Taxonomy
The family Francisellaceae, a member of the
gamma subclass of proteobacteria, consists of
the single genus F rancisella. Francisella
tularensis, Francisella novicida, Francisella
philomiragia, and Francisella noatunensis as well as unclassified Francisella
spp. comprise the
genus. Several publications have reviewed the
history and descriptions of members of the
genus Francisella (48,
165, 166).
F. tularensis is the causative agent of tularemia, a zoonosis affecting a wide
range of animals
and humans. Three subspecies of F. tularensis,
tularensis (type A), holarctica (type B),
and mediasiatica, displaying >99.8%
identity in their 16S rRNA genes, have been described
(165). F. tularensis subsp. tularensis and
F. tularensissubsp. holarctica have been further
subdivided into distinct subpopulations using
molecular typing methods (117). F.
tularensis subsp. tularensis has been separated into three subclades,
termed A1a, A1b, and
A2, and within F. tularensis subsp. holarctica,
10 separate subclades have been identified
(117).
F. novicida and F. philomiragia, originally isolated from salty water,
are only infrequently
associated with human disease (74,
81, 92, 191). F. novicida was classifed as a species
distinct from F. tularensis on the basis of
serologic, virulence, and biochemical comparisons
(92, 133). Subsequently, similarities between F.
novicida andF. tularensis were identified
based on genome reassociation studies (75% DNA
relatedness) and 16S rRNA gene
sequences (>99.8% identity) (74,
165). Recent whole-genome comparisons provide
evidence that the evolutions of F. novicida and
F. tularensis occurred separately, consistent
with F. novicida and F. tularensisbeing
two separate species (93). Between F.
philomiragia and F. tularensis, genome reassociation studies show 39%
average DNA
relatedness and >98.3% identity in their 16S
rRNA gene sequences (57, 74).Francisella
noatunensis has been linked only to environmental sources being recovered from
a number
of saltwater fish species (27,
114, 131). DNA hybridization studies show a mean
reassociation value of 68% between F.
noatunensis and F. philomiragia, with the two species
displaying 99.3% identity in their 16S rRNA genes
(114).
Several additional Francisellaceae members,
isolated in culture, including Wolbachia
persica (165) and Francisellaspp. recovered from
seawater (143), have been described.
Additionally, a Francisella sp. was
isolated from the blood and cerebrospinal fluid (CSF) of
two different patients in the United States in
2005 and 2006 (90), while
another Francisella sp. was isolated from
the blood and urine of a patient in Spain (50).
Characterization data indicate that these Francisella
spp. are distinct from F. tularensis, F. n
ovicida, and F. philomiragia.
Putative Francisellaceae family members
have also been identified based on shared identities
among 16S rRNA gene sequences. These include Francisella-like
endosymbionts (FLEs)
in Amblyomma maculatum and multipleDermacentor
and Ornithodoros tick species
(160, 165). DNA from putative Francisellaceae family
members has also been detected in soil
and water samples (22).
Description of the Genus
The genus Francisella comprises tiny
gram-negative coccobacilli that can be distinguished
from similar genera by several features (Table 1). Members of the genus take up Gram’s
counterstain (safranin) poorly; they are strict
aerobes, weakly catalase positive, urease
negative, nonmotile, and non-spore-forming, and
they react with a limited number of
carbohydrates (Table 2). Only a few sugars
(glucose, maltose, sucrose, and glycerol) are
utilized by most members of the genus. Acid is
produced without gas. Unique cellular fatty
acids are associated with the genus (Table 1) (74, 165, 192). For most species, in vitro
growth is enhanced by sulfhydryl supplementation.
Cultured members of the genus share
>97% identity within their 16S rRNA genes (90,
114,165).
A
few key differences separate species of the Francisella genus (Table
2). F.
philomiragia
and F. novicida are more biochemically reactive than F.
tularensis. F.
philomiragia
is oxidase positive by Kovacs’s modification, whereas F.
novicida and F.
tularensis
are oxidase negative. F. novicida and F. philomiragia differ
from F. tularensis by
their
ability to grow independently of cysteine supplementation and by their
comparatively
large
cell size. Additionally, levels of virulence in mice differ, with <10 cells
of F.
tularensis
being required to kill laboratory mice compared to <100 for F.
novicida (133).
Nucleotide
differences within 16S rRNA genes also discriminate F. tularensis, F.
novicida,
and F. philomiragia (56).
Biochemical,
molecular, and virulence differences also differentiate F. tularensis subspecies.
Glycerol
fermentation and citrulline ureidase activity distinguish F.
tularensis
subsp. tularensis from F. tularensis subsp.holarctica
(Table 2) (127, 128). PCRbased
techniques
as well as 16S rRNA gene sequencing can also differentiate F.
tularensis
subsp. tularensis from F. tularensis subsp. holarctica
(56, 197). Levels of virulence
in
rabbits differ, with the 100% lethal dose of F. tularensis subsp. tularensis
being <10 cells
when
inoculated subcutaneously, compared to 109 cells for F.
tularensis
subsp. holarctica (127). F.
tularensis subsp. mediasiaticadiffers from both F.
tularensis
subsp. tularensis and F. tularensis subsp. holarctica
by its inability to utilize
glucose
and its moderate pathogenicity in rabbits (128, 165).
Epidemiology and Transmission
Francisellaceae
are widely distributed in the natural environment. Some species
are obligate
pathogens
of animals and humans (F. tularensis) (Table 3);
others are present in the
environment
(water) and are opportunistic pathogens of humans (F. novicida and F.
philomiragia)
(Table 3), while others have been associated only with animals (F. n
oatunensis).
F. tularensis isolates have been found only within the Northern Hemisphere
(Holarctic
region), whereas F. novicida and F. noatunensis have been
recovered from both
the Northern and Southern
Hemispheres (27, 165, 166, 194).
The
geographic distribution of F. tularensis varies throughout the Northern
Hemisphere
(128, 165, 166).
Infections caused by F. tularensis subsp. tularensis occur only
in North
America,
whereas F. tularensis subsp. holarctica has a much wider
distribution, causing
disease
in both the Old and New Worlds. F. tularensis subsp.mediasiatica has
been found
only
in regions of central Asia. The geographic distributions of F.
tularensis
subsp.holarctica and subsp. tularensis subclades
also vary. Subclades of F.
tularensis
subsp. holarctica within North America are distinct from
subclades throughout the
rest
of the Northern Hemisphere, excepting Scandinavia (188).
Within the United States, the
geographic
distributions of F. tularensis subsp. tularensis subclades differ.
A1a and A1b
predominate
in the eastern half of the United States, whereas A2 appears restricted to the
western
United States (91).
F.
novicida has been isolated primarily in North America, with occurrences of F.
novicida-like
organisms
in Australia and Thailand (94, 194).
Of the fewer than 20 known isolates of F.
philomiragia,
most have been from North America, with three single incidences reported
from
Central and Eastern Europe and Australia (60, 74,164, 191, 194). F.
noatunensis has
been
isolated from Scandinavia, Chile, and Japan (27, 114, 131).
F.
tularensis is associated with a wide range of hosts; more than 100 species of
wild animals,
birds,
and arthropods have been found naturally infected (24, 75, 80).
Habitats where the
Lagomorpha
(Sylvilagus[rabbits], Lepus [hares], and Oryctolagus [Old
World hares] genera)
and
the Rodentia (water voles, muskrats, lemmings, voles, and beavers) thrive are
believed
to
be important in maintaining enzootic foci (75, 80).
Biting arthropod vectors, primarily
tabanid
flies (Chrysops and Tabanus), ticks (Dermacentor and Amblyomma),
and mosquitoes
(Aedes
and Ochlerotatus in Sweden and Russia), are implicated in the
transmission of
tularemia
(46, 75, 79, 144). Human infections with F. tularensis are acquired usually
via
inhalation
of infective aerosols, ingestion of contaminated water, handling of sick or
dead
animals,
or ingestion of infected meat or from the bite of an infected arthropod. In
contrast
to F.
tularensis, F. novicida and F. philomiragia do not appear to
infect a wide range of hosts.
