Erratum

In the March 2003 issue, the reference for Table 1 on page 193 is incor-

rect. The correct reference is ‘‘Data from Pageat P. Properties of cat’s facial
pheromones. European Patent EP 0 724 832 B1. February 3, 1995.’’

Vet Clin Small Anim

33 (2003) x

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doi:10.1016/S0195-5616(03)00062-7

Preface

Emerging and re-emerging infectious

diseases

Guest Editors

My life-long interest in infectious diseases ﬁrst began under the mentor-

ship of my co-editor, Dr. Ed Breitschwerdt, while I was completing an
internship in small animal medicine and surgery at Louisiana State
University from 1980–81. There, in the bayou, infectious diseases such as
ehrlichiosis, diroﬁlarasis, and blastomycosis were rampant. When I moved
to ‘‘the big city’’ in Philadelphia, I was amazed to ﬁnd infectious diseases
there as well—babesiosis, leptospirosis, and borelliosis, to name a few. In
my present position at Auburn University, I have continued to pursue my
interest in this fascinating area of veterinary medicine. In 1997, we identiﬁed
a new species, Hepatozoon americanum, and in 2001, we helped to character-
ize the emergence of Babesia gibsoni infections in dogs in Alabama and
Georgia. Truly, the study of infectious diseases is one of the most exciting
and dynamic areas of veterinary medicine today.

As global travel increases, previously isolated regional diseases are be-

coming more widespread. In addition, the availability of sophisticated mo-
lecular techniques has led to a number of recent discoveries in the area of
infectious diseases. Many organisms are being reclassiﬁed based on molecu-
lar structure, resulting in new nomenclature in many cases. New diagnostics
are available and new infectious agents are being discovered.

This issue of the Veterinary Clinics of North America: Small Animal Prac-

tice brings together a dedicated group of academicians with an interest in

Vet Clin Small Anim

33 (2003) xi–xiii

Edward B. Breitschwerdt, DVM

Douglass K. Macintire, DVM, MS

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doi:

10.1016/S0195-5616(03)00036-6

infectious diseases. The articles in this issue provide an updated review of
some of the most current knowledge in this exciting and rapidly changing
area of veterinary medicine. I am grateful to the authors for their efforts
in helping to unravel some of the mysteries of the exciting ﬁeld of veterinary
medicine. I am also grateful to my co-editor, Dr. Ed Breitschwerdt, who
continues to set a standard of scientiﬁc excellence for veterinary research
in the area of infectious disease. I am grateful to the American College of
Veterinary Internal Medicine for providing a forum for veterinarians who
share a common interest in infectious disease by allowing us to gather as
a special interest group. It was at this forum that contacts with many of the
authors of this issue occurred. I am also grateful to my family, who continue
to put up with my inability to say ‘‘no’’ to most projects that come across
my desk. Finally, I give thanks to my Lord and Savior, Jesus Christ. Apart
from Him, I could do nothing. I hope that you enjoy this issue and that it
helps you to recognize, diagnose, and treat the emerging and re-emerging in-
fectious disease agents affecting animals in your practices today.

Douglass K. Macintire, DVM, MS

March 18, 2003: As I contemplate a few words of wisdom with which to

preface this issue of the Veterinary Clinics of North America: Small Animal
Practice, our country stands on the brink of war with Iraq. In comparison
with the events of the next several weeks, efforts to characterize and to bet-
ter understand the complexities of emerging and re-emerging infectious dis-
eases seem to be of little consequence. However, infectious diseases continue
to be a major factor that impacts the health, safety, and security of people
throughout the world. Infectious agents substantially compromise the well
being of both animals and people in developing nations as well as more
developed countries. Historically, infectious diseases, some of which are vec-
tor-transmitted, have thrived during periods of war and starvation—periods
during which hygiene and nutrition are deplorable and immunological sur-
veillance systems are taxed to their limits. Unfortunately, this is an aspect of
history that mankind elects to repeat time and time again.

The role of veterinarians as contributors to the global management of in-

fectious diseases continues to expand. This expansion in the role of veteri-
narians as an integral part of the public health infrastructure is certainly
appropriate, because veterinarians are comprehensively trained to function
as comparative infectious diseases diagnosticians. Training in immunology,
microbiology, parasitology, virology, epidemiology, and medicine, across
a spectrum of animal species, provides veterinarians with the unique oppor-
tunity to make important contributions in the context of emerging and re-
emerging infectious diseases. Of comparative medical importance, most
organisms currently listed as potential agents for bioterrorism were ﬁrst
identiﬁed as animal pathogens.

As I reﬂect on my career as a veterinary internist, I am most grateful to

my students. I have been very fortunate to have excellent students. Through-

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D.K. Macintire, E.B. Breitschwerdt / Vet Clin Small Anim 33 (2003) xi–xiii

out my tenure as an academic veterinarian, I have learned many daily
lessons from those whom I was charged to teach. Several of these individuals
are authors of the articles in this issue. It is has been a pleasure to work with
Dr. Dougie Macintire, a friend, a colleague, and an infectious diseases re-
searcher who has made important contributions to our current understand-
ing of several emerging infectious diseases. Finally, I would like to
acknowledge the patience and love of my wife, Anne, and the support of
my two sons, Brett and Kyle. May our collective eﬀorts make the world
a better and safer place in which people and animals can live in peace.

Edward B. Breitschwerdt, DVM

Douglass K. Macintire, DVM, MS

Department of Clinical Sciences

College of Veterinary Medicine

Auburn University, AL 36849, USA

Email Address: macindk@vetmed.auburn.edu

Edward B. Breitschwerdt, DVM

Department of Clinical Sciences

College of Veterinary Medicine

North Carolina State University

4700 Hillsborough Street

Raleigh, NC 27606, USA

Email Address: ed_breitschwerdt@ncsu.edu

xiii

D.K. Macintire, E.B. Breitschwerdt / Vet Clin Small Anim 33 (2003) xi–xiii

Update on molecular techniques for

diagnostic testing of infectious disease

Rance K. Sellon, DVM, PhD

Department of Veterinary Clinical Sciences, Washington State University,

Post Oﬃce Box 7060, Pullman, WA 99164, USA

The application of molecular biology tools to the diagnosis of infectious

disease is increasing in small animal veterinary medicine. The polymerase
chain reaction (PCR), reverse transcriptase PCR (RT-PCR), analysis of
restriction fragment length polymorphisms (RFLP), and others that often
were developed initially for research purposes have increased knowledge of
some infectious disease agents. In the process, there was rapid recognition of
the diagnostic potential of these assays, and some have become available for
diagnostic testing of suspected infections in dogs and cats.

The tools of molecular biology rely on biochemical properties of nucleic

acids (DNA and RNA) imparted by the nucleotide composition (relative
proportions of each of the four nucleotides) and the nucleotide sequence.
The nucleotide composition of nucleic acids inﬂuences their denaturation
(the separation of the two complementary strands that compose DNA or
DNA/RNA hybrids) and hybridization properties (the ability of nucleic acid
sequences to bind to each other to form double-stranded nucleic acid
moieties). Nucleotide sequence also inﬂuences hybridization properties and
dictates susceptibility to nucleic acids being ‘‘cut’’ at speciﬁc locations using
restriction enzymes. This discussion is not intended to provide the details
necessary to perform, or even understand all aspects of the molecular
techniques presented, but rather is meant to convey an overview of some of
the available techniques, the diagnostic power of these applications, some of
the diagnostic limitations of these assays, and the variety of clinical
applications possible with the molecular techniques.

Vet Clin Small Anim

33 (2003) 677–693

E-mail address:

rsellon@vetmed.wsu.edu.

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doi:10.1016/S0195-5616(03)00023-8

Molecular assays for detecting infectious agents

The most widely used of the molecular tools for the diagnosis of

infectious disease are the PCR and the RT-PCR. The PCR is used for initial
detection of DNA and so is most useful for detection of infectious agents
that contain DNA as their primary genetic material. Because the PCR is
incapable of detecting RNA, in order to detect infectious agents that have
RNA as their primary genetic material (many viruses for example), a copy
of DNA (cDNA) can be made from the infectious agent’s RNA through the
process of reverse transcription (RT). Transcription normally produces
a messenger RNA from a DNA template; the enzyme reverse transcriptase
promotes the synthesis of a DNA molecule from an RNA template. Once
the cDNA has been synthesized, a PCR can be performed subsequently, and
the entire process is referred to as RT-PCR.

The PCR uses short single-stranded segments of nucleotides, called

primers, the sequences of which are complimentary to DNA sequences of
the intended target DNA, for example, the DNA of an infectious organism.
Primers serve as the initial template upon which a new DNA molecule can
be synthesized. The primers and other necessary reagents of the PCR are
added to a volume of solution containing representative DNA from the
sample of interest, including host DNA and DNA from the intended target
of detection. The test sample can be anything that could harbor the agent of
interest such as tissue, ﬂuids such as urine or blood, stool, or others. The
reaction mix is heated to separate DNA into its two strands, then cooled to
allow primers to bind to complimentary regions of denatured target DNA.
The reaction then is heated again in an extension step to promote the
addition of nucleotides to the primer ends, thus building a ‘‘new’’ strand of
DNA. The PCR cocktail is subjected to 25 to 40 cycles of heating and
cooling to preferentially amplify target DNA segments. Ampliﬁcation
occurs on a geometric scale; theoretically, one copy of the target DNA
sequence can be ampliﬁed to over 30 million copies in 25 cycles. The PCR
products, or amplicons, are detected most commonly by using gel
electrophoresis. The presence of target DNA in the test sample is suggested
by observation of a speciﬁc and predicted size DNA band in the gel (Fig. 1).

The most common clinical application of the PCR to infectious disease

diagnosis in dogs and cats is the detection of a single suspected infectious
agent. In these instances, some or all of the DNA sequence of the infectious
agent must be known to design agent-speciﬁc primers. The PCR can be
used, however, to cast a wider net in cases in which an infectious agent is
suspected, but for which speciﬁc sequence information is not available, or in
which the presence of an infectious disease is suspected, but one is unsure of
which speciﬁc agent is present. In these situations, instead of performing the
PCR with primers that are speciﬁc for a single agent, universal primers can
be designed to amplify a segment of a gene that retains a high degree of
sequence similarity, or homology, among a large group of related infectious

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organisms. Related is a relative term when used in this sense and could refer
to organisms related by genus, family, or more broadly still as for example
all prokaryotic bacteria. One example of this broad-based approach is
illustrated by detection of the gene encoding the 16S ribosomal RNA
(rRNA) of prokaryotic bacteria, a gene that maintains a high degree of
homology across many genera of bacteria. If a product is ampliﬁed using
these universal primers, the presence of bacterial DNA in the reaction is
established. The reaction amplicons are analyzed by sequencing or other
strategies, and the identity of the agent established by documenting
similarities to organisms already existing in large databases.

Fig. 1. Schematic representation of the polymerase chain reaction (PCR). DNA (a) in the test
sample is heated to separate the two strands (b). The reaction is cooled to allow primer (short
bold lines

) annealing to the target DNA (c), then heated again to allow addition of nucleotides

to extend the nascent DNA molecule (d). At the end of the cycle, the target DNA has been
duplicated (e). Each new DNA can then itself be a targeted molecule for the next cycle of
ampliﬁcation. The process is repeated for 25–40 cycles, and then products are visualized fol-
lowing gel electrophoresis ( f ). The left hand lane represents DNA markers of a known size
against which the size of the amplicons is measured.

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The universal primer PCR can be the ﬁrst step in design of a genus or

species-speciﬁc PCR when detailed genetic information regarding the target
agent is unknown. Amplicons obtained from universal primer PCR can be
sequenced, with the sequence analysis then suggesting a speciﬁc genus, or
perhaps a speciﬁc species, of organism. Sequences that are identiﬁed as
unique to the 16S rRNA gene or other genes of the detected organism then
become the foundation for design of the next set of speciﬁc primers. Thus, one
has gone from casting a broad net with universal primers to the development
of a focused PCR in just a few steps. Such a strategy was employed in the
development of a PCR assay speciﬁc for Haemobartonella felis [1].

Although primer selection is critical for establishing PCR speciﬁcity,

there are steps that should be taken to conﬁrm the speciﬁcity of the
amplicons, primarily to be sure that there are no other unknown DNA
segments that share the same sequence of the primer binding sites of the
target organism. Conﬁrmation of the speciﬁcity is particularly crucial when
assays ﬁrst are developed or are applied to new sample types that represent
a diﬀerent pool of DNA than that in which the assay was developed. Among
the methods that can conﬁrm amplicon speciﬁcity, DNA hybridization,
amplicon sequencing, and analysis of restriction enzyme digestion patterns
are among the most commonly used. DNA hybridization uses a short DNA
segment that is labeled to permit detection and which recognizes a sequence
within the amplicon ﬂanked by the primer binding sites. Detection of the
probe will occur only when the probe has become bound to its comple-
mentary sequence in the amplicon and is retained through several washes
that remove unbound probe. Obtaining a predicted sequence in the amplicon
following sequence analysis also conﬁrms the speciﬁcity of the ampliﬁcation.

With primers designed to be very speciﬁc for the target sequence, the PCR

is very speciﬁc. The PCR is typically also very sensitive because of its ability
to detect very small numbers of target DNA. Modiﬁcations of the PCR have
been developed to further increase the sensitivity of the PCR, typically by
adding a small volume containing amplicons from a ﬁrst reaction to a second
PCR. The primers of the second PCR are complementary to sequences of the
amplicons from the ﬁrst reaction, a technique known as nested PCR. The
sensitivity of any of the PCR strategies facilitates the detection of some
organisms below the threshold of detection of routine microbial cultures,
cytology, histology, or perhaps immunohistochemical detection.

In another modiﬁcation of the PCR, the in situ PCR, the reaction is

conducted on tissue samples (biopsies for example), and the detection of the
amplicons achieved by means other than gel electrophoresis to allow
demonstration of the target sequence in a particular tissue location. In situ
PCR thus allows correlation of agent detection with the presence of
a histological lesion, or localizes the agent to a particular cell type, a ﬁnding
that would provide a compelling argument that the organism detected
played a role in disease causation. In situ PCR can be combined with other
techniques such as immunohistochemistry to more completely characterize

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cells or tissues in which the amplicons are detected, information that can be
useful in understanding disease pathogenesis.

Polymerase chain reaction and RT-PCR are limited to diagnostic or

research laboratories with the equipment and personnel trained to perform
the assays. It is possible that in the future, assays relying on this technology
may be available in a bedside format accessible to the practitioner or staﬀ.
Indeed, such bedside assays are already appearing in the human medical
ﬁeld and are considered standard approaches to the diagnosis of some
infections of people [2]. In the meantime, the practitioner still has access to
some of these laboratory-based assays as samples, such as tissue and ﬂuids
(blood, eﬀusions, and others), usually are easily acquired and submitted.
Any special collection or handling requirements as set forth by the
laboratory should be understood before sample acquisition and submission.
It is important to recognize that not all laboratories will be in a position
to oﬀer all available PCR-based assays for infectious disease diagnosis,
and samples submitted to one laboratory for a given test may be sent to
a diﬀerent laboratory better equipped to perform the assay. Cost of the
assays will vary with the laboratory and can reﬂect how frequently the test is
performed and technical aspects of the assay.

Restriction fragment length polymorphisms

DNA is susceptible to being ‘‘cut’’ at restriction sites by restriction

enzymes. The restriction site recognized by a given restriction enzyme is
deﬁned by DNA nucleotide sequences; thus, each restriction enzyme
recognizes and cuts at a very speciﬁc nucleotide sequence. There are many
restriction enzymes used for cutting DNA, and for a given restriction
enzyme, multiple restriction sites may exist within a particular DNA
segment from a given individual or organism. When DNA that is obtained
from an organism is subjected to the action of restriction enzymes and the
cut DNA subjected to gel electrophoresis, a pattern of restriction fragments
of varying lengths that is unique to the organism is produced (Fig. 2).
Variations in the lengths of fragments, known as polymorphisms, arise from
diﬀerences in DNA sequence between the restriction sites. Diﬀerences in the
restriction fragment lengths alter the migrating properties of the fragments
during gel electrophoresis, allowing comparison between organisms. The
more restriction enzymes, within reason, a given DNA sample is cut with,
the more patterns there can be for analysis. Analysis of restriction fragment
length polymorphisms (RFLP) often is referred to as ‘‘DNA ﬁngerprinting,’’
as RFLP are typically unique to individuals or groups of closely related
organisms. Comparison of polymorphisms between an unknown and
a known pattern can facilitate detection of a new or variant organism
related to the known agent. RFLP analysis often is combined with PCR or
RT-PCR; the PCR-based assays produce an amplicon, the identity of which
can be more precisely established by RFLP analysis.

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Compared with the PCR-based assays, analysis of RFLPs as a strictly

diagnostic technique has had more limited application to the diagnosis of
infectious disease in small animal medicine. RFLP is quite useful in
demonstrating relatedness or divergence of infectious organisms in an
individual or populations of animals. Comparison of RFLP patterns can
demonstrate emergence of variant pathogens within a population, or even
within the same individual in the case of persistent infections. A recent
report describes by RFLP analysis the emergence of variants of Bartonella
henselae

within cats with chronic infection, a ﬁnding that could suggest

a mechanism for persistent infection [3].

In situ hybridization

Another technique that has seen somewhat limited applications in small

animals is the use of labeled genetic probes to identify the nucleic acid of an
infectious agent within a particular cell or tissue type, a technique known as
in situ hybridization. In situ hybridization, like the PCR-based assays, takes
advantage of the fact that complementary DNA sequences will bind to each
other with high aﬃnities. Thus, a probe with a complementary sequence to
a known gene can be labeled to permit easy detection, and the probe added,
under appropriate conditions, to a sample with cells or tissues to see if the
target nucleic acid is present. In situ hybridization thus generally requires
a priori knowledge of the organism that is being sought, as the probes tend
to be organism-speciﬁc. Because in situ hybridization, like in situ PCR, can
associate the nucleic acid of an infectious agent with a particular cell or
tissue type, in situ hybridization can be a powerful tool for elucidating cell
tropism of infectious agents, often a key to the pathophysiology of infectious
diseases. For example, in situ hybridization has been used to localize feline
immunodeﬁciency virus (FIV) to particular cells in the thymus [4], canine and
feline herpesvirus to a number of tissues [5,6], and H felis on erythrocytes [7].

Fig. 2. Schematic representation of restriction fragment length polymorphism (RFLP), or
DNA ﬁngerprinting, analysis. DNA (A) is cut with a restriction enzyme to produce a number of
DNA fragments of varying lengths (B). The DNA fragments are separated by gel electro-
phoresis to produce a pattern unique to the organism and the enzyme used to cut the DNA. In
this representation, six diﬀerent samples were analyzed, with one of the six (in the 4th lane)
clearly exhibiting a pattern diﬀerent from the other ﬁve.

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Representational diﬀerential analysis

Representational diﬀerential analysis (RDA) makes use of the PCR and

hybridization properties of nucleic acids to ‘‘subtract’’ normal DNA from
the total pool of DNA in a test sample [8]. When normal DNA is subtracted
from the total DNA present, what remains are exogenous nucleic acid
sequences such as those of infectious organisms. The exogenous sequences
then can be ampliﬁed by the PCR, the amplicons analyzed by sequencing or
other strategies, and the results compared with information in databases to
identify relationships to known infectious organisms. The author is aware of
no published studies in which RDA has been used to establish the diagnosis
of an infectious disease in small animal patients. This approach has been
used in people to suggest the existence of a novel herpesvirus infection
considered to be a likely cause of Kaposi’s sarcoma in people with HIV-1
infection, however [9].

Analysis of DNA libraries

An interesting approach to establishing the identity of unknown

infectious organisms takes advantage of a host immune response and the
ability to put large segments of DNA into bacterial plasmids or other
organisms like yeast, which can be cultured. The cultured organisms often
are referred to as expression vectors, since the proteins encoded by the
inserted DNA sequence can be produced, or expressed, in detectable
quantities in culture. In so doing, a library of DNA, or cDNA if starting
from RNA, is generated and can be analyzed by a number of methods. With
one technique, nucleic acids are collected from an individual with the
suspected infectious disease, and the nucleic acid cloned, or inserted, into an
expression vector. Each of the clones generated is cultured, with the clones
producing proteins encoded by the inserted genes. The patient’s serum then
is used to screen each of the cultured clones for the production of proteins
recognized by the patient’s antibodies. Clones recognized by patient
antibodies then are analyzed to establish a putative identity of the clone
based on its similarity to other pathogens or organisms. This technique was
used to establish the identity of a new hepatitis virus in people associated
with transfusion-associated hepatitis [10]. To accomplish this feat, the
investigators had to screen approximately one million clones to ﬁnd one that
produced a candidate infectious disease antigen.

Limitations of molecular assays

Like all diagnostic test results, interpretation of results generated from

molecular assays needs to be considered in light of all available information
about the patient, including history, physical examination ﬁndings, and
results of other diagnostic tests. Although very useful in the assessment of
dogs and cats for infectious diseases, the molecular assays are not without

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limitations. For example, the extreme sensitivity of the PCR is also its
biggest limitation, as false positive results can occur with even minute con-
tamination of test samples or reactions. Contamination can occur during
sample collection or during the course of performing the PCR. It is impor-
tant to know that the laboratory has taken relevant steps to eliminate
contamination and to ensure that a positive result is truly positive and not a
reﬂection of assay contamination. The extreme sensitivity of the PCR is
the biggest obstacle to routine use of universal primer PCR as a diagnostic
tool, as contamination with bacteria at any step has the potential to result
in a false-positive reaction. Thus, universal primer PCR is better for de-
tection of organisms in normally sterile sites, such as blood, nonepithelial
tissues, or other locations from which samples can be obtained aseptically.

