neubauer, H., 2006. Growth characteristics of Bacillus anthracis compared to WHO, 2007. World Health Organization. Use of Liquid TB Culture and Drug
other Bacillus spp on the selective nutrient media Anthrax Blood Agar and Suscebility Testing (DST) in Low and Medium Income Settings. Summary of the
Cereus Ident Agar. Syst. Appl. Microbiol. 29, 24–28. Expert Group Meeting on the use of liquid culture media, Geneva, 26 March
2007. of liquid tb culture summary
Tortoli, E., 2003. Impact of genotypic studies on mycobacterial taxonomy: the new report.pdf? ua=1 Last accessed 19. January 2015. 1- 100.
mycobacteria of the 1990. Clin. Microbiol. Rev. 16, 319–354.
WHO, 2012. Research priorities for Chagas disease, human African trypanosomiasis
Tortoli, E., Mantella, A., Mariottini, A., Mazzarelli, G., Pecile, P., Rogasi, P.G., and leishmaniasis. World Health Organ Tech Rep Ser 975, 1-100.
Sterrantino, G., Fantoni, E., Leoncini, F., 2006a. Successfully treated
spondylodiscitis due to a previously unreported mycobacterium. J. Med. WHO, 2013. WHO Expert Consultation on Rabies. Second report. World Health
Microbiol. 55, 119–121. Organ Tech Rep Ser 982, 1-139.

Tortoli, E., Mattei, R., Russo, C., Scarparo, C., 2006b. Mycobacterium lentiflavum, an WHO, 2014. SevenNeglected Endemic Zoonosis - some basic facts. http://www.
emerging pathogen? J. Infect. 52, e185–e187. zoonotic diseases/en/index.html
Zugegriffen am 19.12.2014.
Toz, S.O., Chang, K.P., Ozbel, Y., Alkan, M.Z., 2004. Diagnostic value of rK39 dipstick zoonotic diseases/en/ Zugegriffen am 27.07.2015.
in zoonotic visceral leishmaniasis in Turkey. J. Parasitol. 90,
1484–1486. WHO-TDR-Diagnostics-Evaluation-Expert-Panel, Banoo, S., Bell, D., Bossuyt, P.,
Herring, A., Mabey, D., Poole, F., Smith, P.G., Sriram, N., Wongsrichanalai, C.,

N.G. Schwarz et al. / Acta Tropica 165 (2017) 40–65 65

Linke, R., O’Brien, R., Perkins, M., Cunningham, J., Matsoso, P., Nathanson, C.M., Yamasaki, H., Allan, J.C., Sato, M.O., Nakao, M., Sako, Y., Nakaya, K., Qiu, D., Mamuti,
Olliaro, P., Peeling, R.W., Ramsay, A., 2010. Evaluation of diagnostic tests for W., Craig, P.S., Ito, A., 2004b. DNA differential diagnosis of taeniasis and
infectious diseases: general principles. Nat. Rev. Microbiol. 8, S17–S29. cysticercosis by multiplex PCR. J. Clin. Microbiol. 42, 548–553.
Wilkins, P.P., Allan, J.C., Verastegui, M., Acosta, M., Eason, A.G., Garcia, H.H.,
Gonzalez, A.E., Gilman, R.H., Tsang, V.C., 1999. Development of a serologic Yancey, L.S., Diaz-Marchan, P.J., White, A.C., 2005. Cysticercosis: recent advances in
assay to detect Taenia solium taeniasis. Am. J. Trop. Med. Hyg. 60, 199–204. diagnosis and management of neurocysticercosis. Curr. Infect. Dis. Rep. 7,
Winkler, W.G., Fashinell, T.R., Leffingwell, L., Howard, P., Conomy, P., 1973. 39–47.
Airborne rabies transmission in a laboratory worker. Jama 226, 1219–1221.
Wright, E., McNabb, S., Goddard, T., Horton, D.L., Lembo, T., Nel, L.H., Weiss, R.A., Young, E.J., 1995a. Brucellosis: current epidemiology, diagnosis, and management.
Cleaveland, S., Fooks, A.R., 2009. A robust lentiviral pseudotype neutralisation Curr. Clin. Topics Infect. Dis. 15, 115–128.
assay for in-field serosurveillance of rabies and lyssaviruses in Africa. Vaccine
27, 7178–7186. Young, E.J., 1995. An overview of human brucellosis. Clin. Infect. Dis. 21, 283–290,
Wunner, W.H., Briggs, D.J., 2010. Rabies in the 21 century. PLoS Negl. Trop. Dis. 4, quiz 290.
Yagupsky, P., 1994. Detection of Brucella melitensis by BACTEC NR660 blood Zelazny, A.M., Fedorko, D.P., Li, L., Neva, F.A., Fischer, S.H., 2005. Evaluation of 7SL
culture system. J. Clin. Microbiol. 32, 1899–1901. RNA gene sequences for the identification of Leishmania spp. Am. J. Trop. Med.
Yagupsky, P., Baron, E.J., 2005. Laboratory exposures to brucellae and implications Hyg. 72, 415–420.
for bioterrorism. Emerg. Infect. Dis. 11, 1180–1185.
Yamasaki, H., Allan, J.C., Sato, M.O., Nakao, M., Sako, Y., Nakaya, K., Qiu, D., Mamuti, Zhang, W., Li, J., McManus, D.P., 2003. Concepts in immunology and diagnosis of
W., Craig, P.S., Ito, A., 2004a. DNA differential diagnosis of taeniasis and hydatid disease. Clin. Microbiol. Rev. 16, 18–36.
cysticercosis by multiplex PCR. J. Clin. Microbiol. 42, 548–553.
Zhang, W., McManus, D.P., 2006. Recent advances in the immunology and
diagnosis of echinococcosis. FEMS Immunol. Med. Microbiol. 47, 24–41.

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