Their
primary natural habitat appears to be saltwater. Only a single isolation of F.
philomiragia
from an animal host (muskrat) has been documented (81).
Neither F.
novicida
nor F. philomiragiahas been shown to be transmitted to
humans by the bite of an
infective
arthropod, and arthropods infected with these species have not been identified.
Tularemia
has been reported from many countries of the Northern Hemisphere. Foci of
endemicity
have long been documented in Russia and Kazakhstan, as well as Finland and
Sweden
(166). Annual cases are usually reported from most countries in
Eastern Europe.
Cases
of tularemia are reported less frequently from Western Europe; however, several
outbreaks
of tularemia have occurred over the last decade in Spain (5, 108).
Other areas
reporting
outbreaks comprising many hundreds of cases in the last decade include Sweden
and
Kosovo (139, 150).
In
the United States, the incidence of tularemia has steadily declined to 100 to
200 cases
each
year since the 1940s, when several thousand cases of tularemia were reported
annually
(31).
All states except Hawaii have reported cases. A total of 1,368 human cases from
44
states
were reported to the Centers for Disease Control and Prevention (CDC) between 1990
and
2000, with four states, South Dakota, Arkansas, Missouri, and Oklahoma,
accounting for
the
majority (56%) of cases (31). Most cases in the United States are sporadic, with patients
acquiring
tularemia from tick or deerfly bites or from contact with infected animals
(e.g., by
skinning
rabbits) (31, 91). Inhalation of an infective aerosol during landscaping and
contact
with
infected cats are also important modes of transmission (53, 54, 91).
The majority of
tularemia
cases occur in the summer months. Outbreaks of tularemia occur rarely in the
United
States. An outbreak of pneumonic tularemia associated with landscaping
activities
occurred
on Martha ’s Vineyard in 2000 (54), and since that time,
Martha ’a Vineyard has
recorded
yearly cases (109). The most recent outbreak in the United States occurred in Utah
in
2007 and was attributed to deerfly bites (142).
Cultures recovered from human cases in
the
United States from 1996 to 2001 comprise equal proportions of F.
tularensis
subsp. tularensis and F. tularensis subsp.holarctica
(174).
Clinical Significance
F.
tularensis
Tularemia
is caused by F. tularensis. The disease has been known historically by a
number of
synonyms,
such as rabbit fever, deerfly fever, market men’s disease, the glandular type
of
tick
fever, Ohara ’s or yato-byo disease, and water rat trappers’ disease, attesting
to the
variety
of clinical presentations, the infectious agent’s ubiquitous presence in
nature, and the
means
by which humans may acquire the infection.
The
clinical spectrum of tularemia depends on the mode of transmission, the
virulence of the
infecting
strain, the immune status of the host, and timely diagnosis and treatment (197).
Tularemia
can be misdiagnosed since its symptoms are not unique; a sudden onset of
chills,
fever,
headache, and generalized malaise characterize the onset of illness. The
differential
diagnosis
includes a wide range of infectious diseases, such as cat scratch fever,
mycobacterial
infections, anthrax, brucellosis, legionellosis, and plague (197).
The incubation
period
averages 3 to 5 days.
Patients
may present with any one of the clinical forms of tularemia: ulceroglandular,
glandular,
oculoglandular, oropharyngeal, typhoidal, and pneumonic (197).
The most
common
form is ulceroglandular disease (45 to 80% of the reported cases), where the
portal
of
entry is via an infective arthropod bite or other inoculation through the skin
barrier.
Glandular
tularemia is similar to ulceroglandular disease but lacks the ulcerated site of
infection.
Oculoglandular tularemia occurs when the conjunctiva is the initial site of
infection,
usually
as a result of the mechanical transfer of organisms from an infectious source
to the
eye
by the fingers. Oropharyngeal tularemia occurs from ingestion of contaminated
water or
food
and is associated with pharyngeal lymphadenopathy. Pneumonic tularemia occurs
by
direct
inhalation of the organism and is considered the most severe form of the
disease.
Typhoidal
tularemia is the most difficult form to recognize because there is no identified
portal
of entry and localized signs are absent. If untreated, bacterial dissemination
from the
primary
sites of infection can lead to secondary clinical presentations, such as sepsis
and
meningitis.
The
severity of infection can range from mild and self-limiting to fatal and is
largely
dependent
on the infecting strain. Little to no tularemia-related mortality is reported
in
Europe
and Asia, where only F. tularensis subsp.holarctica causes
tularemia, while mortality
in
the United States ranges between 2.3% (those diagnosed by culture and serology)
and
9%
(culture-confirmed cases only) (52, 174).
Infections due to F.
tularensis
subsp.tularensis are considered to be more severe than
infections caused by F.
tularensis
subsp. holarctica. Molecular epidemiologic studies have
refined this picture and
demonstrate
that among culture-confirmed cases in the United States, human infections due
to
the A1b subclade of F. tularensis subsp. tularensis result in
significantly higher mortality
(24%)
than infections caused by F. tularensis subsp. tularensis subclades
A1a (4%) and A2
(0%)
or F. tularensis subsp. holarctica (7%) (91).
Comparisons of levels of virulence in mice
corroborate
the differences in virulence among F. tularensis subsp. tularensis clades
and also
between
F. tularensis subsp.tularensis clades and F. tularensis subsp.
holarctica (118).
F.
novicida and F.
philomiragia
Human
infections caused by F. novicida and F. philomiragia are rare and
considered
opportunistic
infections, infecting primarily patients with underlying immunocompromising
conditions.
Fewer than 20 cases of F. philomiragia infection have been d escribed
since the
discovery
of this species in 1974 (60, 74, 164, 191). All but one case have involved a host
with
an impaired physical barrier to infection (near d rowning) or an impaired
immunologic
defense
system (chronic granulomatous disease or myeloproliferative disease). In most
cases,
F. philomiragia was isolated from normally sterile sites: blood, bone
marrow,
cerebrospinal
fluid, and pericardial fluid. The drowning and water exposure cases were
associated
with saltwater and brackish water, in contrast to F. tularensis infections,
which
are
associated with freshwater sources. Fewer than 10 cases of F. novicida or
F. novicida-like
infections
have been reported (26, 35, 74, 94, 191, 194). Isolates were recovered from
blood,
lymph node tissue, and wounds.
Collection, Handling, Storage, and Transport of
Specimens
Personnel
handling diagnostic cultures of F. tularensis are at considerable risk
for infection.
Due
to the extremely low infectious dose for F. tularensis, tularemia has
been one of the
most
commonly reported laboratory-associated bacterial infections (132, 145).
Even though
the
use of biological safety cabinets and prophylactic antibiotic therapy (as well
as
vaccination,
where available) provides safeguards for laboratory workers, these precautions
have
not fully eliminated laboratory exposures or modified practices in the clinical
laboratory
to
minimize risks (155, 162). Manipulation of cultures presents the greatest risk to
laboratory
workers, due to the high concentration of organisms. Clinical specimens should
be
handled
under biosafety level 2 (BSL-2) conditions with universal precautions, with
work
being
done using BSL-3 practices as soon as a suspect F. tularensis is
isolated in culture
(44).
The greatest risk of laboratory-acquired tularemia is by aerosol inhalation
while
working
with cultures. Any work with clinical samples that generates aerosols should be
performed
using BSL-3 practices. Laboratories may want to consider developing policies
that
encourage
physicians to alert the laboratory if a diagnosis of tularemia is expected.
The
choice of specimen for diagnostic testing is generally dependent on the form of
clinical
illness:
ulceroglandular, glandular, oculoglandular, oropharyngeal, pneumonic, or
typhoidal
tularemia
(197). Whole blood is an acceptable specimen for all clinical forms of
tularemia,
although
this sample may be negative, particularly if the disease is in the early stages
of
progression.
Serum for antibody detection is a standard specimen taken for diagnosis of all
forms
of illness. A first specimen should be collected as early in the course of
infection as
possible,
followed by a second specimen taken in the convalescent period (at least 14
days
apart
and preferably 3 to 4 weeks after onset of symptoms). Pharyngeal swabs,
bronchial/tracheal
washes or aspirates, sputum, transthoracic lung aspirates, and pleural
fluid
are appropriate specimens for pneumonic, typhoidal, or oropharyngeal tularemia.