As sensitive as they are, the PCR or RT-PCR are nonetheless susceptible

to false-negative results. The PCR-based assays may be falsely negative if
there are PCR inhibitors, which may include proteins such as hemoglobin or
others, if the DNA or RNA in a sample is of poor quality because of
degradation, or if there are technical problems with the assay. Performing
the assay on an inappropriate sample also could cause false-negative results.
For example, the sensitivity of detection of canine distemper virus in one
experimental study improved if whole blood, serum and cerebrospinal ﬂuid
were tested by RT-PCR, as any one of these samples from any given dog
could be negative [11]. Another theoretical cause of false-negative results
would be changes in an organism’s nucleic acid sequences that preclude
primer binding during the annealing step of the PCR. If primers do not
anneal to target DNA sequences, a new DNA strand cannot be synthesized.

Most laboratories performing diagnostic PCR will include known positive

and negative samples as controls for technical problems with the assay, or as
a guard against false-positive results from contamination of the assay. To
help further reduce false-positive results from reaction contamination, some
laboratories will include a control sample that has all the components of the
reaction cocktail except a nucleic acid source, replacing the test sample with
water. If a product is detected in one of these reagent controls, products in
any other sample must be considered as potential contaminants.

Detection of a nucleic acid of an infectious organism by one of the

molecular methods does not necessarily mean that the agent is the cause of
clinical disease. A molecular assay may be positive in instances where the
organism is in fact not a cause of the clinical disease, as might occur with
agents that cause latent infections. Likewise, a negative result would not
guarantee that an infectious agent is not responsible for the clinical disease
observed. A microbial toxin produced at a site distant from the tissue
sampled for the assay may be responsible for the clinical disease and thus
would not be detected in the molecular assays. As previously suggested,
a variant of the organism not recognized by a particular primer pair would
lead to a false-negative result in the face of infection. If the sample submitted
was not appropriate for detection of the agent (eg, submission of tissue

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when blood was needed, or collection of a sample at the wrong stage of
infection), the result of an assay could be negative and not reﬂect the true
infection status of the patient. Detection of Borrelia burgdorferi was better
accomplished by PCR of skin samples than blood samples because of the
low level of spirochetemia associated with that infection [12].

Another limitation of the molecular assays is that, in general, they are not

well-suited to assess responses to therapy. It is likely that nucleic acids
persist in the host after the death of an organism, and although the duration
of DNA persistence often is presumed to be less than a few weeks, the actual
duration of DNA persistence following organism death is not known. One
group of investigators of acute Rocky Mountain spotted fever was surprised
to ﬁnd that PCR results for Rickettsia rickettsii were positive for at least 8
days beyond the last positive culture [13], a combination of results that
would be consistent with the persistence of DNA for a time after all
detectably viable organisms had been eliminated from the host. Modiﬁca-
tions of the PCR can provide quantitative information regarding the copy
numbers (numbers of identical DNA segments in the test sample) of nucleic
acid present, which in most instances would be interpreted to reﬂect the
number of organisms actually present in the sample. Thus, a decline in copy
number following a therapeutic intervention would be consistent with
a therapeutic response. Such an approach could prove more helpful in
assessing responses to therapy as compared with a nonquantitative assay.
The RT-PCR has been suggested as more useful than PCR assays to
monitor responses to therapy [14]. RNA is typically more labile than DNA.
Free RNA, or RNA that is not involved in protein synthesis as might occur
with the death of an infectious agent, typically is quickly destroyed by host
RNAses, enzymes that destroy RNA. Thus, the argument holds that if RNA
from an infectious organism is detected in an RT-PCR assay, the presence of
viable organisms is implied. RT-PCR-based testing of viral load is standard
for people infected with HIV-1 [15].

Clinical applications of molecular diagnostic tools

Molecular diagnostic techniques will not supplant the use of traditional

methods of infectious disease diagnosis such as microbial culture, serologic
assays, and microscopic detection of organisms. Molecular biological
assays, however, do oﬀer particular advantages over traditional methods
of infection diagnosis in certain clinical settings. The molecular assays can
also be complementary to the traditional approaches. Advantages to the
molecular approaches can include more rapid conﬁrmation of the presence
or absence of a particular pathogen; the assays, especially the PCR and
RT-PCR, have the potential to be completed within 24 to 48 hours of
the laboratory’s receipt of the sample, including time needed for sample
processing. This compares favorably to the days or weeks sometimes
required for cultures to be declared positive or negative. The time advantage

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is dependent on the turnaround times of the laboratory, however, which
could reﬂect demand for the assay and strategies that minimize expense of
the assay. Another advantage of the molecular assays is that, since recovery
of viable organisms is typically not a goal of the molecular assays, there is
no need to use special media for transport of many samples, which makes
sample acquisition easier for most in private practice. Depending on the
assay and the laboratory, some PCR assays can be cost-competitive with
traditional approaches.

One of the calling cards of the techniques described previously is their

ability to provide information that could be helpful in the diagnosis of
organisms that cannot be cultured (eg, many viruses, some mycoplasma such
as Haemobartonella felis) or organisms that are diﬃcult or slow to grow in
culture (eg, Mycobacteria). Because detection of these agents by molecular
methods does not require having viable organisms, detection of their nucleic
acid ‘‘footprints’’ can give clues to their existence in a host. As stated
previously, a PCR-based assay exists for documentation of H felis infection.
Likewise, PCR approaches have been used to document infection with my-
cobacterial agents in dogs and cats [16–18]. These assays also have been
applied to the diagnosis of enteric viral infections, as PCR-based assays have
been developed for pathogens such as canine and feline coronaviruses [19,20].

Another advantage of the molecular assays is the fact that they can detect

evidence of infection before an infected patient mounts a detectable
antibody response. Thus, the molecular assays are attractive for establishing
the diagnosis of acute infections before seroconversion, and they could
obviate the need for collection of acute and convalescent samples often
required for demonstration of seroconversion. Assays for Rocky Mountain
spotted fever and leptospirosis have been developed [13,21,22], and these
infections would be good examples of diseases that historically have
required evidence of serologic conversion to provide evidence of infection.

Although serologic conversion can provide strong evidence of infection,

there are some diseases for which antibodies exist because of vaccine-induced
antibodies (eg, leptospirosis) or because natural exposure to organisms is
common (eg, Toxoplasma gondii). For infections such as these, serologic
assays often add supportive evidence of infection, but the conﬁdence provided
by positive serology can be slim in some clinical situations. Thus, detection of
the molecular footprints in a seropositive animal would add an extra degree of
conﬁdence in a diagnosis. For example, detection of leptospiral DNA in the
blood or urine of a dog with clinical signs would provide strong evidence of
infection and could provide evidence of serovar-speciﬁc infection supported
by antibody titers that are high against that particular serovar. These assays
also have the potential to clarify whether young animals that are antibody
positive to an infectious disease are infected, or are positive simply because of
maternally derived antibodies. Clariﬁcation of the infection status of kittens
that are positive for antibodies to FIV could be clariﬁed by PCR assays that
detect the FIV provirus in feline blood cells were one commercially available.

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Another application of the molecular assays would be documentation of

latent infections or animals that are carriers of infectious agents. PCR assays
have documented latent herpesvirus infection in dogs [23] and clinically
normal cats [24]. PCR has been used to document clinically silent ehrlichial
infections in dogs 34 months after experimental inoculation [25].

Molecular assays also can diﬀerentiate between pathogenic and non-

pathogenic forms of organisms or between vaccinal and ﬁeld isolates of
infections for which modiﬁed-live virus vaccines exist. RFLP analysis is used
commonly to document the existence of pathogenic versions of bacteria
such as Escherichia coli [26]. Manifestations of canine distemper virus or
feline panleukopenia virus infections that follow closely on the heels of
vaccination can be examined by use of the molecular techniques to
determine if the clinical disease is caused by the vaccine isolate or a ﬁeld
isolate [27,28].

Clinicians frequently encounter cases of animals that die or are

euthanized before a deﬁnitive diagnosis is established. In some of these
patients, evidence or suspicion of an infectious etiology is generated by
results of necropsy and histological examination of tissues, but blood or
other tissues may not be available, or are no longer suitable, for traditional
diagnostic techniques such as serology and culture. An infectious etiology
can be supported in some of these cases by the molecular assays, since some
of them can be performed on nucleic acid extracted from paraﬃn-embedded
tissue blocks. Thus, another advantage of the molecular assays is the ability
to retrospectively analyze samples for infectious disease [5,6].

Selection of appropriate antimicrobial therapy also can be directed by

PCR-based assays and RFLP analysis. Genes in infectious organisms can
encode antimicrobial resistance proteins, and these genes could be targets of
detection for PCR or RFLP. Thus, resistance to a particular class of anti-
microbials could be documented before the organism has been cultured and
antimicrobial resistance patterns identiﬁed by more routine methods. Mo-
lecular methods are the tests of choice for detection of methicillin-resistant
Staphylococcus aureus

and vancomycin-resistant enterococci in people [2].

Other clinical applications that are supported by molecular diagnostic

tools include the characterization of zoonotic infections. A novel chlamydial
infection in a person was traced, using PCR and RFLP, to an organism
harbored by the patient’s cat [29]. Cat scratch disease caused by Bartonella
clarridgeiae

infection in a person occurred with an isolate identical to one

recovered from the patient’s cat [30]. These assays also have tremendous
potential to deﬁne new causes of infectious disease, be it detection of
variants of well-known organisms, or perhaps even novel infectious causes
of well-characterized clinical syndromes that have deﬁed etiologic de-
scription. Recently, an ehrlichial-like syndrome was observed in dogs that
were negative for Ehrlichia canis by an E canis-speciﬁc PCR routinely used
by the investigators. Using primers speciﬁc for the Ehrlichia genus 16S
ribosomal RNA gene, the investigators were able to amplify ehrlichial DNA

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from the clinical patients [31]. Sequence and DNA hybridization analysis of
this amplicon suggested that the dog was infected with E ruminantium, an
ehrlichial agent not previously associated with clinical disease in dogs. The
investigators were careful to state that detection of the DNA of this
ehrlichial organism was not deﬁnitive proof that the infection was the
cause of the dog’s clinical disease, but their ﬁnding certainly raises that
possibility.

Establishing a relationship between organisms and disease

The power of the molecular assays to uncover the existence of new

organisms or new variants in ill animals raises questions about cause and
eﬀect, since in many of these cases, recovery of viable organisms may not be
accomplished easily. The classic methodology of establishing the relation-
ship between a putative infectious agent and clinical disease was to fulﬁll
criteria established by Koch. Brieﬂy, Koch’s postulates held that a given
agent was the cause of clinical disease if:

The agent was found in every case of the disease.
The agent was not found in other diseases.
The agent could be isolated and cultured, and caused disease in a new

host.

A fourth postulate, that the agent could be isolated from the experimen-
tally inoculated host, is considered an additional point of proof of infectious
disease causation.

Although rigorous satisfaction of the postulates provides convincing

evidence of a cause and clinical eﬀect, the postulates break down with regards
to infectious organisms that cannot be cultured, or are very diﬃcult to
culture. To account for the applications of the tools of molecular biology to
the diagnosis of infectious disease, and especially with regards to discovery of
novel infectious organisms, newer criteria to support disease causation have
been proposed [32]. Researchers have suggested that these criteria include:

The sequence of the putative agent should be detectable in most cases of

the disease.

Hosts without disease caused by the putative agent should have no, or

few, copy numbers.

Resolution of the clinical disease should be associated with either

a decrease in copy number or an inability to detect the agent.

Detection of the agent before onset of clinical disease, or an increasing

copy number that is associated with the onset or severity of clinical
disease, makes cause and effect more likely.

The properties of the putative agent inferred from its sequence or

genetic relationship to other organisms should be consistent with other
agents of that particular type.

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If a phenotype, such as a predictable pattern of clinical signs, laboratory

abnormalities, or histopathological lesions, is predictable based on the
presence of the sequence or increasing copy numbers, a causal link is
strengthened.

The strength of a causal link is enhanced if the sequence is detected

readily in lesions, but not in normal tissues.

The ability to detect the sequence should be reproducible.

Questions arise from a consideration of these criteria when the clinician is

presented with a patient or a literature description of a patient from which
the nucleic acid of an infectious agent has been detected. First and foremost,
the clinician needs to ask if it is likely that the infectious organism is truly
responsible for the clinical signs or other abnormalities detected in the
patient. In many cases, the association will be suggested because of well-
established links between the infectious agent and clinical signs (eg, acute
fever, petechial hemorrhage and thrombocytopenia in a dog with R
rickettsii

). In other cases, the link may be more diﬃcult to discern, as for

example a cat with signs of hemolytic anemia and detection of Haemobarto-
nella

by a PCR. Although it is known that Haemobartonella can cause

hemolytic anemia, it is also known that there are other causes of hemolytic
anemia in cats, and that many cats are asymptomatic carriers of
Haemobartonella

. Thus, a PCR-positive blood sample would not guarantee

that hemolytic anemia was caused by Haemobartonella felis infection.
Likewise, T gondii parasitemia has been documented in experimentally
infected cats that show no clinical signs of infection [33]. Thus, a cat
demonstrating signs of neurological or respiratory disease that was positive
for T gondii on the basis of a PCR on a blood sample may or may not have
clinical toxoplasmosis. The strength of disease association in such an
instance would be bolstered by a positive result from a tissue lesion. Thus, as
is the case with interpretation of other diagnostic tests for infectious
organisms, an understanding of the biology of the infection is also important
for proper interpretation of results of molecular assays.

With increasing clinical application of the PCR-based assays or other

molecular approaches, there likely will be increasing reports of novel
infectious organisms being associated with clinical diseases in dogs and cats.
When reading such reports, it would do the reader well to recall some of the
guidelines suggested above before concluding that the organism detected is
the cause of a new disease. For example, there have been several recent
reports of dogs with Bartonella infections detected by PCR with a diverse
array of clinical diseases including granulomatous diseases and peliosis hep-
atis [34,35]. These conditions have been associated with Bartonella infection
through detection of Bartonella DNA in lesions. Granulomatous disease and
peliosis hepatis in dogs have not yet been shown conclusively to be caused by
Bartonella

infections, however, as few of the criteria for establishing a cause

and eﬀect by molecular diagnostic methods have been fulﬁlled.

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R.K. Sellon / Vet Clin Small Anim 33 (2003) 677–693

Summary

The era of diagnostic molecular biology has arrived for small animal

clinicians, and it is a near certainty that assays such as the PCR and RT-
PCR will become more widely available for a wider array of infectious
agents. Already there is an extensive list of infectious diseases of dogs and
cats that have been investigated with molecular tools. A partial list is
included in box 1.

An understanding of the advantages and disadvantages of the molecular

techniques and some of the questions these techniques can answer for
clinicians can serve practitioners well in their approach to the diagnosis of
infectious diseases in dogs and cats. It is likely that additional applications of
these tools to small animal medicine will become apparent as investigators
use and reﬁne them for their research purposes, or as new uses emerge from
human medical applications. Clinicians also are likely to reap the beneﬁts of
this knowledge. Because samples often are acquired easily from clinical
patients in most practice settings, access to these tools puts all clinicians in

Box 1. Partial list of infectious agents for which molecular
assays have been developed and used in dogs and cats

Viral [36–42]

Bornavirus in dogs and cats
Canine adenovirus
Parvovirus (canine and feline)
Feline calicivirus
Rabies

Bacterial [36,43]

Chlamydia psittaci
Helicobacter

Protozoal [44–47]

Cytauxzoon felis
Leishmania in dogs and cats
Babesia species
Hepatozoon americanum

Fungal [48,49]

Histoplasma capsulatum
Cryptococcus neoformans

These are infectious agents for which the molecular assays described in this

article have been used to study the organism in cats and dogs. To author’s
knowledge, none of these assays are available for routine diagnostic tests.

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the group of discoverers of new, or variations of, infectious diseases and their
clinical manifestations.

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Pythiosis, lagenidiosis, and zygomycosis

in small animals

Amy M. Grooters, DVM

Department of Veterinary Clinical Sciences, School of Veterinary Medicine,

Louisiana State University, Baton Rouge, LA 70803

Pythiosis, lagenidiosis, and zygomycosis are devastating and often fatal

infectious diseases that aﬀect animals living in temperate, tropical, and sub-
tropical climates throughout the world. The oomycete Pythium insidiosum
and the zygomycetes Conidiobolus species and Basidiobolus species have been
recognized as animal and human pathogens for many years. Members of the
genus Lagenidium (a group of oomycetes closely related to the genus
Pythium

), however, have been identiﬁed as a cause of oomycosis in dogs only

recently [1]. Although they are caused by a phylogenetically diverse group of
pathogens, pythiosis, lagenidiosis, and zygomycosis share similar clinical and
histologic characteristics (all cause lesions characterized by pyogranuloma-
tous and eosinophilic inﬂammation associated with broad, irregularly
branching, sparsely septate hyphae), making them diﬃcult to distinguish
from one another. Because of these similarities, many veterinary pathologists
and clinicians continue to use the inclusive but out-dated term ‘‘phycomy-
cosis’’ when describing histologic or clinical ﬁndings that could be caused by
any of these pathogens. Although this term is a convenient label for cases in
which a deﬁnitive culture-based diagnosis has not been made, it is no longer
an appropriate taxonomic designation, and it should be replaced with the
more speciﬁc terms ‘‘pythiosis,’’ ‘‘lagenidiosis,’’ and ‘‘zygomycosis.’’ Clini-
cally, distinguishing between these infections is important because of dif-
ferences in epidemiology, choice and duration of therapy, and prognosis.

Pythiosis

Pythiosis, caused by the aquatic oomycete Pythium insidiosum, is known

best as a cause of gastrointestinal (GI) or cutaneous disease in dogs and of
cutaneous and subcutaneous disease in horses [2–4]. It also has been

Vet Clin Small Anim

33 (2003) 695–720

E-mail address:

agrooters@vetmed.lsu.edu.

0195-5616/03/$ - see front matter

doi:10.1016/S0195-5616(03)00034-2

described as an uncommon cause of cutaneous or subcutaneous lesions in
cats [5,6] and calves [7] and of arteritis, keratitis, or periorbital cellulitis in
people [8–11]. In nondomestic species, P insidiosum has been recognized as
a cause of primary pulmonary disease in a captive Central American jaguar
cub in southern Louisiana [12], of cutaneous and GI lesions in a group of
captive spectacled bears in a South Carolina zoo, (Grooters and Lamberski,
unpublished data), and most recently, as a cause of a subcutaneous
mandibular lesion in a captive dromedary camel in Florida (Grooters and
Wellehan, unpublished data).

History

The clinical manifestations of P insidiosum infection ﬁrst were recognized

in the middle of the 19th century by British veterinarians in India who ob-
served a chronic cutaneous granulomatous disease of horses that they
termed ‘‘bursattee.’’ In the late 19th century, a fungal etiology for the
disease was proposed by several investigators based on histologic ﬁndings
[13,14]. Although the pathogen was isolated as early as 1901 by Dutch
investigators working with horses in Indonesia [15], it could not be induced
to sporulate, and thus was assumed to be a zygomycete or ‘‘phycomycete’’
fungus based on the morphologic characteristics of its vegetative hyphae. It
was not until 1974 that Austwick and Copland were able to produce bi-
ﬂagellate zoospores from isolates obtained from horses in New Guinea,
identifying the pathogen as an oomycete that was likely a member of the
genus Pythium [16]. The binomial P insidiosum was introduced by de Cock
in 1987, who examined several isolates obtained from horses and dogs in
several geographic regions (including the United States) and found them to
be identical [17]. In addition, de Cock was able to produce sexual repro-
ductive structures from these isolates in vitro and was therefore able to
identify the pathogen as a new species of Pythium.

Taxonomy

Members of the genus Pythium are water- or soil-dwelling organisms that

belong to the Kingdom Stramenopila, Class Oomycetes, Order Pythiales,
and Family Pythiaceae. Although many Pythium species are economically
important plant pathogens, P insidiosum is the only mammalian pathogen
recognized in this genus. Recent taxonomic studies based on ribosomal
RNA gene sequence data have conﬁrmed that members of the class Oomy-
cetes

are phylogenetically distant from the Kingdom Fungi, and actually are

related more closely to algae than to fungi [18]. The taxonomic distance
between oomycetes and fungi are reﬂected on the cellular level by diﬀerences
in cell wall and cell membrane composition. Chitin, an essential component
of the fungal cell wall, is generally lacking in the oomycete cell wall, which
instead contains predominately cellulose and b–glucan [19]. In addition,
oomycetes diﬀer from fungi in that ergosterol is not a principal sterol in the

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A.M. Grooters / Vet Clin Small Anim 33 (2003) 695–720

oomycete cell membrane. Not all oomycetes share the same sterol bio-
chemistry, however. Members of the genera Pythium, Lagenidium, and
Phytophthora

are sterol auxotrophs, meaning that they incorporate sterols

from their environment rather than producing them [19–21]. In general,
sterols are required for in vitro production of reproductive structures, but
not necessarily for growth of vegetative hyphae.

Classiﬁcation of oomycetes generally is based on the morphologic cha-

racteristics of their sexual reproductive structures (oogonia and antheridia)
and, to a lesser degree, asexual reproductive structures (zoospores). Because
P insidiosum

was for many years the only zoosporic organism known to be

a mammalian pathogen, some authors mistakenly have suggested that the
production of zoospores speciﬁcally identiﬁes an isolate as P insidiosum
[6,22]. In actuality, there are numerous zoosporic eukaryotes in the class
Oomycetes and among the Chytridiomycetes [18,23]. Many speciﬁc
characteristics of asexual reproduction are shared by members of the genera
Pythium

, Lagenidium and Phytophthora, including the production of motile,

reniform, biﬂagellate zoospores that develop by progressive cleavage within
a vesicle that forms at the end of a discharge tube produced by a zoo-
sporangium (Fig. 1) [24–26]. Diﬀerentiation between these genera and their
identiﬁcation to species level usually is based on morphologic characteristics
of sexual reproductive structures [27]. P insidiosum is characterized by in-
tercalary, smooth, and subglobose oogonia; diclinous antheridia that pro-
duce a rigid fertilization tube from their tip; and oospores that are aplerotic
and often pressed to one side of the oogonium [17]. Unfortunately, because
isolates of P insidiosum rarely produce sexual reproductive structures in the

Fig. 1. Photomicrograph illustrating asexual reproduction of Pythium insidiosum at the edge of
an infected blade of grass. Note that protoplasm (P) can be visualized moving down the
discharge tube into the enlarging vesicle. Abbreviations: DT, discharge tube; VES, vesicle; ZSP,
zoospore. (Courtesy of Amy Grooters, DVM, Louisiana State University)

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laboratory, their deﬁnitive identiﬁcation based on morphologic features is
usually not possible. As a result, in clinical practice and in the veterinary
literature [28], isolates that have colonial and hyphal characteristics con-
sistent with P insidiosum, grow well at 37

C, and produce motile biﬂagellate

zoospores in water culture have been assumed to be P insidiosum by default,
given that no other species of Pythium has been recognized as a mammalian
pathogen. More recently, molecular tools have been developed that allow
deﬁnitive identiﬁcation of many Pythium species (including P insidiosum)
without the need for identifying speciﬁc sexual characters in vitro [29,30].

Biology and life cycle

A description of the life cycle of P insidiosum was ﬁrst proposed in 1983

by Miller [31] and was expanded in 1993 by Mendoza, who described the
details of in vitro asexual reproduction as observed by light and scan-
ning electron microscopy [32]. Although data regarding the maintenance
of P insidiosum in aquatic environments and the speciﬁc factors that lead
to mammalian infection are lacking, a wealth of related information has
been generated by studies of plant pathogenic species of Pythium and
Phytophthora

The infective form of P insidiosum is thought to be the motile biﬂagellate

zoospore, which is released into aquatic environments and likely causes
infection by encysting in damaged skin or GI mucosa. Zoospore production
begins with the formation of zoosporangia, which develop from hyphae that
have colonized certain types of plant material. In the laboratory, production
of zoosporangia can be induced by incubating infected grass blades at
37

C in a dilute salt solution [33,34]. Microscopically, zoosporangia of

P insidiosum

are ﬁlamentous and undiﬀerentiated from vegetative hyphae.

Early in the process of zoosporogenesis, discharge tubes (thin-walled
tubular structures that are long and narrow but widen at the tip) develop
from zoosporangia (Fig. 1). As the process progresses, sporangial proto-
plasm ﬂows down the discharge tube and forms a terminal vesicle that
continues to expand as it is ﬁlled. Progressive cleavage of the protoplasm
within the vesicle then results in the formation of zoospores, which mecha-
nically break through the thin vesicle wall and are released (Figs. 2A,B)
[17,32].

Pythium insidiosum

zoospores (like those of many other oomycetes) have

an anterior tinsel ﬂagellum (which generates the majority of thrust), and
a posterior whiplash ﬂagellum (which acts as a rudder), both of which
originate from a ventral or lateral groove (Fig. 2C). Once released from the
vesicle, zoospores swim in a spiral or helical pattern that is interrupted
intermittently by random changes of direction. This pattern of motility
(which is mediated largely by calcium) is part of a complex homing sequence
that allows the zoospore to locate, move toward, and encyst on speciﬁc host
tissues or other substrates [35]. Processes that have been found to be

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important in the zoospore homing behavior of plant pathogenic Pythium
species include chemotaxis (mediated by certain amino acids or sugars
released from damaged roots and root tips) [36], electrotaxis [37], and auto-
aggregation, in which dense accumulations of zoospores may attract other
zoospores, increasing the inoculum for infection [38]. Once the target tissue
has been located, zoospores encyst (Fig. 2D) and adhere with a precise
orientation such that the predetermined point of germ tube eruption is
facing the host tissue, allowing rapid tissue penetration [35]. Adhesion is
facilitated at the time of encystment by the secretion of a ‘‘sticky’’ glyco-
protein that coats the cyst surface [39]. The use of similar mechanisms by
P insidiosum

zoospores to seek out and invade mammalian tissues is

supported by observations made by Mendoza during baiting experiments
[32]. When horse skin and hair baits were placed in a suspension of
P insidiosum

zoospores, the zoospores immediately moved toward the baits

and eventually encysted on them. Microscopic examination of the skin baits
revealed that most zoospores had adhered to the cut edges of skin and rarely

Fig. 2. Scanning electron micrographs illustrating asexual reproduction of Pythium insidiosum.
(A) Terminal vesicle containing well-developed zoospores; ﬂagella (arrow) can be visualized
through defects that have developed in the vesicle wall before its rupture. (B) Multiple
zoospores (arrow) being released from a vesicle that has just ruptured. (C) Zoospore resting with
its dorsal surface on a nucleopore ﬁlter; note the ventral groove from which both ﬂagella
originate. (D) Encysted zoospore that has produced a germ tube (arrow;) note that the shape of
the zoospore has changed from reniform to spherical, and the ﬂagella have been lost.
Abbreviation:

DT, discharge tube. (Courtesy of Amy Grooters, DVM and William Henk, PhD,

Louisiana State University)

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had encysted on the undamaged skin surface. On the hair baits, encysted
zoospores accumulated at the follicular end of the hair rather than along the
hair shaft (an observation that had also been made by Miller using human
hair baits) [31].

The speciﬁc factors that favor sporulation of P insidiosum in the

environment are unknown, but there is some evidence that it may have an
aﬃnity for certain types of plant material, prompting one author to hy-
pothesize that aquatic plants may play a role in the life cycle as a natural
host [31]. In the early 1980s, Miller used human hair baits to attempt to
isolate Pythium species from water samples collected from swampy habitats
in northern Queensland, Australia [31]. Pythium species were isolated in
pure culture from all samples obtained from 2 of 10 locations that were
studied, suggesting that zoospores may be released in high numbers in
certain areas, when climactic or other environmental factors favor
sporulation. Unfortunately, since the sexual stage of P insidiosum had not
been described at the time that work was done, it is not possible to know
whether the Pythium species isolated by Miller was actually P insidiosum.
Recent attempts by the author’s laboratory to isolate P insidiosum from
a freshwater lake in southeastern Louisiana using canine skin baits were
unsuccessful, yielding only non-insidiosum Pythium species.

Epidemiology and etiology

In the United States, pythiosis is encountered most often in the Gulf

Coast states, but it has been recognized in animals living as far north as New
Jersey, Virginia, North Carolina, southern Illinois, southern Indiana, and
Kentucky, and as far west as Oklahoma, Missouri, and Kansas. The author
also recently conﬁrmed three cases of GI pythiosis in dogs living in Arizona
that had not traveled outside the southwestern United States (Grooters
and Dial, unpublished data). Globally, pythiosis most often is encountered
in southeast Asia (especially Thailand and Indonesia), eastern coastal
Australia, New Zealand, and South America (especially Brazil and Costa
Rica), but it also has been documented in Korea, Japan, and the Caribbean
(Haiti).

Although the clinical syndrome associated with pythiosis has been recog-

nized for more than a century, very little is known about the epidemiologic
factors that contribute to its development. It is identiﬁed most often in
young (1 to 3 years old), male, large breed dogs, and it is especially common
in outdoor working breeds such as Labrador retrievers. Aﬀected dogs are
presented to the veterinarian more often in the fall, winter, and early spring
months [3]. In cats, speciﬁc breed and sex predilections have not been
observed in the few cases that have been reported. Of 10 cats with cutaneous
pythiosis diagnosed through the author’s laboratory since 1999, however,
ﬁve were younger than 10 months old, with an age range of 4 months to
9 years (Grooters and Evans, unpublished data).

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Infected dogs often have a history of recurrent exposure to warm

freshwater habitats. A signiﬁcant percentage of animals with pythiosis have
not had exposure to these types of environments, however, and it is not
uncommon to diagnose pythiosis in suburban house dogs that have not
visited lakes, swamps, or ponds. Until a technique for identifying and
quantifying P insidiosum in aquatic and soil environments can be developed,
the role of environmental exposure in the development of this disease likely
will remain ambiguous.

Other risk factors for developing pythiosis have not been identiﬁed.

Aﬀected animals are immunocompetent and otherwise healthy. Given the
aﬃnity of P insidiosum zoospores for damaged skin [32], it would make
sense to assume that animals with cutaneous wounds or parasite-induced
injury to GI mucosa would be more likely to become infected. Solid epi-
demiologic evidence to support this assumption is lacking, however. The
presence of a traumatic wound before the development of cutaneous
pythiosis has been reported in a few canine and equine cases. Because
traumatic incidents are rarely observed by the owner, however, it is often
diﬃcult to determine whether lesions noted early in the course of disease
resulted from trauma or simply from early infection.

Clinicopathologic ﬁndings

The lesions associated with cutaneous pythiosis in small animals are

typically nonhealing wounds and invasive masses that contain ulcerated
nodules and draining tracts (Fig. 3A) [2,6]. Aﬀected dogs most often are
presented to the veterinarian for evaluation of cutaneous or subcutaneous
lesions involving the extremities, tailhead, ventrum, or perineum. Regional
lymphadenopathy is often present (especially in animals that have recurrent
disease following partial surgical resection) and usually reﬂects extension of
infection rather than just reactive inﬂammation.

Lesions in 10 cats with cutaneous pythiosis recently evaluated though the

author’s laboratory have included subcutaneous masses (some of which
were highly invasive) in the inguinal, tailhead, or periorbital regions, and
draining nodular lesions or ulcerated plaque-like lesions on the extremities,
sometimes centered on the digits or footpad. Two previously described cats
with pythiosis had large subcutaneous masses aﬀecting the extremities and
cranioventral thorax, but no cutaneous involvement [6]. In a third pre-
viously reported case, P insidiosum infection resulted in nasal cavity and
nasopharyngeal lesions and bilateral retrobulbar masses [28].

Gastrointestinal pythiosis in dogs typically is characterized by severe

segmental transmural thickening of the stomach, small intestine, colon,
rectum, or, rarely, the esophagus or pharyngeal region (Figs. 3B,C) [3,
40–42]. Mesenteric lymphadenopathy is common, and this occasionally is
observed without accompanying GI tract lesions. The gastric outﬂow
area (including proximal duodenum) and ileocolic junction are the most

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frequently aﬀected portions of the GI tract, and it is not uncommon to ﬁnd
two or more segmental lesions in the same patient. Inﬂammation in aﬀected
regions is typically centered on the submucosa, with variable mucosal
ulceration and occasional extension of disease through serosal surfaces, re-
sulting in adhesion formation and peritonitis. Involvement of the mesenteric
root may cause severe enlargement of mesenteric lymph nodes, which often
are embedded in a single large ﬁrm granulomatous mass that is palpable in
the mid-abdomen. Extension of disease into mesenteric vessels may result
in bowel ischemia, infarction, perforation, or acute hemoabdomen [43]. In
addition, infection may extend from the GI tract into contiguous tissues
such as pancreas and uterus [3,40]. In one recent report, a prostatic abscess
caused by P insidiosum was observed in a dog with an adjacent colonic
lesion [44]. The route of infection in this dog was not determined.

Clinical signs associated with GI pythiosis include weight loss, vomiting,

diarrhea, or hematochezia. Physical examination often reveals a very thin
body condition and a palpable abdominal mass. Signs of systemic illness
such as lethargy or depression are not typically present unless intestinal
obstruction, infarction, or perforation occurs. Cutaneous and GI lesions

Fig. 3. Photographs of the gross lesions associated with canine pythiosis. (A) Distal extremity
of a 2-year-old female spayed Labrador retriever with multiple draining tracts. (B) Cross section
of a segment of infected colon that had been resected from a 3-year-old female Doberman
pinscher; note the severe mural thickening and decreased luminal diameter. (C ) Lesser
curvature and pyloric antral regions of the stomach of a 6-month-old male Catahoula; note the
extreme thickening of the gastric wall.

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rarely are encountered together in the same patient. Laboratory abnormal-
ities that may be associated with pythiosis include eosinophilia, anemia,
hyperglobulinemia, and hypoalbuminemia. In dogs with GI pythiosis, ab-
dominal radiography and sonography usually reveal segmental thickening
of the GI tract, an abdominal mass, or mesenteric lymphadenopathy [45].

Although GI pythiosis in cats has not been reported previously, the

author’s laboratory recently conﬁrmed P insidiosum infection causing
segmental eosinophilic granulomatous enteritis in two cats (Rakich et al,
unpublished data). In each case, the diagnosis was made by histology,
immunohistochemistry, and immunoblot serology. Complete surgical re-
section of the lesion was curative in one cat.

Systemic dissemination of pythiosis has been described only once [2]. A 2-

year-old male German shepherd was presented with a severe necrotic and
ulcerative lesion surrounding the left stiﬂe. In addition, the dog exhibited
depression, vomiting, diarrhea, and neurologic abnormalities. The patient
died despite amputation and supportive care, and necropsy revealed an
ileocolic mass, intestinal perforation that had resulted in septic peritonitis,
multifocal 2 to 10 mm lesions in the kidneys and heart, and a 1 to 2 cm focal
lesion in the cerebral cortex. P insidiosum hyphae were detected in the
cutaneous, ileocolic, renal, cardiac, and cerebral lesions.

Lagenidiosis

Although the class Oomycetes includes many taxa that are common plant

pathogens, P insidiosum long has been considered to be the only mammalian
pathogen in this class. In March of 1999, however, the author isolated
another oomycete from tissue biopsies taken from a dog with severe invasive
subcutaneous disease. The dog died acutely following rupture of a caudal
vena caval aneurysm, and necropsy revealed severe sublumbar lymphade-
nitis and pyogranulomatous vasculitis [46]. Sequencing of the 18s ribosomal
RNA gene of the isolate recovered from this dog identiﬁed it as an oomycete
in the genus Lagenidium. The author has since identiﬁed more than 40 dogs
with serologic, histologic, or culture evidence of Lagenidium infection.
Complete clinical, necropsy, and ribosomal RNA sequence data are
available on six of these cases [1].

Taxonomy and biology

The genus Lagenidium includes more than 50 species, most of which

occur as parasites of algae, fungi, rotifers, nematodes, crustaceans, Daphne,
and mosquito larvae. The best studied species, Lagenidium giganteum, is
a facultative pathogen of mosquito larvae that is approved by the Environ-
mental Protection Agency for use in the United States as a biological control
agent for mosquito populations [47,48]. The author’s data suggest that the
canine pathogenic Lagenidium species is related closely to L giganteum,

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based on strong antigenic similarities (demonstrated by the degree to which
sera from Lagenidium-infected dogs recognize antigens of L giganteum) and
molecular similarities [1]. Diﬀerences in their in vitro growth characteristics
(Grooters, unpublished data) and the failure of L giganteum to infect
rodents in mammalian safety studies [49], however, suggest that the canine
pathogenic Lagenidium species and L giganteum are likely distinct species.
Although little is known about the life cycle and biology of the canine
pathogenic Lagenidium species, it is likely similar to that of P insidiosum and
L giganteum

Epidemiology and clinicopathologic ﬁndings

The epidemiologic and clinicopathologic features of lagenidiosis that have

been identiﬁed are similar in many respects to those that previously have been
associated with cutaneous pythiosis. Aﬀected animals are typically young to
middle-aged dogs living in the southeastern United States. Although most of
these dogs have been from Florida or Louisiana, the author has identiﬁed
cases in Texas, Tennessee, Virginia, and Indiana also. A number of infected
dogs have had frequent exposure to lakes or ponds. Two of these dogs were
a pair of unrelated housemates [1], suggesting that environmental exposure to
the pathogen may play an important role in infection.

The majority of dogs with lagenidiosis initially are presented for eval-

uation of progressive cutaneous or subcutaneous lesions (often multifocal)
involving the extremities, mammary and inguinal regions, perineum, or
trunk [1]. Grossly, these lesions appear as ﬁrm dermal or subcutaneous
nodules, or as ulcerated, thickened, edematous areas with regions of necrosis
and numerous draining tracts (Fig. 4). Regional lymphadenopathy often is
noted, and in one dog in which cutaneous lesions were absent, submandi-
bular lymphadenopathy was the presenting complaint [1]. Animals with
great vessel or sublumbar lymph node involvement often develop hind limb
edema. Similar to the clinical course typically associated with cutaneous
pythiosis, the skin lesions in dogs with lagenidiosis tend to be progressive,
locally invasive, and poorly responsive to therapy. In contrast to pythiosis,
however, most dogs with lagenidiosis have been found to have lesions in
distant sites, including great vessels, sublumbar and inguinal lymph nodes,
lung, pulmonary hilus, and cranial mediastinum. GI lesions caused by
Lagenidium

species have not been observed, and infection in mammals other

than dogs has not been identiﬁed.

Zygomycosis

The term ‘‘zygomycosis’’ refers to infections caused by fungi in the class

Zygomycetes,

including the genera Basidiobolus and Conidiobolus in the

order Entomophthorales, and the genera Rhizopus, Absidia, Mucor,
Saksenaea

, and others in the order Mucorales. In people and animals, the

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Mucorales

tend to cause acute, rapidly progressive disease in debilitated or

immunocompromised individuals, whereas the Entomophthorales typically
cause chronic localized infections in subcutaneous tissue or nasal submucosa
of immunocompetent patients [50,51]. Culture-conﬁrmed cases of mucor-
mycosis (infections caused by pathogens in the order Mucorales) have not
been described well in small animal patients. Basidiobolus species and
Conidiobolus

species, however, have been reported to cause cutaneous

pyogranulomatous lesions in dogs that are grossly and histologically similar
to those caused by P insidiosum [52]. Unfortunately, clinical and pathologic
information about zygomycosis in small animal patients is lacking because
of the diﬃculty in establishing a deﬁnitive diagnosis, which historically has
required tissue biopsy and culture. As a result, dogs with zygomycosis often
go undiﬀerentiated from those with pythiosis.

Epidemiology and clinicopathologic ﬁndings

Basidiobolus ranarum

(previously B haptosporus), Conidiobolus coronatus,

C incongruus

, and C lamprauges are saprophytes that are distributed widely

in natural environments throughout the world. Conidiobolus and Basidio-
bolus

species are found in soil and decaying plant matter, and Basidiobolus

species also are isolated commonly from insects and from the feces of
amphibians and reptiles [51]. Cutaneous infection with Basidiobolus or

Fig. 4. Photographs of the gross lesions associated with canine lagenidiosis. (A) Ulcerative
dermatitis caused by Lagenidium species infection in a 2-year-old female border collie presented
for progressive skin lesions and generalized lymphadenopathy. (B) Close-up view of left hind
limb showing a large eschar distal to the ulcerative lesion. (Courtesy of Amy Grooters, DVM,
Louisiana State University)

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Conidiobolus

species likely occurs by percutaneous inoculation of spores by

way of minor trauma or insect bites. Secondary capilliconidia of B ranarum
have been identiﬁed on the surface of mites and likely are carried by other
insects also [53]. Dogs with palate lesions caused by conidiobolomycosis
may become infected by direct implantation of spores when chewing on
decaying pieces of wood. Infection also may result from inhalation or
ingestion of spores. Aﬀected animals are typically immunocompetent.

In people, horses, sheep, and other mammalian species, conidiobolomy-

cosis occurs most often as a nasopharyngeal infection with or without local
dissemination into tissues of the face, retropharyngeal region, and retro-
bulbar space. In two dogs evaluated by the author, a presumptive diagnosis
of nasopharyngeal conidiobolomycosis was made on the basis of histologic
and serologic ﬁndings. One of these dogs was presented for signs of chronic
nasal cavity disease, and the other had a severe chronic ulcerative dermatitis
of the nasal planum. In two additional canine patients evaluated in the same
hospital, culture-conﬁrmed conidiobolomycosis was associated with ulcer-
ative lesions of the hard palate (Fig. 5) [54]. Both dogs had radiographically
apparent involvement of the nasal cavity or nasopharynx. In a ﬁfth dog
described in the literature, Conidiobolus infection was associated with multi-
focal nodular draining subcutaneous lesions and regional lymphadenopathy

Fig. 5. (A) photograph of an ulcerative lesion of the hard palate caused by Conidiobolus species
infection in a 1.5-year-old female German shepherd dog. (Courtesy of Stephen LeMarie´, DVM,
Southeast Veterinary Specialists, Metairie, LA and Carol Foil, DVM, Louisiana State
University) (B) Photomicrograph showing Conidiobolus species hyphae (arrow) in a tissue
biopsy from an ulcerative hard palate lesion in a 3-year-old female boxer; note the large amount
of amorphous inﬂammatory material (eosinophilic sleeve) surrounding the hyphal segment;
H&E stain.