Swabs
of
visible lesions or affected areas have been used most commonly for
ulceroglandular and
oculoglandular
tularemia. Aspirates from lymph nodes or lesions can be used for diagnosis of
ulceroglandular,
glandular, and oropharyngeal tularemia. Necropsied materials from animals
that
are appropriate for testing include samples from visible abscesses as well as
samples
from
lymph node, lung, liver, and spleen tissues and bone marrow.
For
specimens to be tested by culture, it is important where possible to
decontaminate the
surface
area prior to specimen collection since contamination of the sample with normal
flora
could
interfere with the interpretation of culture results. To minimize loss in
viability,
specimens
should be delivered to the laboratory within 24 hours and preferably within 2
hours.
In general, if transport is >24 hours, specimens should be stored chilled (2
to 8°C) in
an
appropriate medium until processed in the laboratory. Freezing of samples,
unless in a
preservative
environment, such as tissue specimens or glycerol-containing solutions, is not
recommended
because of the lysis of live bacteria upon thawing. Swabs should be placed in
Amies
agar with charcoal, a commercial transport system designed for anaerobic and
aerobic
pathogens
(Becton, Dickinson and Company, Franklin Lakes, NJ). F. tularensis should
remain
viable
for 7 days at ambient room temperature when stored in Amies medium (82).
Stuart
medium,
designed for transporting gonococcal specimens, and saline are inadequate for
keeping
F. tularensis viable during transport (82).
For serum samples, separation from blood
should
take place as soon as possible, preferably within 24 hours. Sera may be stored
at 2 to
8°C
for up to 10 days. If testing is delayed for a long period, serum samples may
be frozen.
For
PCR, specimens should be collected in guanidine isothiocyanate-containing
buffer, which
preserves
F. tularensis DNA for up to 1 month (82).
Arthropods may be stored intact in 2%
NaCl
for culture analysis or in ethanol for molecular testing.
Direct Detection in Specimens
Fresh
clinical specimens (ulcer and wound swabs, tissues, and aspirates) where the
concentration
of organisms might be expected to be high can be directly examined by
microscopy
by Gram straining, direct fluorescent-antibody (DFA) binding, or
immunohistochemistry.
Under microscopic examination of Gram-stained
specimens,
Francisella cells (single and pleomorphic) appear tiny and counterstain
so faintly
with
safranin that they can easily be missed. Basic fuchsin counterstains F.
tularensis better
than
safranin. Due to the small size of F. tularensis, Gram staining of
clinical specimens is
usually
of little diagnostic value. DFA staining using a fluorescein
isothiocyanate-labeled
hyperimmune
rabbit polyclonal antibody directed against whole, killed F. tularensis cells
can
be
used to presumptively identify F. tularensis subsp. tularensis and
F. tularensis subsp.
holarctica
in clinical specimens (197).
This DFA reagent does not react well or at all with F.
novicida
or F. philomiragia. Immunohistochemical (IHC) staining
using a monoclonal
antibody
directed against the lipopolysaccharide (LPS) has been used successfully to
visualize
F. tularensis in formalin-fixed tissues (71).
Neither the DFA reagent nor the IHC
reagent
is commercially available. DFA testing of specimens is provided by many
reference
laboratories
in the United States.
Because
of the relative rarity of human tularemia, evaluation of molecular diagnostics
with
clinical
specimens has been challenging. PCR-based diagnostic methods have been used
most
commonly for diagnosis of ulceroglandular tularemia, the most prevalent
clinical form.
DNA
detection by conventional PCR directed at thetul4 gene (unique to Francisella
spp.) has
been
successfully and widely applied for diagnosis of ulceroglandular tularemia
(82, 167, 197).
The tul4 PCR assay displays a sensitivity of 75% when applied to wound
specimens
from patients with ulceroglandular tularemia and was shown to be more sensitive
than
culture (sensitivity of 62%) (82). More recently, real-time
PCR assays using the TaqMan
5′
nuclease assay have been applied to detect F. tularensis DNA in a
variety of clinical
specimens,
including ulcer specimens, aspirates, throat swabs, bronchial washes, and
pleural
fluid
(30, 186, 195). These assays target multiple Francisella genes
(ISFtu2element,
iglC, tul4, and fopA genes). A limitation of most PCR-based
diagnostics
for F.
tularensis is the inability to discriminate F. tularensis from F.
novicida. While this may
not
be significant for patient management, the correct identification of species
has both
epidemiologic
and public health value. Routine incorporation of real-time PCR into the
diagnosis
of human tularemia remains in need of further evaluations and standardization.
Detection
of F. tularensis in ectoparasites is based primarily on PCR methods (65, 203).
Of
note,
FLEs, present in a wide range of tick species, have been shown to cross-react
with
molecular
targets used for the detection of F. tularensis, leading to
false-positive results
(89, 160, 173).
If molecular assays are to be used for screening environmental samples
for F.
tularensis, it is important to evaluate the assays for cross-reactivity
with
other
Francisellaceae members present in these sample types (22, 143).
16S rRNA gene
sequencing
can also be helpful for discriminating F. tularensis from FLEs or
environmental
Francisella spp. (22, 89).
Isolation Procedures
F.
tularensis is slow growing and fastidious, requiring supplementation with
sulfhydryl
compounds
(cysteine, cystine, thiosulfate, and IsoVitaleX) to grow on artificial medium. F.
tularensis
grows on several media common to clinical laboratories, including
chocolate agar
(CA)
(Fig. 1A), buffered charcoal-yeast extract agar (BCYE), and Thayer-Martin
agar. It also
grows
in thioglycolate broth. When supplemented with 1 to 2% IsoVitaleX, general
bacteriological
media (tryptic soy broth and Mueller-Hinton broth) support the growth of F.
tularensis.
The organisms grow slowly (60-min generation time); good growth, therefore, is
obtained
by prolonged incubation (48 hours or longer). Suspect cultures should be
incubated
at
35 to 37°C aerobically and observed daily for up to 14 days; CO2 does not
impede growth.
A
specialized medium, cysteine heart agar supplemented with 9% chocolatized sheep
blood
(CHAB),
is often used for the growth of F. tularensis in reference laboratories
(197). F.
tularensis
grows well on CHAB, as this is a high-nutrient medium.
Additionally,F.
tularensis
displays distinctive colony morphology on CHAB5 which can aid in
identification
(Fig.
1). F. tularensis does not grow well or at all on general
bacteriologic media, such as
sheep
blood agar. Nutritionally enriched specimens, such as blood or tissue, provide
an
intrinsic
source of sulfhydryl compounds that may initially permit F. tularensis growth
on
general
bacteriological media. Upon subculture, the fastidious nature ofF.
tularensis will
become
evident as the exogenous compounds are depleted, leading to the loss of the
bacterium’s viability unless
the subculture is propagated on cysteine-supplemented medium.
Clinical
samples from normally sterile sites can be plated on nonselective agars that
support
the
growth of F. tularensis. Care should be taken not to permit laboratory
contamination
of F.
tularensis isolates by environmental bacteria such as Staphylococcus, as
these bacteria
rapidly
outcompete and inhibit the growth ofF. tularensis on nonselective media
(141).
Clinical
specimens obtained from nonsterile sites or autopsy and environmental sources
should
be plated on antibiotic-containing media. Incorporation of an antibiotic
supplement
(7.5-mg/liter
colistin, 2.5-mg/liter amphotericin, 0.5-mg/liter lincomycin, 4.0-mg/liter
trimethoprim,
and 10-mg/liter ampicillin) into the medium has been demonstrated to
prevent
other organisms from overwhelming F. tularensis (141).
Commercially available
antibiotic-containing
media that support the growth of F. tularensis include improved Thayer-
Martin
agar (Remel, Lenexa, KS), BCYE selective agar containing polymyxin B,
anisomycin,
and
vancomycin (Remel or Becton, Dickinson), and cysteine heart agar containing
antibiotics
(polymyxin
B and penicillin) (Remel).
Specimens
for culture should be taken on the basis of clinical presentation and before
administration
of antibiotics. Fresh clinical material that is likely to contain high
concentrations
of F. tularensis organisms, such as ulcer and wound specimens and
lymphoid
tissue
(liver, spleen, or affected lymph node tissue), is inoculated directly onto an
agar plate
by
using a sample-laden swab or bacteriological loop. Larger inocula are necessary
for the
recovery
of F. tularensis from specimens that contain a lower concentration of
the
organisms,
such as aspirates of pharyngeal washes, bronchial wash fluids, pleural fluids,
and
environmental
samples. Whole blood should be directly inoculated into blood culture bottles
(197),
with subsequent culture of blood culture-positive specimens on agar.
Identification
Because
of the rarity as well as the sporadic nature of most tularemia cases, the
organism is
often
not easily identified when it is cultured. The isolation of a very tiny
(individual cells may
be
difficult to discern), poorly counterstaining (by safranin), gram-negative
coccobacillus
(Fig.
2A) that produces 1- to 2-mm-diameter gray to gray-white colonies on
CA after 48
hours
(and scant to no growth on blood agar) should raise suspicion for F.
tularensis. F.
tularensis
is oxidase negative, weakly catalase positive, β-lactamase
positive, X and V factor
negative,
and urease negative. Oxidase, X/V factor, and urease testing can help
differentiate
F. tularensisfrom other similar gram-negative organisms,
including
Brucella spp., Haemophilus influenzae, and Acinetobacterspp.
(Table 1). If further
testing
according to the algorithms given in chapter 12 does
not rule out F. tularensisand the
laboratory
cannot do further confirmatory tests, the isolate should be sent to a reference
laboratory
that can confirm (or rule out) its identification as F. tularensis.
Proper biohazard
shipping
procedures must be followed (see chapter 12 in
this Manual). In the United States,
all
states have at least one reference laboratory that is part of the Laboratory
Response
Network
(LRN). These LRN laboratories are able to confirm the identification of
bacterial
The
colonial morphology of F. tularensis is most distinctive when it is
grown on CHAB. On
CHAB,
F. tularensisexhibits a prominent and unique opalescent sheen due to its
production of
H2S;
this sheen is less prominent in F. philomiragia than in F. tularensis
and is absent in
cultures
of other gram-negative organisms, such
asYersinia,
Brucella, Haemophilus, and Pasteurella spp. On CA, BCYE, and
Thayer-Martin
agar,
F. tularensis colonies have an entire margin; they are gray, smooth,
raised, and moist,
with
a butyrous consistency. F. novicidagrows more robustly than F.
tularensis and is
cysteine
independent (Table 2). Colonies of F. philomiragia on CA are >5 mm in
diameter,
with
an entire margin; they are white, smooth, raised, mucoid, and cysteine
independent.
Isolates
can be identified as F. tularensis using antigen or molecular detection
methods,
including
slide agglutination, DFA staining (Fig. 3),
PCR, or sequencing. The slide
agglutination
test identifies a suspect culture by mixing commercially available (Becton,
Dickinson
and Company) polyclonal rabbit anti-F. tularensisantibody with suspect
cultures;
the
polyclonal rabbit anti-F. tularensis antibody does not react well or at
all
with
F. novicida or F. philomiragia (197).
DFA staining can also be used to identify F.
tularensis
subsp. tularensisand F. tularensis subsp.
holarctica isolates (197). Care should be
taken
to ensure that prepared smears are not too thick, as this can interfere with
the
performance
and interpretation of the test. In general, antigen-based identification methods
work
optimally when fresh cultures (24 hours) are tested. If a culture older than 24
hours is
to
be tested by antigen detection methods, a fresh subculture should be prepared.
PCR
methods
targeting F. tularensis-specific genes can also be used for the identification
of
suspect
cultures (167, 186). As described earlier in this chapter, it is important to
consider
that
many F. tularensis PCR assays cross-react withF. novicida. Therefore,
it may be
important
to rule out F. novicida as the cause of infection, particularly in areas
where
tularemia
has not been previously reported. 16S rRNA gene sequencing can be useful if the
isolate
is not easily identified as F. tularensis (5, 35).
Universal 16S rRNA gene primers
or Francisella-specific
16S rRNA gene primers can be used (57).
Typing Systems
Once an isolate has been identified as a Francisella
sp., supplemental tests can be used for
additional characterization, including typing of
species, subspecies, and strain. Oxidase can
be used to differentiate F. philomiragia from
F. novicida and F. tularensis (Table 2). Glycerol
fermentation and citrulline ureidase activity
distinguish F. tularensis subsp. tularensis from F.
tularensis subsp. holarctica (Table 2); conventional assays for
these biochemical tests have
been described (157). The 96-well automated
MicroLog Micro Station system with GN2
microplates (Biolog Inc., Hayward, CA) can also be
used to assess the glycerol fermentation
of F. tularensis (197).
Genus, species, and subspecies can be typed by sequence analysis of
the 16S rRNA gene (56, 57).
PCR methods (both conventional and real-time) can type
isolates at the level of genus, species,
subspecies (F. tularensis subsp.
tularensis or F. tularensis subsp. holarctica), and
subclades (F.
tularensissubsp. tularensis subclades A1 and A2) (116,
197). Pulsed-field gel electrophoresis
can differentiate the threeF. tularensis subsp.
tularensis subclades, A1a, A1b, and A2 (91).
For discrimination of individual strains, a
multilocus variable-number tandem-repeat assay
(MLVA) for F. tularensis, based on 25
different repeats in the genome (83), has been
utilized.
Serologic Tests
Antibodies may be detected as early as 1 week
after the onset of symptoms (about 2 weeks
after infection). By 2 weeks after onset,
antibodies may be detected in 89 to 95.4% of
samples. Antibodies can persist for more than 10
years (25, 190). Immunoglobulin M (IgM),
IgA, and IgG antibodies may appear simultaneously
(178, 187). IgM antibodies can last for
many years; thus, their presence does not always
indicate early or recent infection
(25, 178).
Agglutination testing, either by the tube
agglutination (TA) or the microagglutination (MA)
method, is a standard serology test for
determining the presence of antibodies in tularemia
(25, 29, 197). Formalin-killed antigen (prepared from F.
tularensis subsp. tularensis strain
Schu S4) is commercially available from Becton,
Dickinson. Formalin-killed F.
tularensis antigen is also prepared within reference laboratories worldwide.
In the United
States, a single specimen with a TA titer of
≥1:160 or an MA titer of ≥1:128 is considered
positive. Formalin-killed F. tularensis whole-cell
antigen may display low-level crossreactivity
with Brucella antibodies (23,
125). No cross-reactivity of F. novicida or F.
philomiragia sera has been observed with F. tularensis-killed cells (133).
Enzyme-linked
immunosorbent assays (ELISAs) have been adopted
for use in the parts of Europe where
tularemia is endemic (25,
146, 159, 197). The LPS and/or outer membrane fraction remains
the primary ELISA antigen used in test
applications. Antigenic differences between F.
tularensis subsp. tularensis and subsp. holarctica have not
been identified for use in serology
assays. Thus, serology assays do not distinguish
the infecting subspecies. This is of most
importance in North America, where both F.
tularensis subsp.tularensis and F.
tularensis subsp. holarctica cause tularemia.
F. tularensis organisms are intracellular bacteria and are capable of eliciting
both humoral
and cell-mediated immunity (178).
The latter response has been known to remain strong 25
years after infection (49).
Host T cells retain proliferative responses to unique F.
tularensis membrane proteins, with concomitant increases in interferon and
interleukin-2
levels (49, 175,
176, 178). Tests for measuring the cell-mediated immune
response are
specialized and are not routinely used for
diagnosis of tularemia (47).
Antimicrobial Susceptibilities
F. tularensis infections are treatable with narrow-spectrum antibiotics. All Francisella
isolates
examined to date are β-lactamase positive, so
penicillins and cephalosporins are not effective
and should not be used to treat tularemia.