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[55]. In addition, the author recently conﬁrmed Conidiobolus species causing
pneumonia in a dog that was receiving immunosuppressive therapy
(Grooters and Hawkins, unpublished data).

Basidiobolomycosis is a rare cause of ulcerative draining skin lesions in

dogs, and it also has been reported in a single case as a cause of respiratory
disease [56]. Disseminated Basidiobolus species infection involving the GI
tract and other abdominal organs has been described in two dogs [3,57]. In
addition, the author has evaluated two dogs (one with a focal preputial
lesion and another with a focal vulvar lesion) in which a presumptive diag-
nosis of basidiobolomycosis was made on the basis of histologic and sero-
logic ﬁndings (Grooters et al, unpublished data). Complete surgical
resection of the infected tissue was curative in both cases.

Culture-conﬁrmed cases of zygomycosis in cats have not been described

well. The author recently made a presumptive diagnosis of conidiobolomy-
cosis (based on histopathology and serology) in a 3-year-old cat with an
ulcerative lesion of the hard palate. Eleven cases of suspected mucormycosis
were reported in a necropsy study from Switzerland [58], but the diagnoses
were based on histologic ﬁndings alone, and there was no attempt to dif-
ferentiate the pathogens involved from oomycetes or members of the order
Entomophthorales

. Lesions in most of these cats involved the GI tract or lungs.

Diagnosis

Because obtaining a deﬁnitive diagnosis of oomycosis or zygomycosis has

been challenging traditionally, diﬀerentiation between these diseases has
remained diﬃcult. Subsequently, most P insidiosum, Lagenidium, and zygo-
mycete-infected veterinary patients continue to be labeled with a pre-
sumptive diagnosis such as ‘‘suspected pythiosis,’’ ‘‘suspected zygomycosis,’’
or even ‘‘phycomycosis’’ on the basis of histopathologic ﬁndings. Although
the presumptive diagnosis of pythiosis is likely correct in most canine, feline,
and equine cases (simply because it is more common), the inability to
distinguish pythiosis from lagenidiosis and zygomycosis consistently is
troublesome for several reasons. First, because of diﬀerences in cell mem-
brane composition between these taxonomically distinct organisms, zygo-
mycosis may be more likely than oomycosis to respond to medical therapy
with traditional antifungal agents that target ergosterol. Second, the prog-
nosis associated with these diseases may diﬀer; the available information
suggests that cutaneous pythiosis and lagenidiosis may be more aggressive
infections than cutaneous zygomycosis. Finally, diﬀerentiating between
these diseases is essential to more fully characterize their clinical, pathologic,
and epidemiologic features. This is especially true for lagenidiosis, a disease
for which epidemiologic data is needed urgently.

Recently, a number of highly speciﬁc serologic, immunohistochemical,

and molecular-based tools for the diagnosis of pythiosis and lagenidiosis
have been developed in the author’s laboratory. Initial evaluations indicate

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that these assays are likely to make the deﬁnitive diagnosis of pythiosis or
lagenidiosis possible for most aﬀected animals, even when culture is un-
successful. In addition, they may provide the veterinary practitioner with an
opportunity to make a diagnosis earlier in the course of disease, when
lesions may be more amenable to surgical or medical therapy.

Cytology

In animals with cutaneous pythiosis, lagenidiosis, or zygomycosis,

cytologic evaluation of exudate from draining tracts, impression smears
made from ulcerated skin lesions, and ﬁne needle aspirates of enlarged
lymph nodes often reveal pyogranulomatous, suppurative, or eosinophilic
inﬂammation. Hyphae are observed occasionally, and their morphologic
appearance in conjunction with a typical inﬂammatory response can provide
a tentative diagnosis of oomycosis or zygomycosis (Fig. 6A). Microscopic
examination of macerated tissue that has been digested in 10% potassium
hydroxide may be more likely to reveal hyphal elements than other cytologic
specimens [52]. Little information is available regarding the usefulness of
cytology for the diagnosis of oomycosis and zygomycosis in noncutaneous
forms of disease. In the one previously reported case of respiratory
Basidiobolus

infection and in an unpublished case of Conidiobolus pneu-

monia, however, cytologic examination of bronchoalveolar lavage ﬂuid re-
vealed characteristic fungal hyphae [56].

Fig. 6. (A) Fine needle aspirate specimen obtained from an enlarged inguinal lymph node in a
1-year-old female spayed Labrador retriever with a large tailhead lesion caused by pythiosis;
note the presence of multiple broad, poorly septate hyphal structures (arrows); modiﬁed
Wright’s stain. (Courtesy of Casey LeBlanc, DVM, Louisiana State University) (B)
Photomicrograph of a GMS-stained section of P insidiosum-infected tracheobronchial lymph
node from a 7-month old male jaguar cub with pulmonary pythiosis.

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Histology

Pythiosis

The histologic ﬁndings associated with pythiosis generally are character-

ized by eosinophilic granulomatous to pyogranulomatous inﬂammation with
ﬁbrosis. Aﬀected tissues typically contain multiple foci of necrosis sur-
rounded and inﬁltrated by neutrophils, eosinophils, and macrophages. In
addition, discrete granulomas composed of epithelioid macrophages, plasma
cells, multinucleate giant cells, and fewer neutrophils and eosinophils may be
observed [2,40,59].Vasculitis is occasionally present. Organisms usually are
found within areas of necrosis or at the center of discrete granulomas [3].
Although P insidiosum hyphae are diﬃcult to visualize on H&E-stained
sections, they may be identiﬁed as clear spaces surrounded by a narrow band
of eosinophilic material [3]. Hyphae are visualized easily in sections stained
with Gomori’s methenamine sliver (GMS) but not with periodic acid-Schiﬀ
(PAS). They are broad (mean, 4 lm; range, 2 to 7 lm), rarely septate, and
occasionally branching (Fig. 6B) [2,3].

Cutaneous pythiosis typically causes severe nodular to diﬀuse ulcerative

dermatitis and panniculitis. Because areas of inﬂammation are found most
often in the deep dermis and subcutis, deep wedge biopsies are preferred to
punch biopsies when pythiosis is suspected. In GI pythiosis, granulomatous
inﬂammation centers on the submucosal and muscular layers rather than the
mucosa and lamina propria [3]. For this reason, the diagnosis of pythiosis
may be missed on endoscopic biopsies that fail to reach deeper tissues. In
addition, pythiosis should be considered as a diﬀerential diagnosis when
endoscopic biopsies reveal eosinophilic or pyogranulomatous inﬂammation
without identiﬁcation of an etiologic agent.

Lagenidiosis

Histologically, the cutaneous lesions associated with lagenidiosis are

characterized by eosinophilic granulomatous to pyogranulomatous in-
ﬂammation of the dermis and subcutis. Some cases have demonstrated
coalescing to diﬀuse pyogranulomatous inﬂammation with fewer eosino-
phils, and areas of necrosis and suppuration are observed occasionally.
Multinucleated giant cells and plasma cells are commonly present. In
contrast to P insidiosum, Lagenidium hyphae are usually recognizable on
H&E-stained sections and can be found intracellularly (within giant cells)
and extracellularly (within areas of inﬂammation or necrosis). On GMS-
stained sections, numerous broad, thick-walled, irregularly septate hyphae
are easily visible (Fig. 7A). The diameter of Lagenidium hyphae typically
varies a great deal, even within the same tissue section, but in general the
diameter is much larger than P insidiosum, ranging from 7 to 25 lm, with an
average of 12 lm. In some sections, hyphae appear as round or bulbous
structures (Fig. 7B), and right angle branching is observed occasionally.

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A scant to thin eosinophilic acellular area resembling a thin eosinophilic
sleeve may be noted around the hyphae.

Zygomycosis

The histologic lesions associated with zygomycosis are characterized by

multifocal to coalescing granulomatous inﬂammation inﬁltrated by signiﬁ-
cant numbers of neutrophils, plasma cells, multinucleate giant cells, and
eosinophils. Multiple areas of necrosis often are observed, and ulceration is
common in cutaneous and GI lesions. Conidiobolus and Basidiobolus hyphae
are identiﬁed on H&E-stained sections as clear or slightly basophilic spaces
surrounded by amorphous to granular eosinophilic material within pyo-
granulomas. On GMS-stained sections, hyphae appear broad, thin-walled,
occasionally septate, and infrequently branch at right angles. The histologic
hallmark of zygomycosis is the presence of a wide (2.5 to 25 lm) eosino-
philic sleeve surrounding the hyphae, which makes them easily located on
H&E-stained sections (Fig. 5B) [60]. This ﬁnding helps diﬀerentiate
zygomycosis from pythiosis and lagenidiosis, in which eosinophilic sleeves
tend to be thin or absent. In addition, the hyphal diameter (as measured in
tissue) tends to be signiﬁcantly larger for Basidiobolus species (mean 9 lm;
range, 5 to 20 lm) and Conidiobolus species (mean 8 lm; range, 5 to 13 lm)
than for P insidiosum (mean, 4 lm; range, 2 to 7 lm).

Culture

The culture-based diagnosis of oomycosis and zygomycosis historically

has been problematic. Because the gross lesions associated with pythiosis are
confused easily with those caused by neoplasia or bacterial infection,
veterinarians may fail to submit samples for fungal or oomycete culture.

Fig. 7. Photomicrographs of GMS-stained sections showing Lagenidium species hyphae in
tissue. (A) Broad, thick-walled, infrequently septate hyphae associated with granulomatous
lymphadenitis. (B) Numerous Lagenidium species hyphae of varying diameter (some of which
have a round or bulbous appearance) in an infected canine lymph node; bar

¼ 20 lm.

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If tissues are submitted for culture, sample handling techniques that are
appropriate for fungal-infected tissues may not be conducive to preservation
of oomycete viability. In addition, many diagnostic laboratories are
unfamiliar with culture techniques for oomycetes, and rapid bacterial
growth from specimens with secondary bacterial infection often prevents
recovery of the pathogen. In the author’s experience, isolation of oomycetes
and zygomycetes from infected tissues is not diﬃcult, if appropriate sample
handling and culture techniques are employed.

Pythiosis

A prospective evaluation of sample handling and culture techniques for

the isolation of P insidiosum from equine tissues recently was completed in
the author’s laboratory [61]. Results indicated that for samples that were
stored or shipped for 1 to 3 days before processing, P insidiosum isolation
rates were highest when samples were stored at ambient temperature and
then plated on antibiotic-containing media. When selective media were not
employed, however, isolation rates were higher for samples that had been
stored at 4

C or on cold packs than for those that had been stored at

ambient temperature. These ﬁndings suggest that using techniques designed
to decrease the growth of bacterial contaminants signiﬁcantly improves the
rate of P insidiosum isolation from infected tissues. In the author’s labora-
tory, V8 agar [62] amended with streptomycin (200 lg/mL and ampicillin
(100 lg/mL) is used routinely for the isolation of P insidiosum. As a
commercially-available alternative, the author has found Campy blood agar
(Remel, Inc., Lenexa, KS), which contains trimethoprim, vancomycin,
polymyxin B, cephalothin, and amphotericin B, to be equally eﬀective for
the isolation of P insidiosum.

For best results, small pieces of fresh, nonmacerated tissue should be

placed directly on the surface of the agar and incubated at 37

C. Growth

typically is observed within 12 to 24 hours. Isolation of P insidiosum from
swabs of exudate collected from draining skin lesions is generally un-
successful, but the author occasionally has been able to isolate the patho-
gen from ﬁne needle aspirates of enlarged regional lymph nodes if the sample
is sprayed directly onto the culture plate. Tissue samples submitted for
culture should be wrapped in a sterile saline-moistened gauze sponge and
shipped at ambient temperature to arrive at the laboratory within 24 to 48
hours of collection whenever possible. When samples cannot be processed
for more than 2 to 3 days after collection, however, they should be shipped
with ice packs, stored in the refrigerator, or stored at ambient temperature
in an antibiotic solution.

Identiﬁcation of P insidiosum isolates in the laboratory should be based

on colonial and hyphal characteristics; growth at 37

C; production of

motile, reniform, biﬂagellate zoospores; and, if possible, speciﬁc polymerase
chain reaction (PCR) ampliﬁcation or ribosomal RNA gene sequencing.

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Colonies on V8, Sabouraud/dextrose, or cornmeal agar typically are sub-
merged, white to colorless, and have an irregular radiate pattern [17,63].
Microscopically, hyphae are broad (4 to 10 lm in diameter), hyaline,
sparsely septate, and tend to branch at right angles [17,63]. Zoospores can
be produced readily by placing boiled grass blades on the surface of a 1- to
2-day-old colony growing on 2% water agar, incubating at 37

C for 18 to 24

hours, and then placing the infected grass blades in a dilute salt solution.
After 2 to 4 hours of incubation at 37

C, terminal vesicles, from which

zoospores are released, can be visualized extending from the cut edges of the
infected grass blades (see Fig. 1).

Lagenidiosis and zygomycosis

Isolation techniques for Lagenidium species and the zygomycetes are

similar to those described for P insidiosum, but with slightly diﬀerent media.
Although the zygomycetes (and occasionally Lagenidium species) will grow
on media routinely used for the isolation of pathogenic fungi (Sabouraud/
dextrose agar), isolation is more successful when speciﬁc media are used.
For isolation of Lagenidium species, peptone-yeast-glucose (PYG) agar is
optimal, whereas potato ﬂake agar (PFA) [64], potato dextrose agar, or
cornmeal agar works best for isolation and sporulation of the zygomycetes.
The author routinely uses PYG and PFA amended with ampicillin (100
lg/mL) and streptomycin (200 lg/mL) for initial isolation of oomycetes
and zygomycetes, respectively.

Identiﬁcation of Lagenidium isolates based on morphologic character-

istics is more diﬃcult than for P insidiosum, because zoospores are produced
only occasionally with the techniques described previously, and because
sexual reproductive structures have not yet been identiﬁed. Colonies of
Lagenidium

species on PYG agar are submerged and white to colorless.

Microscopically, mature hyphae are broad (25 to 40 lm in diameter), occa-
sionally to frequently branching or budding, and often appear as segmented
chains of rectangular to oval structures in mature colonies. Because of the
limitations associated with morphologic characterization of this pathogen,
deﬁnitive identiﬁcation of Lagenidium species should be based on speciﬁc
PCR ampliﬁcation or ribosomal RNA gene sequencing [65].

Identiﬁcation of zygomycetes in the laboratory generally is based on

morphologic characteristics of asexual reproductive structures (conidia) and
sexual reproductive structures (zygospores) [66]. Conidiobolus species on
PFA or half-strength cornmeal agar readily produce primary conidia that
are discharged forcibly and can be visualized on the underside of the Petri
dish lid. Primary conidia of Conidiobolus species are spherical, 12 to 40 lm
in diameter, and have a prominent basal papilla (Fig. 8). Older cultures of
C coronatus

may produce primary conidia with hair-like projections (villose

conidia). Because C coronatus is a heterothallic species, zygospores are not
observed in clinical isolates. C incongruus and C lamprauges, however, are

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homothallic species that produce large (12 to 25 lm), round, smooth, thick-
walled zygospores (Fig. 8).

Reproductive structures of B ranarum are produced readily after a 3- to

5-day incubation period on half-strength cornmeal agar. Zygospores are
identiﬁed easily as large (20 to 50 lm), thick-walled, round intercalary
structures with beak-like protuberances that represent the remains of copu-
latory tubes (Fig. 8). Primary conidia, often with a remnant of the conidio-
phore still attached, secondary conidia (morphologically similar to primary
conidia but often smaller), and capilliconidia (oval to elongate spores with
a terminal adhesive knob that develop at the end of a thin supporting
hypha) may be visualized on the inside of the Petri dish lid (Fig. 8).

Molecular assays

Although the identiﬁcation of zygomycetes based on the morphology of

their reproductive structures is straightforward, deﬁnitive morphologic
identiﬁcation of P insidiosum and Lagenidium species isolates is challenging.
To circumvent these diﬃculties, the author recently developed and evaluated

Fig. 8. Photomicrographs illustrating reproductive structures of Basidiobolus and Conidiobolus.
(A) Primary conidia of Conidiobolus species on the inner surface of the lid of a Petri dish. (B)
primary conidium (PC) of B ranarum with a remnant (R) of the conidiophore wall still attached.
(C) Primary conidium (PC) and secondary capilliconidium (SC) of B ranarum on the inner
surface of the lid of a Petri dish. (D) Primary conidia (PC) and a zygospore (Z) of Conidiobolus
species (oblique light microscopy). (E) Zygospore of B ranarum; note the copulatory ‘‘beaks’’
extending from the edge of the round smooth-walled zygospore. (Courtesy of Amy Grooters,
DVM, Louisiana State University)

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PCR-based assays for identifying P insidiosum and Lagenidium species
[30,65]. These assays can be applied to DNA extracted from cultured iso-
lates or from appropriately preserved infected tissue samples. When ap-
plying these assays to the detection of P insidiosum and Lagenidium species
DNA from skin biopsy samples, the author and coworkers found that
freezing samples at

C or storing them at ambient temperature in 95%

ethanol adequately preserved DNA for subsequent ampliﬁcation [67]. Both
PCR assays were designed to amplify small enough products to permit
ampliﬁcation of DNA extracted from sections of paraﬃn-embedded tissue,
which would allow the evaluation of archival samples. Although this
method is technically diﬃcult and labor-intensive because of the steps
needed to prevent microtome-associated carry-over contamination, the
author has used it successfully to conﬁrm the diagnosis of pythiosis in cases
in which paraﬃn-embedded tissues were the only samples available [44].

Serology

Serologic evaluation of pythiosis in people and animals has been used

occasionally as a research tool. Until recently, however, none of these assays
had been tested extensively in small animals or made available for routine
diagnostic purposes. Agar gel immunodiﬀusion (AGID) detects precipitat-
ing antibodies in the serum of most equine and human patients with active
pythiosis, but it is often negative in aﬀected dogs. Western immunoblot
analysis has been used successfully to demonstrate the ability of sera from
Pythium

-infected horses and dogs to recognize antigens of P insidiosum, and

it has the added advantage of high speciﬁcity and sensitivity. Because of
these advantages, immunoblot analysis is the serologic test of choice in the
author’s laboratory for evaluating canine and feline patients suspected of
having pythiosis, lagenidiosis, or zygomycosis. The immunoblot technique is
more time- and labor-intensive, however, than other techniques (such as
ELISA) that also can be used for antibody detection in serum.

As an alternative to immunoblot analysis, the author recently developed

a soluble mycelial antigen-based ELISA assay for the detection of anti-P
insidiosum

antibodies in small animal patients [68]. This assay was found to

be highly sensitive and speciﬁc when used to evaluate serum samples
obtained from 43 dogs with pythiosis, eight dogs with lagenidiosis, 16 dogs
with nonoomycotic fungal or algal infections, 22 dogs with nonfungal GI or
skin disease, and 55 healthy dogs. This assay also has been used for the
detection of antibodies in P insidiosum-infected cats. In addition to
providing a means for early, noninvasive, and speciﬁc diagnosis, the ELISA
also appears to be useful for monitoring response to therapy in aﬀected
patients. In seven dogs and two cats, a dramatic decrease in antibody levels
approaching the range of healthy animals was detected 2 to 3 months
following successful surgical resection of infected tissues. In contrast,
antibody levels remain high in animals that go on to develop clinical

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recurrence following surgical treatment. Therefore, the ELISA appears to be
a promising tool for the early detection of postoperative recurrence, and it
also may be used to guide postoperative medical therapy duration.

Similar ELISA-based assays using soluble mycelial antigens of Lageni-

dium

species, C coronatus, and Basidiobolus ranarum also have been devel-

oped in the author’s laboratory. At this time, insuﬃcient numbers of sera
from zygomycete-infected dogs are available to evaluate the Conidiobolus
and Basidiobolus ELISAs. The Lagenidium ELISA has been evaluated more
fully, however. Unfortunately, the speciﬁcity of this assay is compromised
by cross-reactivity associated with extremely high anti-P insidiosum
antibody concentrations in some dogs with pythiosis. As a result, positive
reactivity in the Lagenidium ELISA can be considered signiﬁcant only when
interpreted in conjunction with P insidiosum serology results. In addition,
the author has observed some degree of anti-Lagenidium activity in the
serum of healthy dogs and horses; therefore, it is likely that a more speciﬁc
antigen will be needed for future development of a Lagenidium-speciﬁc
ELISA. The Lagenidium ELISA has been found to be a useful tool for
monitoring response to medical or surgical therapy in dogs with culture-
conﬁrmed lagenidiosis, however.

Immunohistochemistry

Immunohistochemical techniques based on polyclonal antibodies devel-

oped ﬁrst by Brown [69] and later by Newton [42] have been used regularly
over the past 10 years as conﬁrmatory tests for pythiosis. These techniques
have the advantage of being applicable to paraﬃn-embedded tissues,
permitting the evaluation of archival samples. Although Brown’s antibody
failed to cross react with Conidiobolus and Basidiobolus hyphae in equine
tissues, Newton’s antibody showed some cross-reactive staining when used
to evaluate tissues from dogs with conidiobolomycosis [55]. In addition, the
author observed mild-to-moderate staining of Lagenidium hyphae when
Newton’s antibody was applied to tissues from dogs with culture-conﬁrmed
lagenidiosis [1]. Therefore, the speciﬁcity of this antibody for the im-
munohistochemical diagnosis of pythiosis may be questionable. A new
polyclonal anti-P insidiosum antibody raised in chickens was produced and
evaluated recently in the author’s laboratory [70]. After extensive adsorption
of this antibody with sonicated Lagenidium and Conidiobolus hyphae, it
appears to be highly speciﬁc for the immunohistochemical detection of P
insidiosum

hyphae in tissues. Blinded evaluation of this antibody in a large

number of canine, feline, and equine tissues infected with P insidiosum,
Lagenidium

, Conidiobolus, Basidiobolus, or other fungal organisms has been

completed recently.