Antibiotics recommended for treatment and
prophylaxis include chloramphenicol,
ciprofloxacin, gentamicin, streptomycin, and
tetracycline. Antimicrobial susceptibility testing
(broth microdilution and Etest) of a large
collection of F. tularensis isolates
worldwide has demonstrated no antimicrobial resistance to
drugs used for treating tularemia (77,
177, 181, 182, 184). Some isolates of F.
tularensis subsp. holarctica from Europe and Russia are erythromycin
resistant.
Antimicrobial susceptibility testing of F.
tularensis is not usually performed in clinical
microbiology laboratories because of safety
concerns in working with this organism and
because resistance to antibiotics used for
clinical treatment of tularemia has never been
reported (179). The Clinical and
Laboratory Standards Institute (CLSI) has published
interpretative criteria and quality control limits
for broth microdilution of F. tularensisusing
Mueller-Hinton medium supplemented with 2%
IsoVitaleX (36, 37).
Evaluation, Interpretation, and Reporting of
Results
Serology is generally the most common method for
laboratory confirmation of F.
tularensis infection, due largely to the organism being slow growing and
fastidious.
Nonetheless, culture provides a conclusive
diagnosis of infection and whenever possible
should be attempted using appropriate biosafety
measures. Isolation of a very tiny gramnegative
bacterium that shows fastidious growth
characteristics and is oxidase negative,
weakly catalase positive, urease negative, X/V
factor negative, and β-lactamase positive
should be strongly suspected as F. tularensis and
referred to a reference laboratory.
Confirmation of F. tularensisinfections
includes (i) identification of a culture as F.
tularensis and/or (ii) a fourfold difference in titers in acute- and
convalescent-phase serum
samples, with one of the paired samples having a
positive titer. A positive test result for a
primary clinical specimen using antigen or
molecular detection methods, including DFA
staining, IHC staining, or PCR, provides only a
presumptive diagnosis of F. tularensis. A
single positive serum sample is also considered
presumptive for tularemia. For all cases
presumed to be tularemia, it is necessary to
verify that the patients’ symptoms are
compatible with tularemia and, in the case of a
single positive titer, that the patient has not
been previously vaccinated.
In the United States, F. tularensis is
classified as a select agent. To transfer, receive, or
possess F. tularensis,laboratories must be
registered both with the CDC and with the Animal
and Plant Health Inspection Service (APHIS) of the
U.S. Department of Agriculture. The
registration process includes a U.S. Department of
Justice investigation of all personnel
having access to select agents. Clinical
laboratories are exempt from the registration
requirement provided that within 7 calendar days
of identifying one of these agents, they
transfer it to a registered entity and/or destroy
the agent on site. Laboratories identifying an
organism as F. tularensisare required to
report this finding immediately to the CDC. Report
forms, contact information, laboratory
registration information, and pertinent citations of the
U.S. Federal Code may be found athttp://www.cdc.gov/od/sap.
BRUCELLA Back to top
Brucella spp. are common zoonoses among domestic animals and among
wildlife, including
novel species of marine mammals. Brucella spp.
also cause infections in humans and can
mimic other infectious and noninfectious diseases,
posing challenges to physicians in
reaching a diagnosis. The remittent/undulant fever
of brucellosis was first confused with
other diseases, such as malaria and typhoid fever,
and was called many synonyms pertaining
mainly to the geographic locations where the
disease occurred: Mediterranean fever, Malta
fever, Gibraltar fever, and Cyprus fever (198).
Over the last decade, there has been renewed
interest in this organism due to its inclusion in
the potential biological weapons lists of most
authorities (8, 64,
100, 136,202;www.who.int/csr/resources/publications/Brucellosis.pdf).
Taxonomy and Genome
Brucellaceae is a family of phylogenetically closely related free-living soil
organisms
composed of Brucella,Ochrobactrum, and Mycoplana
spp. The Brucellaceae are part of the
order Rhizobiales, which includes other
genera involved in human disease: Bartonella,
Afipia, Methylobacterium, and Roseomonas. (41,
61).
The taxonomy of Brucella spp. remains to be
clarified. Studies indicate that
terrestrial Brucella spp. are homogeneous
species harboring >90% interspecies homology by
DNA-DNA hybridization studies, identical 16S rRNA
gene sequences, and >98% sequence
homology by comparative genomics. Because of these
findings, a suggestion was made to
consider Brucella a monospecific genus and
the different species as biovars ofBrucella
melitensis (73).
The average size of the genome is 2.37 × 109 Da,
with a DNA G+C content of 58 to 59
mol%. Currently, the genus Brucella encompasses
nine recognized species, six terrestrial
and three marine (58, 193).
The six terrestrial Brucella species are B. melitensis (three
biovars) (preferred hosts are goats, sheep, and
camels), B. abortus (seven biovars) (cattle,
bison, and buffalo), B. suis (five biovars)
(swine and a range of wild animals),B.
canis (dogs), B. ovis (rams), and B. neotomae (desert and
wood rats). The three identified
marine species, B. delphini, B. pinnipediae, and
B. cetaceae, were recovered from marine
mammals (e.g., seals, whales, and dolphins) and
were found to differ phenotypically from
the six terrestrial species by their patterns of
substrate-mediated metabolic activity. Brucella
maris has been suggested as a name to encompass these marine isolates,
but to date this
has not been accepted (38).
Though preferred or predominant hosts are recognized
for Brucella spp., cross-infection of other
mammalian species, including humans, may occur
(41).
Description of the Genus
Brucella spp. are facultative, intracellular, small (0.5- to 1.5-μm),
gram-negative coccobacilli
that lack capsules, flagellae, endospores, or
native plasmids. They are aerobic (some prefer
CO2 for their growth), do not ferment sugars, and
are positive in a few oxidative metabolic
tests. Brucella spp. can grow on a wide
range of culture media, and colonies appear after 24
to 48 hours of incubation as mostly smooth colonies,
but rough variants can occur (4, 41).
Antigenic Components
Several antigenic determinants of Brucella,
related mainly to LPS and protein antigens, have
been characterized. The LPS is the major antigen
that dominates the antibody response. LPS
of rough strains is very similar to LPS of smooth
strains. Based on their O side chain, smooth
strains were reported to be composed of two
antigenic epitopes: A (B. abortus) and M (B.
melitensis). The smooth-strain LPS has been reported to be responsible for
observed crossreactions
in both the agglutination and complement fixation
tests between smooth species
of Brucella and Yersinia enterocolitica O:9,
Escherichia hermannii, Escherichia
coliO:157, Salmonella enterica serovar O:30, Stenotrophomonas
maltophilia, and Vibrio
cholerae O:1. Cross-reaction has been attributed to the similarities of the
O-specific side
chains of the LPS molecules of these organisms (45).
The characterized protein antigens include outer
and inner membrane, cytoplasmic, and
periplasmic antigens. Some are recognized by the
immune system during infection and are
potentially useful in diagnostic tests (41,66,
120). For example, Omp25 is an outer
membrane structural protein that is highly
conserved in all brucellae and is associated with
both LPS and peptidoglycan. In addition, some
proteins, such as ribosomal proteins (e.g.,
L7/L12) and fusion proteins, demonstrate a
protective effect against Brucella based on
antibody and cell-mediated responses (41,
126). These molecules may be useful in potential
vaccines.