Although the author’s laboratory also has produced polyclonal anti-

bodies against Lagenidium species, C coronatus, and B ranarum, adequate

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removal of cross-reactivity from these antibodies without unacceptably
decreasing staining intensity for their intended targets has been elusive. As
an alternative, the author’s laboratory recently has produced 30 Lagenidium-
speciﬁc murine monoclonal antibodies. Three of these antibodies have
proven to be sensitive and speciﬁc tools for the immunohistochemical
detection and identiﬁcation of Lagenidium species hyphae in tissue sections.

Treatment

Aggressive surgical resection remains the treatment of choice for

pythiosis, lagenidiosis, and zygomycosis in small animal patients. Because
it provides the best opportunity for long-term cure, complete resection of
infected tissue should be pursued whenever possible. In animals with GI
pythiosis, segmental lesions should be resected with 3 to 4 cm margins
whenever possible. Although mesenteric lymphadenopathy is almost always
present, P insidiosum hyphae are often absent in enlarged mesenteric nodes.
Therefore, the presence of nonresectable mesenteric lymphadenopathy
should not dissuade the surgeon from pursuing complete resection of
a segmental bowel lesion. In this situation, enlarged lymph nodes should be
biopsied and cultured for prognostic information. Unfortunately, most dogs
with GI pythiosis are not presented to the veterinarian until late in the course
of disease, when complete excision is not possible. In addition, the anatomic
location of the lesion may prevent complete surgical excision when the
esophagus, gastric outﬂow tract, rectum, or mesenteric root are involved.

In animals with cutaneous pythiosis or lagenidiosis in which lesions are

limited to a single distal extremity, amputation is recommended. In animals
with cutaneous or subcutaneous lesions in other areas of the body, the
surgeon should pursue aggressive resection with wide margins. Because dogs
with lagenidiosis often have occult systemic lesions, radiographic imaging of
the chest and abdomen and sonographic imaging of the abdomen are
recommended before attempting surgical resection of cutaneous lesions that
may be caused by pythiosis or lagenidiosis.

Local postoperative recurrence of pythiosis and lagenidiosis is common,

occurring at the site of resection or in regional lymph nodes. For this reason,
medical therapy with itraconazole (10 mg/kg by mouth every 24 hours) and
terbinaﬁne (5 to 10 mg/kg by mouth every 24 hours) is recommended for
at least 2 to 3 months after surgery. To monitor for recurrence, ELISA
serology should be performed before and 2 to 3 months after surgery. In
animals that have had a complete surgical resection and go on to have no
recurrence of disease, serum antibody titers typically drop dramatically
within 3 months of surgery [68]. If this occurs, medical therapy can be
discontinued, with subsequent re-evaluation of serum antibody levels every
3 months for up to 1 year. If antibody levels remain elevated 2 to 3 months
after surgery, medical therapy should be continued, with re-evaluation of
ELISA serology every 2 to 3 months.

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Medical therapy for oomycosis traditionally has been unrewarding. This

is likely because ergosterol (the target for most currently-available anti-
fungal drugs) is generally lacking in the oomycete cell membrane. Despite
this fact, clinical and serologic cures have been obtained following medical
therapy with ergosterol-targeting drugs in a few dogs with P insidiosum
infection, and in a recently-described case in a 2-year-old child [10]. In the
author’s experience over the past 4 to 5 years at Louisiana State University,
approximately 15% of dogs with GI pythiosis have responded to itra-
conazole (10 mg/kg every 24 hours for 6 to 9 months) or amphotericin B
lipid complex (2 to 3 mg/kg administered every other day to a cumulative
dose of 24 to 27 mg/kg). More recently, the author has observed clinical and
serologic improvement or resolution in several of cases of canine and feline
cutaneous pythiosis treated with a combination of itraconazole (10 mg/kg
every 24 hours) with terbinaﬁne (5 to 10 mg/kg every 24 hours). Although
the percentage of animals responding is still poor (less than 20%), based on
the author’s subjective observations, the combination protocol seems supe-
rior to itraconazole or amphotericin B alone. Caspofungin, the ﬁrst anti-
fungal in the newly-developed echinocandin class of b–glucan synthase
inhibitors to gain US Food and Drug Administration (FDA) approval, has
the potential to be a much more eﬀective drug for the treatment of oomyco-
sis because of the large amount of b–glucan present in the oomycete cell
wall. Unfortunately, its extremely high cost makes it unlikely to be used
except on rare occasions.

Recommendations for the treatment of conidiobolomycosis and basidio-

bolomycosis are less straightforward, because these organisms, as true fungi,
should be more likely than oomycetes to respond to medical therapy.
Although anecdotal information and a small number of cases in the litera-
ture [54] support this idea to some degree, progression of cutaneous lesions
and sometimes even dissemination despite treatment, also have been de-
scribed in dogs with zygomycete infection [55]. This may reﬂect variability in
the susceptibility of Conidiobolus and Basidiobolus isolates, as suggested by
results of in vitro susceptibility testing performed in a limited number of
isolates [71,72]. Probably the most appropriate recommendations for the
treatment of cutaneous zygomycosis in small animal patients are aggressive
surgical resection of infected tissues whenever possible, followed by itra-
conazole therapy for 2 to 3 months. If resection is not possible, therapy with
itraconazole or amphotericin B lipid complex should be recommended.

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A.M. Grooters / Vet Clin Small Anim 33 (2003) 695–720

Update on canine and feline

fungal diseases

Marie E. Kerl, DVM

Department of Veterinary Medicine and Surgery, University of Missouri–Columbia,

379 East Campus Drive, Columbia, MO 65211, USA

Systemic fungal infections remain a signiﬁcant cause of disease for dogs

and cats in most regions of the United States. Systemic fungal pathogens
gain entry through a single portal (commonly the respiratory tract) and
disseminate to aﬀect multiple body systems. Antifungal antimicrobials may
be eﬀective for treatment; however eﬃcacy is variable among pathogens,
treatment periods are prolonged, and drugs are costly. This chapter will
focus on clinical signs, diagnosis, and treatment of the most common
systemic mycoses of dogs and cats including blastomycosis, histoplasmosis,
coccidiomycosis, and cryptococcosis, and will review antifungal drugs
currently available for treatment.

Blastomycosis

Blastomycosis is caused by infection with fungal spores of Blastomyces

dermatitidis

, most commonly by way of inhalation and respiratory

colonization. Environmental conditions favoring fungal growth include
moist, acidic soil with decaying vegetation or animal feces. Environmental
moisture is thought to play a major role in dissemination of infective spores
[1]. Geographic regions with the greatest prevalence of blastomycosis include
the Mississippi, Missouri, and Ohio river valleys. The middle Atlantic and
southern states and southern Canadian waterways also have high disease
prevalence, but outbreaks have been reported in other regions also [2–4].

Infection occurs most commonly when an animal inhales conidiophores

from an appropriate environment, but inoculation by penetration also
can cause localized disease (Fig. 1). Dogs and people are aﬀected more
commonly than other species [2,3]. Following inhalation, infective conidia

Vet Clin Small Anim

33 (2003) 721–747

E-mail address:

kerlm@missouri.edu

0195-5616/03/$ - see front matter

doi:10.1016/S0195-5616(03)00035-4

are phagocytized by alveolar macrophages and are transformed from the
mycelial to the yeast phase. Normal body temperatures promote formation
of yeast. The yeast forms are thick-walled structures 8 to 12 lm in diameter
that lack a capsule and bud to form daughter cells with broad-based
attachments (Fig. 2). Yeast may produce a localized infection or may
disseminate hematogenously or by way of lymphatics to distant sites [5].

Fig. 1. Lingual nodular ulceration caused by localized blastomycosis in a dog.

Fig. 2. Cytologic appearance of Blastomyces dermatitidis from a lymph node aspirate. (Wright-
Giemsa stain,

100). (Courtesy of Linda Berent, DVM, Columbia, MO.)

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M.E. Kerl / Vet Clin Small Anim 33 (2003) 721–747

Clinical signs

Dogs

Typical signalment for dogs with blastomycosis includes young adult,

large breeds. Most cases occur in dogs 2 to 4 years of age, with males and
females equally represented. Retrievers and Doberman pinschers were over-
represented in one study [6,7].

Clinical signs develop weeks to months after exposure to the organism.

Anorexia, depression, lethargy, weight loss, cachexia, and fever are com-
mon features of disease. Dissemination occurs by way of lymphatic and
hematogenous spread to colonize distant sites including the eye, skeletal
system, skin, and lymph nodes. Symptoms often are associated with
infection in more than one site at a time. Respiratory signs including
tachypnea, dyspnea, cyanosis, or respiratory distress occur in 65% to 85%
of dogs in association with pulmonary infection. Lymphadenopathy occurs
in 30% to 50% of cases and may be confused with lymphosarcoma if lymph
node aspirate or biopsy is not performed [2,6,7]. Ocular involvement,
manifesting as chorioretinitis, anterior uveitis, retinal detachment, and
secondary glaucoma occurs in 20% to 50% of dogs [8]. Bone involvement is
noticed in 10% to 15% of cases, with lesions most commonly occurring over
epiphyseal regions below the elbow or stiﬂe [5]. Cutaneous signs are
reported in 30% to 50% of cases, including nodules, papules, or plaques of
varying sizes that may drain serosanguineous to purulent exudate.
Paronychia may occur. Calcinosis cutis may be an unusual manifestation
of disease [9]. Central nervous system (CNS) involvement is an uncommon
ﬁnding in dogs with blastomycosis, occurring in fewer than 5% of cases.
Pulmonary thromboembolism may be associated with respiratory blasto-
mycosis [10].

Cats

Blastomycosis is an uncommon fungal disease in cats [3]. It is unclear

whether diﬀerences in age, breed, or sex contribute to likelihood of infection
in cats. Immunosuppression with feline leukemia virus does not seem to
increase predisposition [3]. Clinical signs of disease are similar to dogs, except
cats more commonly exhibit CNS disease and develop large dermal abscesses
[3,5,11].

Diagnosis

Routine screening blood tests of ill patients do not provide deﬁnitive

diagnoses but may show evidence supporting fungal disease. Complete
blood count (CBC) results are often unremarkable but may demonstrate
mild nonregenerative anemia, mature neutrophilia, or neutrophilia with
left shift. Serum biochemical proﬁle is often within reference ranges.

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M.E. Kerl / Vet Clin Small Anim 33 (2003) 721–747

Hypoalbuminemia is the most commonly identiﬁed abnormality, occurring
in approximately 75% of cases; 50% develop hyperglobulinemia, and hy-
percalcemia occurs in 10% of cases [2,7,12].

Thoracic radiographs identify characteristic diﬀuse or nodular interstitial

pattern, alveolar inﬁltrate, or hilar lymphadenopathy in 70% of cases (Fig. 3).
Pleural space abnormalities (ﬂuid or air) are less common. Bone involvement
most commonly aﬀects the appendicular skeleton. Osteolysis with periosteal
proliferation and soft tissue swelling is demonstrated on radiographic exami-
nation. These lesions must be diﬀerentiated from primary osteosarcoma,
which presents with a similar radiographic appearance [2,5,7].

Deﬁnitive diagnosis is made by identifying organisms retrieved from

aﬀected sites by aspirate or biopsy. Site of involvement dictates method of
sampling. Lymph node aspirate of infected nodes or evaluation of exudates
or aspirates from dermal lesions yields organisms reliably. Vitreous aspirate
or ocular histopathology following enucleation frequently provides a diag-
nosis [13]. Respiratory procedures on infected dogs, including lung aspirate,
tracheal wash, or bronchoalveolar lavage, are nondiagnostic at least 50% of
the time [7]. Low yield of these procedures is explained by interstitial location
of the organism. Culture for diagnosis of blastomycosis is not necessary if
cytologic or histopathologic examination demonstrates characteristic organ-
isms. Mycelial growth occurs slowly, and cultures may take several weeks to
become positive [5]. Blastomycosis has zoonotic potential, and caution
should be exercised when handling infected tissues [14,15].

Serologic testing should be considered if multiple attempts to identify the

organism by cytologic or histopathologic examination have failed. A variety
of serologic tests have been developed, none of which will yield a diagnosis
correctly in all cases. Agar gel immunodiﬀusion (AGID) against the

Fig. 3. Lateral thoracic radiograph of a dog with disseminated blastomycosis demonstrating
classic military nodular to diﬀuse pulmonary interstitial inﬁltrate.

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M.E. Kerl / Vet Clin Small Anim 33 (2003) 721–747

A-antigen of Blastomyces dermatitidis is the most commonly used serologic
test, with sensitivity reported to be 60% to 90%, and speciﬁcity up to 96%
[16–18]. AGID is often negative early in the course of disease and may
become negative with treatment or remain positive even with clinical reso-
lution of disease, depending upon the antibody response of the individual
animal. AGID in cats is unrewarding [3]. Recent information on radioim-
munoassay (RIA) testing for WI-1 antigen of B dermatitidis shows a possible
advantage in early diagnosis of blastomycosis in dogs. Sensitivity of this test
was 91%, and speciﬁcity was 100% in dogs from diﬀerent regions of the
United States. The degree of positive titer did not correlate with severity of
disease, and titers remained positive for more than 1500 days following
successful treatment in some dogs. Therefore, this titer would not predict
clinical resolution of disease. This test is not commercially available [18].

Treatment

Although some dogs exposed to blastomycosis clear the organism, dogs

and cats that present with clinical disease will not improve spontaneously
and require therapeutic intervention [2,5]. Approximately 70% to 75%
of dogs that receive treatment with antifungal medications survive. Re-
searchers have found that dogs with severe respiratory infections or multi-
ple body system involvement were more likely to die within the ﬁrst week of
therapy. Brain involvement was signiﬁcantly associated with treatment
failure [19].

Itraconazole (Sporanox) is the treatment of choice for blastomycosis

because of its eﬃcacy, relative safety, and convenience of administration. In
a study of 112 dogs comparing itraconazole with historical controls treated
with amphotericin B (AMB, Fungizone), response and recurrence rates were
similar among all groups [19]. Other treatment options include ketoconazole
(Nizoral), AMB, and lipid-complexed AMB. Ketoconazole is less eﬀective
than itraconazole, with lower response rates, higher relapse rates, and longer
treatment periods [5,20]. AMB has been used successfully to treat blas-
tomycosis. Drawbacks of AMB include parenteral administration and risk of
nephrotoxicity [21]. Lipid-complexed AMB (AmBisome) is eﬀective for
treatment of blastomycosis in dogs, with less risk of nephrotoxicity [22].
Combinations of AMB and itraconazole or ketoconazole may be used in
cases of severe respiratory infection [5]. Table 1 contains dose recommen-
dations.

General medical management of dogs and cats with blastomycosis is

dictated by the location of fungal infection. Supportive therapy for
respiratory involvement includes oxygen therapy in hypoxemic animals,
bronchodilators, and possibly antibiotics, if secondary bacterial infection is
suspected [23]. Ocular involvement may require speciﬁc therapy for anterior
uveitis or secondary glaucoma [5,8,13,24]. Animals with orthopedic lesions
causing lameness and pain should receive analgesic therapy. Dermal wounds

725

M.E. Kerl / Vet Clin Small Anim 33 (2003) 721–747

Table

Drug

ther

apy

for

mmon

syste

mic

mycoses

Species

photericin

(AM

Lipo

somal

AMB

Flucy

tosine

Ketocon

azole

Itracon

azole

Flucon

azole

Blast

omyc

osis

Cani

0.5

mg/kg

/wk;

cumulat

ive

se:

4–6

mg/kg

mg/k

/wk;

cumulat

ive

dose:

mg/kg

5–15

mg/k

for

least

nths,

ith

amph

oter

icin

initially

mg/kg

for

ﬁrst

days,

then

for

60–9

day

days

yond

reso

lution

mg/kg

for

least

days,

days

beyo

resolution

Feline

0.25

mg/kg

/wk;

cumulat

ive

dose:

mg/kg

mg/k

for

least

nths,

ith

amph

oter

icin

initially

mg/kg

for

60–9

day

days

yond

reso

lution

Histo

plasmosis

Cani

0.25–

0.5

mg/kg

/wk;

cumu

la-

tive

dose:

5–10

mg/kg

mg/k

12–2

for

leas

nths,

days

beyond

reso

lution

mg/kg

for

4–6

nths,

day

yond

reso

lution

2.5–5

mg/kg

12–24

for

4–6

months,

days

beyo

resolution

Feline

0.25–

0.5

mg/kg

/wk;

cumu

la-

tive

dose:

4–8

mg/kg

can

ine

rec-

ommen

dation

canine

rec

ommen

datio

See

canine

rec

omme

ndatio

Cryp

tococ

cosis

Cani

0.25–

0.5

mg/kg

/wk;

cumu

la-

tive

dose:

4–10

mg/kg

mg/k

/wk;

cumulat

ive

dose:

8–12

mg/kg

6–8

for

1–12

months

mg/k

12–2

ing

amph

otericin

B/ﬂ

ucytos

ine,

for

4–6

month

5–15

mg/kg

12–24

for

6–10

months,

days

beyo

resolution

726

M.E. Kerl / Vet Clin Small Anim 33 (2003) 721–747

Feline

0.1–0

mg/kg

0.5–0.8

mg/kg

/wk;

cumu-

lative

dose:

4–10

mg/kg

25–5

mg/kg

6–12h

for

1–9

months

canine

rec

ommen

datio

5–10

mg/kg

for

6–10

nths,

days

beyo

resolution

See

canin

rec

omme

ndatio

Coc

cidiomyc

osis

Cani

0.4–0

mg/kg

/wk;

cumu

lative

dose:

8–11

mg/kg

5–10

mg/k

for

8–12

nths

mg/kg

months

mg/kg

month

Feline

cat

12–24

nths

25–50

per

cat

12–2

month

25–5

cat

12–2

month

Monit

for

phrotoxic

ity.

ombine

with

amph

oter

icin

treatmen

Adm

inister

ith

food.

bbrevia

tions:

IV,

intravenou

SQ,

subcutan

eous.

Data

Refs.

[2,5,7

,19,22,23,27,29

,32,4

0,45,51,53

,59].

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M.E. Kerl / Vet Clin Small Anim 33 (2003) 721–747

caused by blastomycosis should be shaved and kept clean and dry.
Personnel handling animals with draining wounds should exercise caution
to avoid accidental infection [14,15].

Treatment with itraconazole or AMB should be continued for a minimum

60 days or at least 1 month beyond clinical or radiographic resolution of
clinical signs. Animals with severe lung involvement should receive therapy
for at least 90 days. Recurrence rates of 20% are reported for dogs treated
with itraconazole for 60 to 90 days or with AMB [19]. Similar treatment
recommendations should be followed for cats, although information about
response and long-term follow-up are lacking [3].

Prognosis

Blastomycosis is associated with an overall mortality rate of 25% to

30%. The prognosis is worse when severe pulmonary involvement is present,
or more than three body systems are involved [19]. Most animals that die
after treatment for blastomycosis do so within the ﬁrst 5 days after therapy
is initiated, likely because of the burden inﬂicted by the sudden death of
many fungal organisms and the subsequent inﬂammatory response.
Pulmonary thromboembolism has been reported as a complication of
blastomycosis that causes sudden deterioration or death [10].

Histoplasmosis

Histoplasmosis occurs as a result of infection with the soil-borne,

dimorphic fungus Histoplasma capsulatum. This organism can survive wide
variations of environmental temperature and is most prevalent in moist soil
containing bird or bat waste. Regions of the United States with greatest
frequency of cases are the Ohio, Missouri, and Mississippi river valleys;
however outbreaks may occur in other regions, if environmental conditions
favor fungal growth [25–27].

Histoplasma capsulatum

has a free-living mycelial stage in soil, with free-

living microconidia (2 to 5 lm) or macroconidia (5 to 18 lm) that serve as
a source for mammalian infection. Histoplasma organisms are 2 to 4 lm in
diameter, with a thin clear halo surrounding a round or crescent-shaped
basophilic cytoplasm. Route of entry is thought to be respiratory in most
cases. Oral exposure may be possible, since some animals have gastrointes-
tinal [GI] signs only [26,28]. With establishment of active infection, dissemi-
nation proceeds to any organ. Lungs, GI tract, lymph nodes, spleen, liver,
bone marrow, eyes, and adrenal glands are aﬀected most commonly. The
incubation period is 12 to 16 days in dogs and people [5]. Exposure to highly
contaminated environments may cause point–source outbreaks in dogs and
people. Cats and dogs are equally likely to develop histoplasmosis [27].

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M.E. Kerl / Vet Clin Small Anim 33 (2003) 721–747

Clinical signs

Dogs

Similar to dogs with blastomycosis, most dogs with histoplasmosis are

large breed, young adults. Males are slightly predisposed, and hunting
breeds including Brittanys, Pointers, and Weimaraners, may be over-
represented [5,26]. Clinical signs are dictated by the organ systems involved;
dogs manifest signs of GI or respiratory disease, but seldom both.
Disseminated histoplasmosis with GI involvement accounts for most
clinical presentations of histoplasmosis [5,27,28]. GI signs are usually
consistent with small and large intestinal diarrhea, and include weight loss,
hypoalbuminemia, intestinal blood loss (melena or hematochezia), and
tenesmus. Hepatosplenomegaly occurs in up to 50% of cases (Fig. 4) [28].
Coughing, tachypnea, dyspnea, or pleural eﬀusion occurs with pulmonary
involvement. Less speciﬁc ﬁndings include fever, anorexia, depression, and
severe weight loss. Unlike blastomycosis, however, histoplasmosis seldom is
associated with bone, ocular, or dermal lesions [27].

Cats

Unlike blastomycosis, histoplasmosis is as likely to occur in cats as in

dogs. Histoplasmosis occurs most commonly in cats younger than 4 years of

Fig. 4. Dorsoventral abdominal radiograph of a dog with disseminated histoplasmosis causing
diﬀuse hepatomegaly.

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M.E. Kerl / Vet Clin Small Anim 33 (2003) 721–747

age, with no breed or sex predilection [25,27]. Infection with feline leukemia
virus is not associated with increased occurrence of infection [25]. Clinical
signs are those of nonspeciﬁc disease, including weight loss, depression,
fever, anorexia, and anemia. Weight loss and emaciation are common
ﬁndings. Unlike dogs, speciﬁc GI signs are identiﬁed less commonly.
Pulmonary involvement results in clinical signs of dyspnea, tachypnea, or
abnormal lung sounds. Lymphadenopathy and hepatosplenomegaly occur
with dissemination. In the author’s experience, bone marrow involvement
can occur frequently, and can be associated with various cytopathies.
Dermal, ocular, orthopedic, and oral lesions occur uncommonly. Bone
involvement may cause osteolytic lesions of the distal appendicular skeleton
and result in lameness [5,27].

Diagnosis

As with other fungal infections, there are no pathognomonic ﬁndings on

routine laboratory evaluation for histoplasmosis. Nonregenerative anemia is
the most common ﬁnding on CBC. Causes include chronic inﬂammation,
GI blood loss, and bone marrow infection [5]. Histoplasma organisms are
seen rarely on CBC. Thrombocytopenia may occur. Abnormalities on serum
biochemical proﬁle include hypoalbuminemia, elevated hepatic enzymes,
total bilirubin, and hypercalcemia [5]. Thoracic radiographs reveal diﬀuse or
nodular interstitial pattern, or hilar lymphadenopathy in animals with
pulmonary involvement [27]. Abdominal ultrasound is helpful to evaluate
organomegaly (Fig. 5).

Deﬁnitive diagnosis is established by identiﬁcation of H capsulatum on

cytologic or histopathologic evaluation. Organisms typically are found
within cells of the mononuclear phagocyte system. Cytologic diﬀerentiation
from other fungal pathogens is facilitated by the fact that Histoplasma
capsulatum

organisms are much smaller and typically are clustered within

cells (Fig. 6). Diagnostic samples may be obtained from a variety of
locations. Rectal mucosal scraping in dogs with GI involvement is
frequently diagnostic for histoplasmosis. For disseminated disease, aspira-
tion of lymph nodes, dermal nodules, bone marrow, liver, spleen, or
endotracheal wash should be considered. Tissue biopsy and histopathology
may demonstrate organisms if cytology is unrewarding.

Serologic tests to diagnose histoplasmosis are unreliable, with false-

negatives occurring in active disease and false-positives occurring in animals
without active disease. No reliable immunodiagnostic test is available for
identiﬁcation of histoplasmosis in companion animals [27].

Treatment

Although pulmonary histoplasmosis may resolve spontaneously, treat-

ment is recommended to prevent dissemination from occurring early in the

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M.E. Kerl / Vet Clin Small Anim 33 (2003) 721–747

course of disease. Similar to treatment for blastomycosis, itraconazole is the
treatment of choice for histoplasmosis [27,29]. In a case series of cats with
histoplasmosis, itraconazole was more eﬀective than ketoconazole, with
fewer adverse eﬀects [29]. In dogs, itraconazole is also likely the treatment of
choice; however it has not been studied extensively. GI drug absorption has
not been predicted accurately in animals with GI or disseminated
histoplasmosis. Fluconazole (Diﬂucan) has better penetration into the eye

Fig. 5. Ultrasonographic image demonstrating bright hepatic parenchyma and hepatomegaly
as evidenced by liver contacting dorsal and lateral surfaces of the right kidney.

Fig. 6. Cytologic appearance of Histoplasma capsulatum from hepatic aspirate (Wright-Giemsa
stain,

100). (Courtesy of Linda Berent, DVM, Columbia, MO.)

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M.E. Kerl / Vet Clin Small Anim 33 (2003) 721–747

and CNS than itraconazole, but in people with histoplasmosis, ﬂuconazole
is less eﬀective than itraconazole [27]. This drug has not been studied
extensively for treatment of histoplasmosis in dogs and cats [27].
Fluconazole should be considered with CNS involvement, or in individuals
refractory to treatment with AMB and itraconazole. With severe GI or
disseminated disease, treatment with parenteral AMB combined with
itraconazole or high-dose itraconazole has been recommended for more
rapid control of the fungal disease [27].

Treatment should be continued for at least 60 days, or until 1 month

following resolution of clinical signs. Complete resolution of GI or
disseminated histoplasmosis is diﬃcult to determine, and serologic testing
cannot help identify response to therapy. Animals that experience relapse
with discontinuation of therapy should resume antifungal drug treatment.
Table 1 contains dosing recommendations.

Ancillary therapy for GI histoplasmosis includes dietary modiﬁcation for

small or large bowel disease (highly digestible diet for small intestinal
disease, increased ﬁber diet for large intestinal disease) and antibiotic
therapy to control concurrent small intestinal bacterial overgrowth. Anti-
diarrheal therapy may be helpful in conjunction with antifungal therapy for
symptomatic relief [5,27]. If malabsorption is causing malnutrition in severe
GI histoplasmosis, nutritional support with partial or total parenteral
nutrition should be considered until normal GI function resumes.
Concurrent respiratory therapy for animals with pulmonary involvement
includes oxygen and bronchodilator therapy for hypoxemic patients,
minimizing handling that exacerbates respiratory distress, and possibly
short-term anti-inﬂammatory corticosteroids [23,30].

Prognosis

Statistics on mortality with histoplasmosis have not been reported for

dogs since the advent of itraconazole therapy. In one report, eight cats that
had failed initial treatment with ketoconazole were cured with itraconazole
[29]. In the author’s experience, prognosis is guarded to good depending
upon the nature of systemic involvement of organ systems.

Coccidiomycosis

Coccidiomycosis is a systemic fungal infection caused by Coccidioides

immitis

, a soil saprophyte that grows in areas with sandy, alkaline soils and

semiarid conditions. In the environment, C immitis grows as a mycelium
with thick-walled, barrel shaped arthroconidia, 2 to 4 lm wide and 3 to
10 lm long. Following exposure by inhalation, the arthroconidia enlarge
to form a spherule 20 to 200 lm in diameter. (Fig. 7). The disease has
been reported in most mammals. In companion animals, dogs are more
frequently infected than cats [31,32].

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M.E. Kerl / Vet Clin Small Anim 33 (2003) 721–747

Growth of C immitis is localized to regions in the lower Sonoran life

zone. This includes the southwestern United States, Mexico, and Central
and South America. A pattern of increased rainfall followed by drought
conditions can cause epidemics. Dust storms or earthquakes facilitate
spread. Coccidiomycosis also is known as valley fever or San Joaquin valley
fever, so named for regions in which the disease is reported frequently [5,32].

Exposure and infection occur by way of the respiratory route; however,

direct inoculation of infective spores can cause localized subcutaneous
infection [31,32]. Inhaled arthrospores migrate through the pleural tissue to
the subpleural space. Incubation period ranges from 1 to 3 weeks in dogs.
An intense inﬂammatory response develops, resulting in clinical respiratory
signs. The disease most commonly involves the respiratory tract. If
dissemination occurs, involvement of other organ systems may be found
in bones, eyes, heart, pericardium, testicles, brain, spinal cord, and visceral
organs. People are considered immune following resolution of infection;
however the same is not known for dogs and cats [31,32].

Clinical signs

Dogs

Similar to other systemic mycoses, young adult large breed dogs that are

housed outdoors seem to be predisposed. Breeds that may be at increased
risk for infection include the boxer, pointer, Australian shepherd, beagle,
Scottish terrier, Doberman pinscher, and cocker spaniel [5]. Symptoms in
the dog are primarily respiratory and may be inapparent or manifest only as
mild respiratory signs following exposure. In a small subset of animals,
immune response is ineﬀective, and clinical signs become more severe.
Chronic cough is the most common presenting complaint. The cough may

Fig. 7. Histopathologic appearance of Coccidioides immitis (Hematoxylin & eosin stain).
(Courtesy of Susan Turnquist, DVM, Columbia, MO.)

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M.E. Kerl / Vet Clin Small Anim 33 (2003) 721–747

be dry if it occurs as a result of hilar lymphadenopathy secondary to fungal
infection, or it may be moist and productive with alveolar involvement.
Respiratory lesions may progress to severe pneumonia. Fever, weight loss,
and anorexia are common ﬁndings with clinical disease [5,32].

In addition to respiratory involvement, coccidiomycosis may become

disseminated. Osseous lesions occur in 65% of dogs and may be associated
with draining skin nodules over the lesions [5]. Regional lymphadenopathy
associated with appendicular skeletal lesions may be seen, but peripheral
lymphadenopathy is considered rare. Renal or GI systems may become
involved. Myocardial or pericardial infection may occur, causing cardiac
arrhythmias or restrictive pericarditis, resulting in right or left-sided heart
failure [32,33]. CNS infection may cause seizures, behavior change, or coma.
Granulomatous meningoencephalitis has been associated with coccidiomy-
cosis [34]. Ocular lesions involving the anterior and posterior segments
occur; however ocular involvement is less common with coccidiomycosis
than with other systemic fungal infections [8].

Cats

Cats are relatively immune to coccidiomycosis compared with dogs.

There is no obvious age, breed, or sex predilection for coccidiomycosis in
cats. Skin lesions from dermal inoculation with fungus are the most
common presentation. Lesions may form masses or be associated with
abscessation or drainage [35]. Localized lymphadenopathy associated with
draining lesions may occur. Fever, inappetence, and weight loss commonly
occur in aﬀected cats. Respiratory symptoms similar to those seen in dogs
occur in about 25% of aﬀected cats [35]. Ocular involvement including
retinal detachment and uveitis or iritis occurs in approximately 12% of cats
[11,35]. CNS localization is uncommon.

Diagnosis

Characteristics of CBC and serum biochemical proﬁle are suggestive of

chronic inﬂammatory disease. Mild, normocytic, normochromic, non-
regenerative anemia, neutrophilia, left shift, and monocytosis may be
identiﬁed on evaluation of CBC. Serum biochemical proﬁle results vary
depending upon organ systems involved; hypoalbuminemia and hyper-
globulinemia are common; hepatic transaminase elevation occurs with
hepatic involvement, and azotemia occurs with renal involvement. Hyper-
calcemia has been reported, but is less common than with other fungal
diseases [5].

Thoracic radiographs often reveal diﬀuse interstitial or peribronchilar

pattern, frequently with hilar lymphadenopathy. Alveolar inﬁltrate some-
times occurs (Fig. 8). Pleural involvement occurs in 65% of cases, either as
pleural eﬀusion or ﬁbrosis and thickening. Hypertrophic osteopathy of long
bones may occur with pulmonary involvement [32].

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M.E. Kerl / Vet Clin Small Anim 33 (2003) 721–747

Demonstration of the oﬀending organism by cytologic or histopathologic

examination provides a conclusive diagnosis for coccidiomycosis. Because
of the relatively low numbers of organisms, location of lesions, or
invasiveness of procedures necessary for obtaining samples, organism
identiﬁcation is often diﬃcult, however. Evaluation of pleural ﬂuid or pus
from draining skin nodules aﬀords the greatest frequency of organism
identiﬁcation. Tracheal wash, bronchoalveolar lavage, lung aspirate, or
lymph node aspirate yield organisms less commonly, and bone aspirate is
largely unrewarding. Lung biopsy and histopathology may reveal organisms
in pulmonary microabscesses; obtaining samples from more than one site is
recommended [5,32].

Fungal culture is not a clinically useful tool for diagnosis of coc-

cidiomycosis. C immitis grows readily on commercial agars at room tem-
perature; however deﬁnitive identiﬁcation requires inoculation into animals
to induce spherule formation. Laboratory personnel must exercise pre-
cautions to prevent accidental exposure and infection [36].

Serologic diagnosis of coccidiomycosis is more rewarding than for blasto-

mycosis or histoplasmosis. If symptoms are consistent with coccidiomycosis,

Fig. 8. Ventrodorsal thoracic radiograph of a dog with coccidiomycosis causing focal alveolar
inﬁltrate of the right caudal lung lobe.

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M.E. Kerl / Vet Clin Small Anim 33 (2003) 721–747

and organisms cannot be identiﬁed, a variety of serologic tests may be used to
detect IgM and IgG antibodies. Compliment ﬁxation (CF) testing represents
IgG antibodies and should become positive within 4 to 6 weeks after
exposure in clinical infection. Tube precipitin (TP) testing represents IgM
antibody response. The TP test will become positive earlier in the course of
infection (2 weeks after exposure) and becomes negative within 4 to 6 weeks
[32]. CF titers generally increase with severe or disseminated disease. Low
titers (less than 1:16) are suggestive of early or past infection, while titers of
1:32 or greater are consistent with active infection. Other methods of
serologic testing have been reported for people, with varying rates of success
[37]. In cats, CF and TP testing can become positive, and can remain positive
for long periods of time even with treatment of disease. [35] Serologic testing
should be interpreted in light of clinical signs consistent with active infection
to conﬁrm diagnosis when attempted visualization of fungal organisms is
unrewarding. Repeat CF testing in 2 to 4 weeks to demonstrate increasing
titer is warranted in questionable cases [32].

Treatment

Coccidiomycosis is diﬃcult to cure compared with other fungal

infections, and lifelong therapy may be necessary. Commonly recommended
treatments for dogs and cats include azole antibiotics (ketoconazole,
itraconazole, and ﬂuconazole), and AMB; however, controlled therapeutic
trials with these agents are lacking. Respiratory infection can be cleared
spontaneously by the host immune response; therefore debate exists over the
criteria indicated to initiate prolonged therapy with expensive and
potentially toxic medication [5,32]. Early initiation of therapy in primary
respiratory coccidiomycosis may be appropriate, since dissemination is
possible. The decision to discontinue therapy is based upon resolution of
clinical signs and resolution of elevated titers; CF titers may become
negative, or may remain positive at 1:2 to 1:4.

Ketoconazole traditionally has been the drug of choice in dogs and cats

for treatment of coccidiomycosis. Table 1 contains dose recommendations.
Serologic testing should be repeated in 4 to 6 weeks of initiation of therapy.
If the titer is increasing, or clinical signs deteriorating, alternative therapy
should be chosen. Treatment may need to be continued for 8 to 12 months
[5,32].

Itraconazole may be an alternative to ketoconazole with fewer adverse

eﬀects; however eﬃcacy remains undetermined for dogs and cats. Some
animals seem to have a more favorable response, while others require
change in therapy to ketoconazole following unsuccessful itraconazole
treatment [32]. AMB is indicated in animals that cannot tolerate the adverse
eﬀects of azole drugs. Liposome-encapsulated AMB formulations may have
fewer adverse eﬀects while retaining eﬃcacy; however, they have not been
studied for this disease [32].

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M.E. Kerl / Vet Clin Small Anim 33 (2003) 721–747

Relapse is common after discontinuation of therapy, particularly in cats

[35]. Therapy duration generally is recommended for months. Decisions to
discontinue therapy should be based upon resolution of clinical signs and
serologic testing. Positive CF test results are not unusual even with
treatment; however increasing titer can be interpreted as treatment failure or
relapse following discontinuation of therapy [32].

Chitin synthesis inhibitors interfere with fungal cell wall formation. In

addition, they are cidal and may require relatively low doses for shorter
periods of time than azole antibiotics. This class of drug is under inves-
tigation in people as antifungal therapy [38]. Lufenuron (Program) is a chitin
synthesis inhibitor approved for veterinary use to control ectoparasites. A
case series of dogs with coccidiomycosis treated with daily doses of
lufenuron showed clinical improvement in 1 week, and resolution of
radiographic lesions occurred in 8 weeks [32,39]. The main drawback to
lufenuron therapy is drug cost; however, this might be oﬀset if the treatment
duration is shorter than with traditional antifungal therapy. Lufenuron is
not approved for this use [39].

Prognosis

Coccidiomycosis remains a challenge to treat and is diﬃcult to cure

compared with other systemic mycoses. Localized respiratory infections may
resolve spontaneously and generally carry a good prognosis. Disseminated
infections will result in death if not treated. An overall recovery rate of 60%
has been noted with ketoconazole therapy; however, multiple bone or CNS
involvement carries a worse prognosis [32].

Cryptococcosis

Cryptococcosis is caused by a variety of species of Cryptococcus.

Cryptococcus neoformans

, which thrives at normal body temperature, is

the most clinically signiﬁcant agent. Unlike other fungal infections,
cryptococcosis does not occur in a deﬁned geographic region [40]. C
neoformans

is a saprophytic, round, yeast-like fungus 3.5 to 7 lm in

diameter, with a large heteropolysaccharide capsule of 1 to 30 lm that does
not uptake common cytologic stains (Fig. 9) [41]. Cryptococcus reproduces
by budding from the parent cell. Buds can break oﬀ at diﬀerent stages of
growth, resulting in size variation of organisms in the tissues. Most likely
environmental sources are locations near avian habitats or in leaf and bark
litter of eucalyptus trees. The pigeon is thought to be the most important
vector; the elevated body temperature of the pigeon is thought to protect it
from disease. Cryptococcosis has become an important health problem in
people with HIV and AIDS [42].

Unlike other fungal infections, cryptococcosis occurs with equal or

greater frequency in cats compared with dogs [43]. The most likely route for

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M.E. Kerl / Vet Clin Small Anim 33 (2003) 721–747

infection is the respiratory tract. Cryptococcus is unencapsulated in the en-
vironment and may be as small as 1 lm, enhancing respiratory colonization.
Following deposition in tissues, organisms colonize the upper or lower
respiratory tract, where they regenerate their capsules. The capsule interferes
with the normal host immune response and organism elimination. CNS
involvement is common in cats and dogs and may occur as a result of
hematogenous spread or extension from nasal cavity disease [44–47].

In people, natural disease resistance is strong, and infection is a sign of

immunosuppression [42]. In dogs and cats, corticosteroid therapy has
exacerbated infection [41]. In cats, concurrent feline leukemia virus (FeLV)
and feline immunodeﬁciency virus (FIV) infections decrease the likelihood
of favorable response to treatment [48]. It is unknown whether cats with
FeLV or FIV are predisposed to cryptococcosis. Immunosuppression in
dogs has not shown to increase risk for cryptococcosis.

Clinical signs

Cats

Cryptococcosis is the most common systemic fungal infection of cats [43].

There is no obvious age or sex predilection. Siamese cats are over-
represented in some studies [49]. Upper respiratory infection is evident in
50% to 60% of cases [41,48,50]. Symptoms may include nasal or facial
deformity, mass protruding from nares, nasal discharge, sneezing, respira-
tory noise, or change of voice (Fig. 10). Skin lesions occur in 40% to 50% of
cases, and ocular and CNS signs occur in approximately 15% of cases [43].
Ocular signs consist of blindness caused by retinal detachment and
granulomatous chorioretinitis [11]. Neurologic symptoms include depres-
sion, temperament changes, ataxia, vestibular signs, and blindness [40,46].

Fig. 9. Cytologic appearance of Cryptococcus neoformans. (Wright-Giemsa stain,

100).

(Courtesy of Linda Berent, DVM, Columbia, MO.)

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M.E. Kerl / Vet Clin Small Anim 33 (2003) 721–747

Lungs are aﬀected less commonly, and cats typically do not show signs of
lower respiratory tract disease. Nonspeciﬁc ﬁndings of weight loss,
listlessness and anorexia are common with chronic disease.

Dogs

Aﬀected dogs are generally younger than 4 years of age, although an

occasional older dog may become aﬀected. Great Danes, Doberman

Fig. 10. (A) Nasal swelling caused by cryptococcosis in a cat. (B) Same cat following successful
therapy with itraconazole.

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M.E. Kerl / Vet Clin Small Anim 33 (2003) 721–747

pinschers, Labrador retrievers and American cocker spaniels are over-
represented [5,40]. Clinical signs most often are localized to the CNS, with
presentations of seizure, ataxia, central vestibular disease, papilledema,
cervical pain, tetraparesis, or multifocal cranial nerve involvement [45].
Dogs also may have ocular lesions to include granulomatous chorioretinitis,
retinal hemorrhage, and optic neuritis [8]. Skin lesions, fever, and peripheral
lymphadenopathy may be seen in occasional cases [40]. Weight loss and
lethargy are common but nonspeciﬁc ﬁndings.

Diagnosis

Results of CBC and serum biochemical proﬁle are usually unremarkable

with cryptococcosis. Nonregenerative anemia, neutrophilia, or left shift may
be identiﬁed. Thoracic radiographs occasionally reveal nodular interstitial
inﬁltrates, hilar lymphadenopathy, or pleural eﬀusion. Skull radiographs or
CT can demonstrate nasal bone destruction and soft tissue swelling [5].
With CNS involvement, cerebrospinal ﬂuid (CSF) evaluation commonly
exhibits increased protein and mixed mononuclear and neutrophilic
pleocytosis [45].