Virulence Factors, Pathogenic Mechanisms, and
Immune
Response
The incubation period is variable but generally is
1 to 4 weeks. The intracelluar location and
survival of the organism contribute to its
virulence and pathogenesis. The exact
pathophysiologic aspects of infection remain to be
defined (201). Briefly, once the brucellae
enter the body by various routes, they are
encountered by polymorphonuclear and
mononuclear phagocytes, to which lectin
facilitates a ttachment. In the process, several
factors are involved in enabling a brucella to
enter a host, escape from phagocytic killing by
inhibiting the phagosome-lysosome fusion, and
evade the immune system, and they aid in
its survival and propagation within macrophages
and other cells. This is followed up by
brucellae being transferred through regional lymph
nodes into the circulatory system and
subsequently being seeded throughout the body,
with tropism for the reticuloendothelial
system, resulting in different clinical phases of
disease (69). Virulence determinants include
urease to avoid stomach stress through oral
passage (158) and Brucella-containing vacuoles
that enable escape from immune system recognition
and provide an acidic environment to
hamper antibiotic activity. ABrucella LPS
cell component (containing a poly N-formyl
perosamine O chain and a CuZn superoxide
dismutase) and outer membrane protein 25
(OMP 25) were reported to help the bacteria
survive within mononuclear phagocytes
(41, 55). Also, the uniqueness of Brucella LPS
lies in its being a poor inducer of gamma
interferon and tumor necrosis factor alpha, both
of which are essential for T-helper 1 (Th1)-
type-cell-mediated immunity for the elimination of
the organism (67). The overall
inflammatory process results in a slow degradation
ofBrucella cell wall components by the
polymorphonuclear leukocytes and can lead to
granuloma formation, which is more often
associated with B. abortus than B.
melitensis (69).
Protective immunity, though not long term, is
conferred by antibodies to LPS and T-cellmediated
macrophage activation, triggered by protein
antigens (41). A study showed a
significant increase in the levels of
interleukin-12 and gamma interferon in patients
with Brucella infection compared to levels
in controls, indicating that there is induction of
Th1-type cytokines during human brucellosis (1).
The immune response
against Brucella involves antigen-specific
T-cell activation, CD4+ Th lymphocytes, CD8+ T
cells, and humoral responses (67).
Lymphocytes are the main stimulant of the immune
response. The Th1 response stimulates IgG2a
production, which is involved mostly in
protection against intracellular pathogens through
cell-mediated immunity, and is critical for
the clearance of Brucella infection. The
Th2 response stimulates the production of IgG1 and
is mainly responsible for protection against
extracellular pathogens through the humoral
immune response (67, 201).
Recently, a study of B. abortus infection in rats showed that the
IgG2a response (indicative of a Th1 response) persisted
and dominated over the IgG1
response (88). However, the exact
nature of the immune response and protective factors
involved in this disease are still being
investigated, and the pathogenic mechanisms of
reinfection remain unknown.
Epidemiology and Transmission
Although Brucella can be killed by
pasteurization, exposure to UV light, acidity, or many
antiseptics and disinfectants, it can survive for
long periods under various conditions, e.g.,
10 weeks in soil, 11 weeks in aborted fetuses, 17
weeks in bovine stool, around 3 weeks in
milk and ice cream, and several months in fresh
cheese (41, 202). In terms of the total
numbers of infected cases, B. melitensis
dominates the world arena (especially in the
Mediterranean and Arabian Gulf countries). However,
B. abortus and B. suis supersede it in
certain geographic locations. B. canis has
also been reported to cause human diseases,
while B. ovis and B.neotomae
have not (41, 100, 137). Brucella spp. associated with marine
animals have been reported to cause disease in
humans (28, 111, 170).
The epidemiologies of human brucellosis differ
between areas of endemicity and
nonendemicity in terms of age, sex, season, and
risk factors. In regions of endemicity, such
as the eastern Mediterranean basin, Middle East, the
Arabian peninsula, Mexico, Central and
South America, the Balkan Peninsula, and the
Indian subcontinent, the disease occurs
among the general population. In the general
population, levels of infection are almost equal
among adults and children of both sexes and mostly
due to ingestion of unpasteurized goat,
sheep, cow, and camel milk or its products (e.g.,
soft cheese) (59, 100, 137, 163, 202).
In areas where the disease is not endemic,
infection is seen predominantly among adult
males, acquired occupationally by transmission
through direct skin contact (e.g., through
cuts and abrasions) with infected animal parts,
inhalation of aerosolized infected particles,
and accidental inoculation (e.g., sprays or
aerosols inoculated into the eye, mouth, and
nose). These infections occur mostly among dairy
industry professionals, veterinarians,
abattoir workers, and clinical and research
microbiology staff (16, 200).
Very rare cases transmitted through blood and bone
marrow transfusion, suspected sexual
intercourse, and banked human sperm have been
reported (121, 138, 153, 180, 185). Also,
a few cases of neonatal brucellosis have been
reported, and the isolation of Brucella from
human milk may explain it (99,
134).
Laboratory-acquired infection is an important
source of transmission. Brucella has a very low
infectious dose (≤102 organisms), and personnel
should adhere to strict safety precautions,
especially when handling cultures suspected of
containing the organims in clinical, research,
and production laboratories (124,
200). Most cases of laboratory-acquired disease
result
from mishandling and misidentification of the
organism (63, 124). The frequent failure of
clinical laboratories to correctly identify
isolates as Brucella species is particularly worrisome
from the perspectives of laboratory safety and
potential use as a bioweapon. B. melitensis,
B. abortus, and B.suis are category B select
agents (70, 136).
Clinical Categories of Human Brucellosis
The clinical categories of human brucellosis are
based on arbitrary criteria. In 1956, Spink
based them on the duration of manifestations
(acute, ≤2 months; subacute, 2 to 12 months;
chronic, ≥12 months) (172).
Subsequently, others based them primarily on clinical
manifestations (e.g., subclinical, localized,
chronic, and active, with or without localized
disease, including bacteremic and serological
classifications) (102, 202). To date, no uniform
definition has been adopted.
The incubation period is variable but usually
ranges between 1 and 4 weeks. The disease
onset is usually insidious, but its presentation
encompasses a wide spectrum of nonspecific
clinical manifestations, such as fever, sweats,
arthralgias, myalgia, fatigue, loss of appetite,
weight loss, hepatomegaly, and splenomegaly.
Complications can involve many organs and
tissues with signs of focal disease. The routine
hematology and biochemical profiles are
usually within normal limits, with some elevation
in erythrocyte sedimentation rate and liver
function tests. Thus, to the unaware physician,
the diagnosis of brucellosis can be a dilemma
and could protract for weeks and, in some
complicated cases, for years
(59, 100, 106, 156, 202). Increased business and leisure travel to
countries where the
disease is endemic has led to diagnostic
challenges in areas where brucellosis is uncommon,
especially when the presentation is unusual (42,
100, 112, 113, 202). Overall, the mortality
is very low, but morbidity is high. Previously,
brucellosis in childhood was thought to be
uncommon, but now it seems to be as prevalent as
and presents in a manner similar to that
in adults in areas of endemicity (98,
100, 163).
Because of these nonspecific clinical features,
human brucellosis was labeled the disease of
“mistakes.” It can be misdiagnosed and confused
with other diseases, such as typhoid fever,
rheumatic fever, tuberculosis, malaria, infectious
mononucleosis, endocarditis,
histoplasmosis, ankylosing spondylitis, pyelitis,
cholecystitis, thrombophlebitis, chronic
fatigue syndrome, collagen vascular diseases,
autoimmune diseases, and tumors
(100, 106, 135, 202).
Complications
The most commonly encountered focal complications
are osteoarticular (10 to 70%) (mostly
joints), genital in both males (6 to 8%) and
females (2 to 5%), neurological (3 to 5%),
cardiac (1 to 3%), pulmonary (1 to 2%), and renal
(< 1%). Mortality is very low (<1%) and
is almost exclusively due to cardiac complications
(39, 42,100, 101, 102, 106, 135, 163, 202).
Osteoarticular complications occur mostly as
arthritis (10 to 70%) and rarely as
osteomyelitis (<1%). The joints most frequently
involved are, in descending order,
sacroiliac, knee, hip, vertebra, ankle, and
multiple other joints. Generally, Brucella arthritis
can be misdiagnosed as rheumatoid arthritis,
rheumatic fever, tuberculosis, and systemic
lupus erythematosis.
Neurobrucellosis (3 to 5% of cases) can affect
both adults and children with diverse
presentations, including fever, headache,
meningeal signs, coma, or paresis. Depression and
mental fatigue are not uncommon complaints (51,
97, 161). CSF analysis, of both adults and
children, is nonspecific and can overlap with
other central nervous system diseases, such as
mycobacterial, viral, syphilitic or fungal
infections, or with noninfectious diseases, such as
psychiatric problems, multiple sclerosis, and
cancer (51, 97, 161). The yield
ofBrucella culture from CSF is low (5 to
30%). Therefore, the use of Brucella serology tests,
especially ELISA, on CSF specimens is essential to
diagnose neurobrucellosis (9, 161). With
appropriate treatment, the prognosis is usually
good for acute presentations and varies in
the setting of chronic disease.
Genital complications in males (6 to 8% of cases)
are mostly orchitis or epididymoorchitis
(40, 76, 100). In females, abortion (2 to 5%) has been
reported mostly in the first trimester
(103). Other rare complications reported for females
include cervicitis, salpingitis, tuboovarian
abscess, and ovarian dermoid cyst (103,
183).
Relapse is considered one of the most important
features of brucellosis and its complications
(19, 100, 102,171). Factors associated with relapse include the use
of less effective
antibiotic therapy, a positive blood culture
during the initial presentation, and ≤10 days’
duration of symptoms before initiation of
treatment.
Collection, Handling, Storage, and Transport of
Specimens
Specimens for the laboratory investigation of
cases with brucellosis may be sent for culture,
serology, and/or molecular testing. Culture can be
performed on a wide range of specimen
types, including blood (at least two sets), bone
marrow, CSF, pleural and synovial fluids,
urine, abscess specimens, and tissue specimens.
Adequate volumes should be secured prior
to initiation of antimicrobial therapy. Blood
(serum) and, when relevant, CSF specimens are
used for serologic testing. Molecular testing,
though usually for research purposes, can be
performed on blood (serum or whole blood, CSF, and
bone marrow) specimens.
The guidelines for proper specimen collection,
handling, transport, and processing are
generally similar to those reported for blood
cultures and other specimens submitted for
bacterial culture (refer to chapter 16 in this Manual). If delay in delivery to the lab is
anticipated, specimens can be held in the
refrigerator. To avoid/minimize laboratory
exposures to the pathogen, specimens from patients
suspected of having the disease should
be labeled appropriately and referred to a
reference laboratory, with the label specifying that
the laboratory should rule out brucellosis (44,
196).
Direct Detection
To circumvent the limitations of routine culture
and serodiagnostic tests for human
brucellosis, in-house-developed conventional PCR
and real-time (RT) PCR assays can directly
detect Brucella from clinical specimens.
Several Brucella-specific gene targets have been
used, including BCS P31 (encodes a 31-kDa cell
surface protein) and BP26 (encodes a 26-
kDa periplasmic protein), 16S rRNA, and the
insertion sequence IS711. The sensitivities of
these assays are quite varied, ranging from 50% to
100%. This variation might be related to
different DNA extraction methods, detection
formats, and different types of specimens
(43, 85, 110, 115, 119,122, 123, 147, 148, 189). The ribosomal 16S-23S ITS region
constitutes a suitable target in clinical
specimens and formalin-fixed paraffin-embedded
archived tissue, as well as for identification of
isolates from culture to the species level (85).
Molecular assays constitute a useful adjunct and
have promising potential for the diagnosis
of human brucellosis in a clinical laboratory
setting. Their routine incorporation in the
diagnosis of human brucellosis remains in need of
further optimization, standardization, and
improvement (85, 149).
Culture
Culture is considered the gold standard in the
laboratory diagnosis of brucellosis.
Conventional methods require long incubation times
(6 weeks) and are generally of variable
yields, being higher among patients with acute
brucellosis (40% to 90%) than in patients in
the chronic, focal, and complicated stages (5 to
20%) (10,41, 100, 199). When positive,
culture provides the definitive diagnosis. Bone
marrow cultures result in a 15- to 20%-higher
yield than p eripheral blood cultures. The
conventional standard m edium for the
nonautomated blood culture broth has been the
biphasic Ruiz-Castaneda bottle. The growth
of the pathogen takes between 7 and 35 days to
become positive, and the bottles should be
held for 6 weeks, with frequent visual inspection
(every 3 days) and terminal subculture
before the specimen is discarded as negative (10,
199).
Automated continuously monitored blood culture
systems such as Bactec (BD Diagnostics,
Sparks, MD) and BacTAlert (bioMerieux, Durham, NC)
show higher yields than the
conventional culture method and expedite the
detection of bacterial growth (majority
recovered within 1 week). There is no need to
incubate bottles longer than 10 to 14 days
(9a; 199). The lysis centrifugation system showed
improved and faster yields than
conventional methods in those labs that do not
have automated blood culture systems (199).
However, due to the need for centrifugation and
manipulation before direct plating, the
system may entail exposure and contamination
hazards. Rarely, some patients with
brucellosis have a positive blood culture in the
absence of positive serology (199, 202).
Recovery of Brucella from other clinical
material, such as bone marrow, CSF, joint fluid,
homogenate of tissues, and bones, in addition to
blood specimens, can be achieved by
inoculation of specimens into broth media (such as
those used for blood cultures) in addition
to plated media (blood and CA). The latter medium
is incubated at 37°C, preferably under 5
to 10% CO2, for up to 10 days prior to reporting
as negative.
Identification
Clinical microbiology laboratories should report
identifications of colonies suspected of
being Brucella spp. on the basis of a few
morphologic, biochemical, and serologic tests. All
manipulations of Brucella cultures should
be done in a biological safety cabinet. In these
setups, the colonies are generally recovered
directly from inoculated clinical specimens or as
a result of subculture from broth media (e.g.,
biphasic Ruiz-Casta neda medium and blood
culture medium showing signs of growth) on blood (Fig. 1C) and CA. Colonies can grow on
other media as well (e.g., Mueller-Hinton agar [Fig. 1D] and MacConkey agar [can show
variable growth]). Thayer-Martin or Martin-Lewis
medium can be used to
isolate Brucella spp. from contaminated
specimens. Generally, colonies are visible after 24 to
48 hours of aerobic incubation or incubation with
5 to 10% CO2 at 37°C, and there is no
need to keep the plates more than 72 to 96 hours
before discarding them as negative. The
colonies are 1 to 2 mm in diameter, entire,
smooth, and glistening. Rough variants can occur
with B. caniscolonies. The presumptive
identification of Brucella spp. from these colonies
entails demonstrating small gram-negative
coccobacilli (0.5 to 0.7 μm in diameter and 0.6 to
1.5 μm in length) (Fig. 2B). Biochemical reactions
show positive oxidase, catalase, and
urease tests, as well as a positive slide
agglutination reaction with specific B.
abortus and/or B. melitensis antisera (4,
41). Once these tests are performed and
completed, the clinical laboratory may report the
organism as presumptively Brucella spp.
Further characterization and speciation of the
pathogen involves extensive testing not
routinely performed in most clinical laboratories
(4,95). In the United States, LRN reference
laboratories are able to confirm and identify Brucella
to the species level, and these
laboratories can provide guidance and additional
pertinent information. See
alsohttp://www.bt.cdc.gov/lrn/.
When definitive identification is indicated,
conventional and molecular characterizations can
be used. Conventional classification/identification
to the species level of Brucella spp. can be
determined from results of certain reactions, such
as dye inhibition (thionin, fuchsin,
safranin), CO2 requirement, Tbilisi phage lysis,
oxidative metabolic tests (glutamic acid,
ornithine, ribose and lysine), and reaction to
monospecific sera.Brucella is usually subtyped
into biovars using multilocus enzyme
electrophoresis, pulsed-field gel electrophoresis,
randomly amplified polymorphic DNA analysis,
enterobacterial repetitive intergenic
consensus sequence PCR, repetitive intergenic
palindromic sequence PCR, amplified
fragment length polymorphism analysis, monolocus
(such as omp2a and omp2b) sequence
analysis, or multilocus sequence typing (3,
86, 113,193).
Typing Systems
One of the most highly discriminatory
methodologies for epidemiological subtyping of
isolates belonging to monomorphic bacterial
species is MLVA (96). In Brucella, MLVA
schemes with 21 loci (MLVA-21) and MLVA-16, which
use a combination of repeat markers
distributed across the Brucella genome,
were able to distinguish isolates of Brucella spp. of
widespread temporal and geographical origins or of
very close origins (3, 86, 193).