The most reliable method to establish diagnosis of cryptococcosis is

direct visualization of the causative organism on cytologic or histopatho-
logic evaluation of specimens from an aﬀected area. Cytologic examination
may be performed on nasal discharge, skin exudates, CSF, tissue aspirate, or
samples obtained by ocular paracentesis. Dogs may have subclinical renal
infection; therefore microscopic evaluation of urine sediment is warranted.
Wright’s stain may cause some distortion of Cryptococcus species, while
Gram’s stain may facilitate visualization [40].

Serologic testing is available and useful to compliment other diagnostic

procedures. Recommended serologic testing consists of latex agglutination
(LA) testing to identify cryptococcal capsular antigen. This is an important
distinction from serologic testing for other fungal infections, which relies on
evaluation of antibody response of the infected individual. Antigen testing is
considered positive at a titer of 1:16 or greater. Response to treatment is
correlated with declining titer results [40,50]. Testing for cryptococcal
antigen in CSF can conﬁrm diagnosis of dogs with CNS involvement when
the organism cannot be demonstrated in CSF [44].

Histopathology of aﬀected tissue is indicated if cytology is negative;

however, impression smears always should be made of biopsy samples
because of the comparative ease of cytologic diagnosis. Histopathologically,
the large capsule diﬀerentiates Cryptococcus from Blastomyces, while
budding and lack of endospores diﬀerentiate it from C immitis.

Fungal isolation is available if deﬁnitive diagnosis cannot be achieved by

other means, but it is not clinically useful because of prolonged growing
time (up to 6 weeks may be required for positive cultures). Interpretation

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M.E. Kerl / Vet Clin Small Anim 33 (2003) 721–747

demands caution, since results from nasal swabs may be positive in dogs and
cats without any clinical evidence of disease [5]. CSF culture may be
appropriate if CNS cryptococcosis is suspected, and cytology is negative.
Culturing large volumes (10 to 15 mL) of CSF may enhance diagnosis, but
this is prohibitive in small animal species. [40,45].

Treatment

Several dosing protocols and regimens have been developed for treatment

of cryptococcosis in dogs and cats; choice of therapy depends upon available
drugs, location of infection, and adverse eﬀects in an individual [5].
Itraconazole therapy has been shown to be curative in 57% of cats [51].
Ketoconazole also has been shown to cure infection; however, deleterious
adverse eﬀects occurred more frequently with ketoconazole than with
itraconazole [52]. Subcutaneous AMB has been used with ﬂucytosine or
ketoconazole in feline and canine cryptococcosis [53].

Therapy duration is generally prolonged. Azole antifungals typically are

administered for 6 to 10 months [40]. The decision to discontinue therapy
should be based on resolution of clinical signs. Recommendations to
discontinue therapy 1 month after resolution of clinical signs, and decrease
in antigen titer by two orders of magnitude or until negative have been
proposed for cats [40]. Table 1 contains dose recommendations.

Prognosis

Cats have a good prognosis when disease occurs outside of the CNS [43].

Progressive decrease of antigen titer by tenfold over 2 months has been
associated with favorable prognosis in cats. Dogs with any form of disease,
and cats with CNS disease, have a guarded prognosis [5,44].

Drug therapy for systemic mycoses

A limited number of drugs are available to treat fungal infections.

Antifungal drugs are expensive; treatment protocols dictate long-term
therapy for cure or control of systemic mycoses, and rate of drug toxicities is
relatively high. Cell-mediated immunity is crucial for host defense against
systemic mycoses. Without a functional immune system, fungal infections
occur more commonly, and deﬁnitive cure with any therapy is diﬃcult or
impossible [42,48].

Mainstays of antifungal therapy include polyene antibiotics and azole

derivatives. Supportive care for dogs and cats with fungal disease dictates
administration of a variety of therapeutics in addition to antifungal drugs.
Medications with known nephrotoxicity should be avoided with concurrent
AMB administration. Drugs that are metabolized by the hepatic p450

741

M.E. Kerl / Vet Clin Small Anim 33 (2003) 721–747

enzyme system (especially histamine-2 receptor antagonists) may delay
metabolism of azole antifungals, especially ketoconazole, thereby resulting
in higher plasma drug concentrations [54]. Some clinicians have used this
drug interaction to delay metabolism of ketoconazole in an eﬀort to
administer a lower, and therefore less costly, dose of ketoconazole in large
breed dogs.

The use of corticosteroids to treat animals with fungal disease is

controversial. There is no question that administering corticosteroid drugs
can lead to dissemination and worsening of fungal infection. Corticosteroids
profoundly impair the cell-mediated immunity that is crucial for protection
from fungal infection, and even to facilitate clearance of infection in animals
treated with antifungal drugs [55]. Corticosteroids are also potent anti-
inﬂammatory medications, however [56]. Much of the morbidity and
mortality that accompanies treatment of fungal disease results from massive
inﬂammation as a response to the death of fungal organisms within the ﬁrst
week of treatment. For animals with respiratory compromise resulting from
pulmonary fungal infection, many clinicians administer anti-inﬂammatory
dosages of corticosteroids either simultaneously with the instigation of
antifungal medication, or if respiratory signs worsen within the ﬁrst few days
of antifungal therapy. In either case, steroids are used in conjunction with
antifungal medications and are continued for only a brief time, typically 1 to
2 weeks [30].

Polyene antibiotics useful for treating systemic mycoses include AMB and

lipid-complexed AMB. AMB is a polyene macrolide antibiotic produced by
the microorganism Streptomyces nodosus, and it is considered the standard
by which other antifungal therapies are judged [5,57]. GI absorption is poor;
therefore AMB must be administered parenterally. Following intravenous
(IV) administration, AMB is highly protein-bound. It then redistributes
from the blood to the tissues. Metabolic pathways of AMB are unknown.
Biphasic elimination occurs, with an initial half-life of 2 to 4 days, and
a terminal half-life of 15 days [54]. Only a small amount undergoes renal and
biliary elimination. CNS penetration is poor. AMB binds to sterols, in-
cluding ergosterol in fungal cell membranes, to increase permeability and
eventually cause cell death. Aﬃnity is greater for ergosterol in fungal cell
membranes than for cholesterol found in mammalian cell membranes;
however, aﬃnity for cholesterol explains AMB toxic eﬀects. Subcutaneous
administration at higher doses has been used in an attempt to delay
absorption and reduce toxicity [57]. In animals with severe GI fungal disease
and poor drug absorption, polyene antifungals administered parenterally
may be preferred to oral therapies.

Typical dosing protocols for AMB include intermittent administration

until a cumulative dose has been achieved, with interruption of therapy in
the event of azotemia. Cats typically receive lower intermittent and
cumulative doses than dogs (Table 1). Bolus IV administration is possible,
but occurrences of nephrotoxicity can be reduced if AMB is infused in 5%

742

M.E. Kerl / Vet Clin Small Anim 33 (2003) 721–747

dextrose and administered over 1 to 5 hours. Serum urea nitrogen (BUN)
and urine sediment evaluation should be measured before administration of
each dose. Identiﬁcation of tubular casts in urine sediment is an earlier
indicator of ongoing renal tubular damage than serum biochemical tests,
and the treatment regimen should be altered. With BUN greater than 50
mg

/dL, the drug should be discontinued until azotemia resolves [5].

Administration of 0.9% IV saline before AMB administration decreases
the incidence of nephrotoxicity in people [58].

Lipid-complexed AMB drugs are available, and these are signiﬁcantly

less nephrotoxic than AMB. Three formulations are approved for use in
people: AMB lipid complex (ABLC, Abelcet), AMB colloidal dispersion
(Amphotec), and liposome-encapsulated AMB (AmBisome) [59]. The
advantage of these preparations is the ability to administer higher
intermittent and cumulative doses with less nephrotoxicity. There are
relatively few head-to-head comparisons of these drugs, so comparisons of
eﬃcacy are diﬃcult [60]. Abelcet has been used successfully to treat
blastomycosis in dogs [22]. The disadvantage of these drugs is increased cost
compared with AMB.

Flucytosine (Ancobon) is a pyrimidine originally developed as an

antineoplastic agent for people [57,60]. Although this drug was ineﬀective
for that use, antifungal activity was discovered in 1973. The drug is taken
up by the fungal cell and converted to 5-ﬂuorouracil, which then interferes
with DNA and protein synthesis. Drug resistance develops rapidly. It has
been used in combination with AMB as a treatment for cryptococcosis
before the availability of newer azole antifungals [53]. Toxicities include
dermal eruptions in dogs, and hematologic changes at high doses in people
[57,60].

The azole antifungals include ketoconazole, itraconazole, and ﬂucon-

azole. Azole antifungals act by inhibiting the fungal P450 enzyme necessary
for development of ergosterol [60]. Itraconazole and ﬂuconazole were ap-
proved by the US Food and Drug Administration (FDA) in the early
1990s, and they have become mainstays of therapy for veterinary systemic
mycoses. All are administered orally, and peak plasma concentrations do
not occur for 6 to 14 days after initiating treatment with azole antifungals.
Ketoconazole and itraconazole are weak bases, lipophilic, and protein-
bound. Absorption is improved in an acid environment, and uptake may be
impaired with concurrent use of antacids or H2 receptor antagonists.
Distribution occurs through most tissues except the CNS and urine.
Fluconazole is minimally protein-bound and highly water soluble. It crosses
the blood-brain, blood-ocular, and blood-prostate barriers well [5,57].
Dosing of ﬂuconazole should be adjusted in animals with reduced
glomerular ﬁltration rate (GFR) [57].

Ketoconazole has been eﬀective as a sole agent for treatment of systemic

mycoses, but in general it is less eﬃcacious than AMB. In serious systemic
fungal infections, combination therapy with AMB and ketoconazole may

743

M.E. Kerl / Vet Clin Small Anim 33 (2003) 721–747

allow reduced dosage and toxicity of AMB while still maintaining eﬃcacy.
Adverse eﬀects of ketoconazole therapy include GI upset, which may be
reduced by administering it with meals and dividing the dose into multiple
smaller doses daily. Hepatic transaminases and alkaline phosphatase
elevations may occur, as well as a clinical hepatitis that may be fatal
[54,57]. Ketoconazole also can suppress testosterone and cortisol synthesis,
and it has been used as a treatment for pituitary-dependent hyperadrenocor-
ticism [61].

Itraconazole has been eﬀective as a sole treatment agent in blastomycosis,

histoplasmosis, and cryptococcosis. It can be used to treat coccidiomycosis;
however, some dogs with this infection fail to respond to this drug and have
a more favorable response to ketoconazole. Absorption is most consistent
with administration following a full meal. Itraconazole selectively inhibits
fungal P450 enzymes and not mammalian enzymes; therefore toxicities
occur less frequently. Mild elevations of hepatic transaminase activity can
occur [29,57]. Cutaneous reactions consisting of localized ulcerative
dermatitis and vasculitis occur in a small percentage of dogs receiving
itraconazole treatment; dermal lesions resolve following discontinuation of
therapy [5].

Fluconazole crosses the blood–brain barrier better than the other azole

antifungals, and it has more consistent oral absorption on an empty
stomach. Therefore, it would be indicated for CNS involvement in systemic
mycoses and for anorexic animals. Because of excellent CNS penetration,
ﬂuconazole is the treatment of choice for people with meningeal
coccidiomycosis, and it should be considered in canine and feline meningeal
coccidiomycosis. Fluconazole has been used to treat cryptococcosis
successfully in cats [49]. Fluconazole crosses the blood–ocular barrier better
than itraconazole, but itraconazole has been used successfully to treat ocular
histoplasmosis in cats [29].

Summary

Systemic fungal diseases cause signiﬁcant morbidity and mortality in dogs

and cats. Blastomycosis, histoplasmosis, coccidiomycosis, and cryptococ-
cosis represent the four most common systemic fungal diseases. Young
adult, large breed dogs generally are predisposed; cats usually do not have
predictable predispositions. Intact cell-mediated immunity is essential to
initial resistance to infection and response to treatment in animals. Several
body systems can be aﬀected. Diagnosis can be conﬁrmed on the basis of
clinical signs and demonstration of the causative organism. Serology is
helpful with coccidiomycosis and cryptococcosis. Treatment is complicated
by limited availability of fungicidal antimicrobials and the necessity of long-
term treatment with expensive drugs.

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M.E. Kerl / Vet Clin Small Anim 33 (2003) 721–747

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Update on antifungal therapy

Amy M. Grooters, DVM*, Joseph Taboada, DVM

Department of Veterinary Clinical Sciences, School of Veterinary Medicine,

Skip Bertman Drive, Louisiana State University, Baton Rouge, LA 70803

Over the past 20 years, fungi have emerged as pathogens of expanding

importance in human medicine, largely because of the increased prevalence
of immunocompromise associated with HIV infection and organ trans-
plantation. The need for more eﬀective and less toxic alternatives for the
treatment of systemic mycoses has prompted drug companies to search for
new ways to increase the safety and eﬃcacy of traditional antifungal agents
[1,2]. In addition, development of new agents that act selectively against
fungal targets has become a priority [3,4]. Historically, antifungal drug
development has lagged behind that of antibacterial agents, in part because
as eukaryotic organisms, fungi contain few drug targets that are not present
in mammalian cells. Traditional antifungal drugs (such as amphotericin B
and the azoles) target ergosterol, an essential component of the fungal cell
membrane. The selectivity of these drugs is based on their greater aﬃnity for
ergosterol in the fungal cell membrane than for cholesterol in the mam-
malian cell membrane. This selectivity limits but does not eliminate their
toxic eﬀects on mammalian cells. To circumvent this problem, the ideal
antifungal agent would be one that targets structures found in fungal
pathogens but not in other eukaryotic cells. The fungal cell wall is a structure
that is unique and essential to fungi. For this reason, compounds that
interfere with the synthesis of important fungal cell wall components (such
as b-glucan and chitin) have become a focus in the development of new
antifungal agents.

Amphotericin B lipid complex

Amphotericin B is a polyene antibiotic that causes cell death by binding

to ergosterol in the fungal cell membrane and disrupting membrane

Vet Clin Small Anim

33 (2003) 749–758

* Corresponding author.
E-mail address:

agrooters@vetmed.lsu.edu (A.M. Grooters).

0195-5616/03/$ - see front matter

doi:10.1016/S0195-5616(03)00038-X

stability. Because of its eﬃcacy against a broad spectrum of yeast and
ﬁlamentous fungal pathogens, amphotericin B traditionally has been the
treatment of choice for invasive mycoses in people and small animals. Its
usefulness has been restricted by nephrotoxicity, however, which limits the
cumulative dose that can be administered and prevents its administration to
patients with underlying renal dysfunction [5]. Recently, the use of novel
delivery systems has been eﬀective in reducing nephrotoxicity and improving
site-speciﬁc delivery of amphotericin B [6]. These new formulations allow
higher doses to be administered, in part because they increase the drug’s
uptake by tissues of the reticuloendothelial system, preventing its ac-
cumulation in the kidneys [7–10]. There are three new formulations of
amphotericin B available for clinical use in people. Amphotericin B lipid
complex (ABLC; Abelcet, The Liposome Company) consists of a mixture of
amphotericin B with two phospholipids, dimyristoyl phosphatidylcholine
(DMPC) and dimyristoyl phophatidylglycerol (DMPG), in a 7:3 molar ratio
[7]. When mixed together, the amphotericin B, DMPC, and DMPG form
ribbon-like lipid complexes with a bilayered membrane. Amphotericin B
colloidal dispersion (ABCD; Amphotec, Sequus Pharmaceuticals) is com-
posed of ﬂat disk-like structures of cholesterol sulfate in stable com-
plexes with amphotericin B [8]. Liposome-encapsulated amphotericin B
(AmBisome, Nexstar) consists of small unilamellar vesicles and is the only
true liposomal formulation [7,11]. Of these formulations, amphotericin B
lipid complex (ABLC) has been evaluated the most extensively in small
animals and is the least expensive.

The improved therapeutic index of ABLC in comparison to amphotericin

B deoxycholate has been demonstrated in preclinical and clinical studies. In
normal dogs receiving multiple doses, ABLC was determined to be 8 to 10
times less nephrotoxic than conventional amphotericin B [12]. This de-
creased nephrotoxicity can be attributed to several factors:

1. Lipid binding results in abrogation of amphotericin-induced direct

tubular toxicity.

2. Lipid binding reduces the amount of free amphotericin in solution [7].
3. Lipid complexes provide the opportunity for selective transfer of am-

photericin from its lipid carrier to ergosterol in the fungal cell membrane
(in contrast, the aﬃnity of amphotericin for cholesterol in the mam-
malian cell membrane is lower than its aﬃnity for the lipid carrier) [8].

4. Binding of lipid-complexed amphotericin to high-density lipoproteins

results in decreased uptake by renal cells [13].

The increased eﬃcacy of ABLC is caused largely by the rapid uptake of

lipid complexes by the reticuloendothelial (RE) system. As a result, the drug
is able to target sites of inﬂammation [14] and organs of the RE system, such
as the liver, spleen, and lungs [9,15]. Preferential uptake by organs of the RE
system allows higher doses of amphotericin to be administered with mini-
mal increases in the degree of renal uptake. When increasing doses of

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lipid-complexed amphotericin B were administered to mice, drug concen-
trations in the liver, spleen, and lung rose dramatically, whereas there was
little change in renal concentrations and no change in plasma levels [12,16].
Once at the target site, lipases from fungal or inﬂammatory cells may release
the amphotericin from its lipid complex, allowing it to bind to and disrupt the
fungal cell membrane.

Clinical trials in people have documented the eﬃcacy and improved

therapeutic index of lipid-complexed amphotericin B for the treatment of
many common fungal pathogens, including Candida, Aspergillus, Crypto-
coccus, Mucor, Histoplasma, Blastomyces

, and Coccidioides immitis [17–20].

In small animal patients, ABLC has been used successfully for the treatment
of blastomycosis, coccidioidomycosis, histoplasmosis, cryptococcal menin-
gitis, protothecosis, and pythiosis [21,22]. Speciﬁc indications for ABLC
administration include: initial therapy for patients with cryptococcal men-
ingitis (in which case it may be combined with ﬂucytosine), treatment of
animals with rapidly progressive or severe systemic mycosis (in which
itraconazole is unlikely to act quickly enough), and treatment of animals in
which persistent vomiting precludes the administration of oral medications.
In dogs, the authors administer 2 to 3 mg/kg of ABLC intravenously (IV)
three times weekly for a total of 9 to 12 treatments (cumulative dose of 24 to
27 mg/kg). In cats, a lower dose of 1 mg/kg three times weekly for a total of
12 treatments (cumulative dose of 12 mg/kg) is used. Amphotericin B lipid
complex is diluted in 5% dextrose to a concentration of 1 mg/mL and
infused over 1 to 2 hours. As with amphotericin B deoxycholate, serum cre-
atinine, serum urea nitrogen (BUN), and potassium should be checked
before each administration. Infusion-related adverse eﬀects such as pyrexia
and nausea can be diminished by pretreating patients with an antihistamine
such as diphenhydramine or with aspirin.

In addition to its fungicidal action, there is convincing evidence that

amphotericin B has signiﬁcant immunomodulating eﬀects that may play an
important role in its antifungal activity. The results of in vitro and in vivo
investigations suggest that amphotericin is a powerful macrophage ac-
tivator, potentiating their phagocytic, tumoricidal, and microbicidal actions
[23–25]. One important mechanism for this potentiation appears to be
enhancement of macrophage killing activity by way of nitric oxide-
dependent pathways mediated by amphotericin-induced production of
tumor necrosis factor alpha (TNF–a) and interleukin-1 (IL-1). In addition,
amphotericin has been shown to augment the macrophage oxidative burst
induced by TNF–a.

Azole antifungals

The azoles are a rapidly expanding class of antifungal agents that act by

inhibiting ergosterol biosynthesis at the C-14 demethylation stage, which

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interferes with fungal membrane function by causing depletion of ergosterol
and accumulation of lanosterol and other 14-methylated sterols [26]. A
major advantage of azole therapy has been the ability to treat endemic
mycoses such as histoplasmosis and blastomycosis on an outpatient basis
with oral medication, providing an alternative to IV amphotericin B.

Itraconazole

Itraconazole (Sporanox, Janssen, Ortho Biotech), a triazole released in

the United States in 1992, has become the treatment of choice in people and
small animals for most endemic or opportunistic mycoses that are not
immediately life-threatening [27,28]. In small animals, it has been used to
treat a variety of systemic mycoses at a dose of 5 to 10 mg/kg orally once
daily or divided twice daily (BID) [29–31]. Because itraconazole is a weak
base that requires an acid environment for maximal oral absorption, bio-
availability is increased when it is taken with food or cola beverages.
Concentrations of itraconazole are typically low in urine and cerebrospinal
ﬂuid (CSF), and it does not cross the blood–brain, blood–prostate, or
blood–ocular barriers well. Despite this fact, fungal infections involving
the central nervous system (CNS), prostate, or eye often respond well to
itraconazole therapy, perhaps because its lipophilic nature allows even small
amounts of the drug that move across inﬂamed barriers to accumulate in
these lipid-laden tissues.

Adverse eﬀects associated with itraconazole administration are typically

dose-dependent and include gastrointestinal (GI) toxicity, hepatic toxicity,
and cutaneous vasculitis. Recently, itraconazole has been solubilized in
cyclodextrins in an eﬀort to increase its absorption after oral dosing and to
allow development of a parenteral formulation. Pharmacokinetic studies in
people and cats have demonstrated increased absorption of itraconazole
after administration of the oral solution in comparison to capsules [32].
Sporanox Oral Solution (10 mg/mL) and Sporanox Injectable (250 mg
ampules diluted in 50 mL 0.9% NaCl for IV administration) are available.