A Brucella MLVA database is hosted at http://mlva.u-psud.fr and contains data derived from
more than 500 animal and human Brucella isolates.
Molecular subtyping methods, especially
the promising MLVA, may potentially be useful not
only for epidemiological trace back
purposes or outbreaks but also for distinguishing
relapses from reinfection, thereby
influencing clinical therapeutic decisions (86).
Serologic Tests
Serologic assays are the most commonly relied upon
tests in the laboratory diagnosis of
brucellosis. Serological results are optimally
interpreted in the context of the evolution of
antibody responses after infection with Brucella
spp. IgM first appears, followed by the
appearance of IgG within 10 to 14 days. The
general evolution of these and other
immunoglobulins depends on response to treatment:
in recovery, a gradual and slow decline
in titers is observed, while persistent titers
alert the clinician to a poor response to treatment
due to focal complications, chronic infections, or
relapse (18, 62, 140). Persistence, (i.e.,
detection of antibodies, mostly IgG and some IgM, for
a very prolonged time [months and
sometime years]) is observed in 15 to 20% of
asymptomatic patients who have undergone
treatment and cure. The explanation for this
remains elusive (100).
Several antigens are used for serologic diagnostic
assays, generally obtained from B.
melitensis and B. abortus. Whole-cell preparations are the antigens
used in the agglutination
and the indirect fluorescent-antibody (IFA) tests,
while sonic extracts, purified LPS or protein
extracts of Brucella, are used mainly in ELISAs
(4, 7, 10, 12, 13, 87). Detection of antibodies
against infections due to B. canis and B.
ovis require using major outer membrane protein
antigens because these strains exist in a rough
colony form and do not share cross-reacting
antigens with the other Brucella spp. (113).
Since there is no standardized reference
antigen, it is important to note that the source
of the antigen, commercial or otherwise, can
influence the test results (14).
A wide range of in-house serologic tests and
formats have been used for investigating
patients with brucellosis (Table 4). These include direct agglutination tests in tubes, e.g., the
serum agglutination test (SAT), and on slides,
e.g., the rose bengal test, indirect Coombs
test, Brucellacapt tests, IFA test, and ELISA for
detection of immunoglobulin classes and
subclasses (6, 11,
13, 14, 15, 68, 154). Generally, agglutination-based tests cannot
differentiate the types of antibodies involved,
while the enzyme immunoassay (EIA) and IFA
test can. Commercial EIAs detecting Brucella IgG
and IgM with a high degree of sensitivity
and specificity have been available for a number
of years (17) and are considered an
excellent method for screening sera forBrucella antibodies
(13, 16).
Anti-Brucella Therapy
Appropriate antimicrobial therapy for treatment of
human brucellosis reduces morbidity,
prevents complications, and minimizes relapses.
Several anti-Brucella agents have been
used (e.g., doxycycline, rifampin,
trimethoprim-sulfamethoxazole, streptomycin-gentamicin,
some quinolones, and cephalosporins) with various
rates of success. Currently, the most
effective treatment regimen and optimal duration
of treatment remain unclear
(20, 59, 84, 98, 168).
Fewer relapses with combined regimens than with
monotherapy have been reported. For
adults with uncomplicated infection, the WHO
recommends oral doxycycline and rifampin for
6 to 8 weeks. Triple regimens using doxycycline,
rifampin, and an aminoglycoside for 2 to 3
months are recommended for patients with
endocarditis and neurobrucellosis.
Treatment regimens with fluoroquinolones and
broad-spectrum cephalosporins have been
used. Although these agents have good MICs in
vitro against Brucella spp., patients treated
with these regimens have higher rates of relapses
than patients on the standard regimen.
The use of fluoroquinolones in combination with
rifampin for the treatment of bacteremia and
complicated brucellosis has yielded varied results
(2, 168).
A recent systematic review and meta-analysis study
covering 30 trials and 77 treatment
arms showed that among patients with bacteraemia
and complicated brucellosis, higher
failure and relapse rates and shorter treatment
durations (less than 6 weeks) were observed
with monotherapy than with multidrug therapy. The
preferred treatment should be with dual
or triple regimens, including an aminoglycoside (168).
The question about postexposure prophylaxis
(doxycycline and rifampin therapy for 3 to 6
weeks) after a high-risk exposure in the lab
remains debatable. Guidelines for postexposure
management are empirical (32).
Upon possible exposure, however, recommendations were
made to take a baseline blood sample, monitor for
symptoms weekly for 6 months, and
perform serological surveillance at 0, 2, 4, 6,
and 24 weeks (33).
Prevention
Vaccines have been successful in the control of
livestock infections, which can subsequently
reduce infections in humans. Most veterinary
vaccines focus on live, attenuated B.
abortus (strain S19) and a more stable rough mutant of B. abortus (strain
RB51) for
cows, B. melitensis (strain Rev-1) for
sheep and goats, and B. suis 2 for swine. However,
developed vaccines have had limited efficacy in
humans and have been associated with
serious medical reactions. Heating of dairy
products and related foods has also been effective
in preventing disease transmission. The most
cost-effective approach to control and prevent
brucellosis relies on raising public awareness
about the disease and greater cooperation
between human and animal health sectors (204).
Evaluation, Interpretation, and Reporting of
Results
Interpretation of serologic test results in
relation to exposure, diagnosis, and prognosis of
the disease necessitates an accurate assessment of
the clinical history and current status of
patients and understanding the usefulness and
pitfalls of the laboratory tests (7, 202).
Positive cutoff titers in the Brucella agglutination
test for diagnosis have generally been
considered to be ≥160 in symptomatic patients.
However, much lower titers with the SAT
have been reported for patients with active
disease (100). Moreover, one has to be careful
when negative serology is encountered when
brucellosis is suspected, since this could be
due, for example, to infection with B. canis, which
can be missed by serologic assays
using B. abortus or B. melitensis antigen.
In addition, this could be due to very early disease
presentation, and thus repeat testing after 1 to 2
weeks is warranted (18, 85).
In acute brucellosis, elevation in Brucella-specific
IgG, IgM, IgA, IgE, IgG1, and IgG3 is
shown, while in those patients with chronic
brucellosis, elevations in IgG, IgA, IgE, IgG1, and
IgG4 are usually seen (6,
10, 15, 62). Monitoring the treatment response requires a
sequential follow-up for patients with serologic
titers. A decline indicates good prognosis,
persistently high titers necessitate continuous
monitoring, and a resurgence in antibody
titers most likely indicates relapse or
reinfection. Slide and TA titers fall faster than with the
EIA (18,21,
105, 140). Relapse has also been diagnosed by a detection
of a resurgence
in Brucella-specific IgG and IgA
antibodies, not IgM (18, 62, 140). Markers for differentiating
active from inactive disease are being sought. For
example, anti-Brucella cytoplasmic or
periplasmic protein antibodies, as determined by
ELISA, increased only in patients with
active brucellosis and were a better predictor of
cure than antilipopolysaccharide antibodies
(21,66, 151). Also, some interleukins show a decrease
posttherapy (104).
Though serologic tests are currently of high
importance for the investigation of patients with
brucellosis, several limitations can be
encountered, mainly lack of standardized antigen
preparations and assay methodologies, as well as
the detection of sustained high antibody
titers in some patients, despite treatment and
cure (7, 18, 62, 68, 140). False-positive
serologic results are rare. However, two cases
were recently reported in the United States, a
finding which led to initiating not only
unnecessary treatment but also a wide range of public
health investigations (34).
Based on the above, laboratories should use a
combination of two agglutination tests,
namely, the SAT and indirect Coombs test, the SAT
and Brucellacapt, or ELISAs for IgG and
IgM. In doing so, one would be able to detect
antibodies in different stages of the disease,
since in the acute stage any test can be positive,
while in chronic, complicated, or focal
disease cases, the SAT can be negative while the
Coombs test, Brucellacapt, and ELISA using
IgG can be positive (Table 5). Again, one should keep in mind that any serologic test findings
need to be interpreted in the context of the patient’s clinical
history.
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