Fluconazole

Fluconazole (Diﬂucan, Pﬁzer), a triazole released in 1990, has become

the drug of choice for treatment of mucocutaneous candidiasis in patients
with AIDS or neutropenia and for maintenance therapy of cryptococcal
meningitis in AIDS patients [33,34]. In small animals, ﬂuconazole is used
most often in patients with CNS or ocular manifestations of systemic my-
coses, and it is indicated speciﬁcally for the treatment of cryptococcosis [35].
In addition, it is used often to treat systemic mycoses in cats when
itraconazole is contraindicated or has been ineﬀective. Fluconazole is
available in oral and parenteral formulations. In the cat, it typically is used
at a dose of 50 mg per cat per day for nasal or dermal cryptococcosis, or

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100 mg per cat per day (usually divided BID) for systemic mycoses. In dogs,
ﬂuconazole typically is used at 2.5 to 10 mg/kg per day, most often for
cryptococcal meningitis. Fluconazole penetrates the blood–brain, blood–
prostate, and blood–ocular barriers well, and high concentrations are found
in CSF, urine, and ocular ﬂuids [36]. Because most administered ﬂuconazole
is excreted unchanged in the urine, its dose should be reduced in animals
with decreased glomerular ﬁltration rate (GFR).

New triazoles

The emergence of azole-resistant fungal pathogens (C krusei, C glabrata,

some strains of C albicans, Fusarium, dematiaceous molds such as Alternaria
and Exophiala, and some Aspergillus species) in people has prompted a search
for new triazoles with greater potency and a broader spectrum of activity
[37]. Three of these drugs, voriconazole, posaconazole, and ravuconazole,
are the ﬁrst of the newly developed triazoles to undergo clinical evaluation.

Voriconazole (Vfend, Pﬁzer) is a ﬂuconazole derivative that has greater

potency and a broader spectrum of activity than ﬂuconazole [38,39]. It
has potent in vitro and in vivo activity against common endemic and
opportunistic fungal pathogens (including molds other than the zygomy-
cetes), and has demonstrated eﬃcacy for invasive aspergillosis and oro-
pharyngeal candidiasis in phase III clinical trials [40]. Voriconazole has
excellent oral bioavailability, allowing it to be administered either orally or
IV. Its indications in people are for the treatment of invasive aspergillosis
and other infections caused by molds (such as Fusarium, Pseudallescheria
boydii

, and dematiaceous fungi) in immunocompromised individuals [41].

Posaconazole (Noxaﬁl, Schering-Plough) is an itraconazole analog that

has more potent in vitro activity than itraconazole or amphotericin B
against Aspergillus species, and it has the potential for good eﬃcacy for the
treatment of infections caused by less common molds, including Fusarium,
Pseudallescheria

, and Acremonium, and zygomycetes in the order Mucorales

[1,42,43]. It has demonstrated good eﬃcacy in animal models of candidiasis,
pulmonary and disseminated aspergillosis, histoplasmosis, cryptococcal
meningitis, coccidioidomycosis, zygomycosis, leishmaniasis, and Chagas dis-
ease [1]. In early open-label clinical studies, posaconazole administration
resulted in complete or partial responses in 50% to 80% of people with
systemic mycoses that were refractory to standard therapy [44]. Posacona-
zole is in phase III clinical trials.

Ravuconazole (formerly BMS 207147, Bristol-Meyers Squibb) is a ﬂuco-

nazole derivative that has good oral bioavailability and a very long half-life
(8.8 hours in dogs) [45]. It has demonstrated good in vitro activity against
yeasts, Aspergillus fumigatus, dermatophytes and dematiaceous fungi but
limited activity against Sporothrix, Pseudallescheria, Fusarium, and the
zygomycetes [1]. Ravuconazole is in phase II clinical trials for oropharyngeal
candidiasis.

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Terbinaﬁne

Terbinaﬁne (Lamisil, Sandoz) is an allylamine antifungal that blocks

ergosterol biosynthesis by inhibiting squalene epoxidase [46]. A keratino-
philic compound that is distributed well in skin, it has been used primarily
for the treatment of dermatophytosis and onychomycosis in human and
veterinary patients [47]. Descriptions of its use in the veterinary literature
are limited, however. Mancianti et al evaluated its use in 12 cats with der-
matophytosis caused by M canis and found that terbinaﬁne administered
orally at 30 mg/kg once daily for 2 weeks resulted in a clinical cure 1 to 3
months post-treatment in 10 of the 12 cats [48]. The eﬃcacy of terbinaﬁne
for invasive or systemic fungal infections has not been evaluated well. A
recent case report described successful treatment of a periorbital infection
caused by Pythium insidiosum in a 2-year-old child with a combination of
terbinaﬁne and itraconazole [49], however. In addition, the authors’ clinical
observations support the use of this drug combination for the treatment
of canine and feline pythiosis when complete surgical resection of infected
tissues is not possible. Sporotrichosis and phaeohyphomycosis are other
cutaneous mycoses for which terbinaﬁne may have eﬃcacy [50]. Potential
adverse eﬀects include GI toxicity and hepatitis [51].

b–glucan synthase inhibitors

Echinocandins and pneumocandins represent a new class of antifungal

agents (the lipopeptides) that hold perhaps the greatest promise for
changing the way that systemic mycoses will be treated in the next decade
[52]. These fungicidal compounds act by inhibiting b–glucan synthase,
blocking the synthesis of 1,3-b–D-glucan, a structural fungal cell wall
component that is not present in mammalian cells [53,54]. The most well-
studied of these agents, caspofungin (Cancidas, Merck), is a potent broad-
spectrum parenteral formulation that has potent activity against Aspergillus
species and Candida species [55,56]. In addition, it is highly eﬀective for the
treatment of Pneumocystis carinii pneumonia because of its ability to pre-
vent development of the glucan-rich cyst form. The primary limitation of
this class of antifungals is its ineﬀectiveness against Cryptococcus neofor-
mans

, which contains very little glucan synthase. Although caspofungin was

approved for the treatment of refractory invasive aspergillosis, it is also
highly eﬀective for oropharyngeal and esophageal candidiasis, and likely
also will be used for the treatment of azole-resistant candidiasis, suspected
candidemia, and febrile neutropenia [57]. The use of caspofungin in vet-
erinary patients has not been evaluated, and its extremely high cost certainly
will limit its otherwise tremendous potential. Another drug in this class that
has completed phase III clinical trials recently is micafungin (Fujisawa).
Anidulafungin (Versicor) is a third echinocandin that is undergoing clinical
trials.

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Chitin synthase inhibitors

Nikkomycins are competitive inhibitors of chitin synthase that have been

evaluated most extensively for their activity against Coccidioides immitis,
a fungal pathogen with high chitin content. One member of this group,
Nikkomycin Z, has been highly eﬀective for the treatment of coccidiomycosis
in animal models. Unfortunately, the spectrum of activity of nikkomycins for
other systemic mycoses is limited, and it is no longer being developed [54].

Lufenuron (Program, Ciba-Geigy) is a chitin synthase inhibitor of the

benzoylphenyl urea class. These compounds have a nonspeciﬁc inhibitory
eﬀect on chitin synthesis that is thought to be related to serine protease
inhibition. In one retrospective study conducted by Ben-Zion and Arzi,
single-dose oral administration of lufenuron signiﬁcantly decreased the time
to resolution of clinical signs associated with canine and feline dermato-
phytosis [58]. Results obtained in clinical practice have been mixed, how-
ever. Lufenuron has not been evaluated well for the treatment of systemic
mycoses in veterinary patients, but the limited spectrum of the nikkomycins
suggests that the usefulness of lufenuron will be limited. It has been
evaluated for the treatment of pulmonary coccidioidomycosis in 17 dogs
treated with 5 to 10 mg/kg once daily for 16 weeks [59]. Clinical and radio-
graphic improvement was noted in most of these dogs; however, because
spontaneous remission may occur in infected dogs without treatment, it is
unclear whether the clinical improvement was attributable to lufenuron
administration. Dogs with disseminated coccidioidomycosis have not
responded well to lufenuron therapy.

Fungal protein synthesis inhibitors

Sordarins are a class of antifungal agents that inhibit fungal protein

synthesis by interacting with EF-2, a translation elongation factor that is
necessary for elongation of the polypeptide chain during protein synthesis
[60,61]. Initial studies have indicated good activity against Candida albicans,
and they also may be useful for the treatment of Pneumocystis pneumonia
[62]. They have not performed well in animal models of disseminated as-
pergillosis, however, [63].

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Update on feline calicivirus: new trends

Kate F. Hurley, DVM, MPVM

Jane E. Sykes, BVSc(Hons), PhD

Center for Companion Animal Health, 270 Veterinary Medicine 11,

University of California, Davis, CA 95616, USA

Department of Medicine and Epidemiology, University of California, VM:

Medicine and Epidemiology, 2108 Tupper Hall, Davis, CA 95616, USA

Feline calicivirus (FCV) is a common cause of feline upper respiratory

tract disease (URTD), accounting for 20% to 53% of cases. Typically,
infection is manifested by fever, conjunctivitis, rhinitis, oral ulcerations, or
chronic stomatitis, although occasionally, skin ulcerations, lameness, and
pneumonia may occur.

The causative agent is a nonenveloped, single-stranded RNA virus with

a spherical capsid studded with cup-shaped depressions. Although consider-
able antigenic diversity occurs among FCV isolates, the degree of cross-
reactivity is suﬃcient for them to be classiﬁed as a single serotype. Similarly,
results of nucleotide sequence analysis have suggested that isolates world-
wide should be considered as a single, diverse group [1]. Like other RNA
viruses, the genome of FCV continually undergoes rapid mutation, with
minimal rates of repair, increasing the diversity of strains over time [2].

Cats that recover from URTD may develop a persistent oropharyngeal

infection, which has been termed the carrier state for FCV. Virus is shed
continuously from this site, although the magnitude of shedding varies with
time and between individual cats [3]. These cats may serve as a source of
infection for other susceptible cats. In many cats, shedding terminates weeks
to months after infection, but in a few cats, shedding is life-long. A single cat
may be infected with multiple variants of FCV at the same time, each derived
from the original infecting strain as a result of genetic mutation, drift, and
selection pressures [4]. Because of the chronic carrier state, the prevalence of
FCV infection in healthy cats is high, ranging from 8% of household cats to
24% of show cats [5].

Vet Clin Small Anim

33 (2003) 759–772

* Corresponding author.
E-mail address:

jesykes@ucdavis.edu (J.E. Sykes).

0195-5616/03/$ - see front matter

doi:10.1016/S0195-5616(03)00025-1

Upper respiratory tract disease caused by FCV is especially a problem in

cats residing in multiple cat households and breeding and boarding catteries,
where it may be associated with a high morbidity and mortality. Although
the introduction of vaccines targeting FCV in the middle of the 1970s may
have reduced the severity of clinical signs, the vaccines do not prevent
infection or persistent shedding of FCV, and URTD may still occur. Fur-
thermore, there has been some concern that vaccine pressure may have led to
selection of FCV strains that have poor cross-reactivity with the vaccine
strain F9, and incorporation of additional strains into vaccines has been
proposed [6].

Over the last 5 years, several highly virulent strains of FCV have been

isolated from outbreaks of a systemic, hemorrhagic-like fever in North
American cats. This condition was described initially in northern California
by Pedersen et al, and subsequently by other investigators across the United
States [7]. The disease has been associated with mortalities of up to 50% in
some outbreaks. Furthermore, vaccination with FCV vaccines has not been
protective. In several respects, this condition has resembled rabbit
hemorrhagic disease, which emerged in the middle of the 1980s, causing
widespread mortality in rabbits [8,9]. The purpose of this article is to discuss
the clinical signs, epidemiology, diagnosis, control, and treatment of FCV
infections, with particular emphasis on these recently reported hemorrhagic-
like fever FCV infections.

Epidemiology

Infection with FCV may be acquired by direct contact with an acutely

infected cat, organisms persisting within the environment, or a carrier cat.
The chance of contact with an acutely infected cat is increased when many
cats are housed together, such as in cattery situations and multiple cat
households. FCV is shed primarily in oral, nasal, and ocular secretions, and
also can be found in blood, urine, and feces of infected cats. Transmission
over distances of about 4 ft may occur by way of droplets generated by
sneezing cats. Airborne transmission is considered unlikely, possibly because
cats lack the tidal volume to generate an eﬀective aerosol [10].

A febrile hemorrhagic disease caused by a FCV was described ﬁrst in

northern California in 1998 [7]. Thirty-three percent to 50% of infected cats
died, and this strain was highly contagious, spreading by way of con-
taminated fomites despite implementation of aggressive disinfection mea-
sures. The condition was characterized by anorexia, ulcerative facial
dermatitis, and diﬀuse cutaneous edema. A novel FCV strain, FCV-ari,
was isolated from aﬀected cats. Laboratory cats experimentally infected with
FCV-ari developed a hemorrhagic disease syndrome identical to that seen in
naturally infected cats. Since FCV-ari was described, at least ﬁve additional
outbreaks have been recognized in Pennsylvania, Massachusetts, Tennessee,

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Nevada and Southern California (J.R. Richards, personal communication;
J. Dinnage, personal communication; N.C. Pedersen, personal communica-
tion; M. Kennedy, personal communication; S. Anderson, personal commu-
nication; K.F.H., unpublished data). FCV-kaos, a genetically distinct strain
from FCV-ari (and the vaccine [F9] strain and several other FCV ﬁeld
strains), was isolated from cats in the southern California outbreak (K.F.H.,
unpublished data, 2002), and the strain isolated in the Massachusetts outbreak
was genetically distinct from FCV-kaos and FCV-ari. This may suggest that
the mutation(s) associated with hemorrhagic disease are diﬀerent in each case,
or that these mutations occurred in a part of the FCV genome that was not
sequenced by all investigators. An FCV strain from the Tennessee outbreak
also has been isolated and characterized.

The six suspected or conﬁrmed outbreaks of hemorrhagic calicivirus

disease share several signiﬁcant features:

In every outbreak where a suspect index case was identiﬁed (ﬁve of six

outbreaks), a hospitalized shelter cat appeared to be the source of
infection.

Otherwise healthy, adult, vaccinated cats were affected prominently,

whereas kittens tended to show less severe signs.

Spread occurred very readily by way of fomites to cats belonging to

hospital employees and clients.

Spread of disease was limited to the affected clinic or shelter, with no

spread within the community reported.

The outbreak resolved within approximately 2 months.

Although the index case was most commonly a shelter cat, only the

Pennsylvania outbreak occurred in a shelter. Three outbreaks involved only
a single clinic. One spread to a veterinary research facility in the course of the
outbreak investigation, and the largest reported outbreak aﬀected 54 cats in
three veterinary practices and a rescue group in the West Los Angeles area
from June through August of 2002 (Hurley et al, unpublished data, 2002).
The wider spread of this southern California outbreak was attributed to the
travel of clients and staﬀ between multiple practices located within a 1-mile
radius. Two outbreaks occurred in the spring, two in the summer, and two in
the fall. No outbreak has been reported in winter, a time when the kitten popu-
lation is at its lowest, reﬂecting the possible role of mildly or subclinically
aﬀected kittens in the propagation of hemorrhagic calicivirus infection.

Transmission of hemorrhagic FCV strains occurs very readily, and

disease has been observed to spread by fomite transmission by way of
veterinary hospital staﬀ and pet owners, by movement of clinically and
subclinically infected cats between clinics and private homes, and by indirect
contact of outpatients with subclinically infected inpatients. Even brief
indirect contact has been adequate to spread the disease, such as restraint of
a cat for physical examination by a person wearing contaminated clothes,
even if hands have been washed between handling cats.

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An epidemiological analysis of the outbreak that occurred in Southern

California showed that adult cats (older than 1 year) were at signiﬁcantly
higher risk than kittens (younger than 6 months) for severe disease or death
(odds ratio 9.56, conﬁdence interval [CI] 2.82 to 32.39, P < 0.001). No sex or
breed predilection was noted. Many cases occurred in otherwise apparently
healthy, fully vaccinated indoor cats; at least 26 of the 54 aﬀected cats had
a history of vaccination. Virtually all exposed cats in an aﬀected clinic may
develop infection, with an attack rate of 94% documented in one outbreak
(Hurley et al, unpublished data, 2002). Vaccination does not appear to be
protective against any hemorrhagic FCV strain, although it remains possible
that only outbreaks severe enough to warrant reporting result from infection
with vaccine-resistant strains. Further studies are required to determine
whether vaccine resistance results from a change in antigenic structure
linked to mutation(s) causing enhanced virulence and hemorrhagic-like
disease. In one small study, four kittens orally immunized with the current
vaccine strain (F9) experienced less severe clinical signs when experimentally
infected with FCV-ari [7]. Neither intranasal nor parenteral vaccination
provided signiﬁcant protection against infection or severe disease caused by
natural exposure to FCV-kaos, however (K.H., unpublished data, 2002).

Many cats infected with FCV remain chronic carriers [3,11], and cats

infected with hemorrhagic strains have been demonstrated to shed virus for
at least 16 weeks after infection (Hurley et al, unpublished data, 2002). Thus
it seems possible that a carrier state for hemorrhagic FCV strains could
occur, and chronically infected cats could pose a threat long after recovery.
Despite these observations, there has been no report of transmission of the
hemorrhagic disease from a fully recovered cat, and in every outbreak,
surviving cats have returned to homes in the community. No outbreak has
persisted for more than 2 to 3 months, nor have sporadic cases ever been
reported in a community following resolution of an outbreak.

Ultimately, the reason for the spontaneous resolution of all reported

hemorrhagic fever-like outbreaks has not been determined. Widespread
susceptibility to infection has been observed regardless of age, health, or
vaccination status, so it seems unlikely that infection dissipates because of
a lack of susceptible hosts. If the virulence and transmissibility of virus shed
by persistently infected cats is retained, sporadic cases, or even additional
outbreaks, should arise from the potential reservoir of chronically shedding
cats. Failure of veterinarians to report these cases seemed unlikely in the
southern California outbreak, as there was widespread awareness of the
condition in the veterinary community in the area (Hurley et al, unpublished
data, 2002). It is possible that the mutation(s) resulting in enhanced virulence
are lost during passage in these cats. Variant strains arise in persistently
infected cats, possibly as a result of immune pressure [4] and genetic
variation, was noted between successive isolates of FCV-kaos obtained over
the course of the southern California outbreak. It is also possible that
implementation of eﬀective control measures played a role in outbreak

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K.F. Hurley, J.E. Sykes / Vet Clin Small Anim 33 (2003) 759–772

resolution. Once these measures were instituted, the likelihood of an
infectious cat contacting a susceptible cat were reduced greatly, and perhaps
thereafter, no persistently shedding survivor had suﬃciently close or
prolonged contact to transmit infection to a susceptible cat.

Clinical signs

Acute upper respiratory tract disease (URTD) caused by FCV occurs

after an incubation period of 2 to 10 days. The nature and severity of disease
may depend on the infecting strain, although examination of large numbers
of FCV strains failed to show a clear link between disease manifestation and
antigenic and heterogeneity of the FCV capsid protein [12,13]. Typical signs
of URTD include serous or mucopurulent nasal and ocular discharge,
sneezing, conjunctival hyperemia, blepharospasm, and chemosis. Depres-
sion, anorexia, hypersalivation and pyrexia also may be seen, and
occasionally, pneumonia may develop with coughing and dyspnea. Ulcera-
tive glossitis develops in some aﬀected cats. A small proportion of FCV
carriers develop chronic lymphoplasmacytic or ulceroproliferative stomati-
tis, which may be refractory to treatment.

Cats exposed to FCV strains causing the systemic hemorrhagic-like

febrile disease while hospitalized in veterinary clinics have developed signs 1
to 5 days after exposure to aﬀected cats. Peracute disease has developed in
less than 24 hours. The incubation period has been longer (up to 12 days) for
cats exposed in the home environment (Hurley et al, unpublished data,
2002). Aﬀected cats frequently show signs typical of caliciviral URTD,
including anorexia, oral ulceration, and nasal or ocular discharge, although
these signs are often severe. Fever is present in most cases and may be
profound, often exceeding 105

F (40.6

C). Distinctive clinical signs of the

systemic hemorrhagic-like febrile disease include cutaneous edema, alopecia,
crusting, and ulceration (Hurley et al, unpublished data, 2002) [7]. Pustules
also have been noted in some cases. Edema is most commonly noted on the
head and limbs, and crusting and ulceration are most prominent on the
nose, lips, pinnae, periocularly, and on the distal limbs. Some cats may be
reluctant to walk. Cats in the early stages of developing facial edema have
been described as having a ‘‘Roman nose’’ appearance. Severe respiratory
distress, sometimes caused by pulmonary edema or pleural eﬀusion, devel-
ops in some cats. Icterus has been noted in many aﬀected cats. Clinically
apparent jaundice and dyspnea have been associated with a poor prognosis.
Involvement of the gastrointestinal [GI] tract, including the liver and
pancreas, may be associated with signs of vomiting or diarrhea. Late in the
course of disease, signs of coagulopathy may be seen, including petechiae
and ecchymoses; rarely, epistaxis and hematochezia have been noted.
Coagulopathy may result from disseminated intravascular coagulation.
Peracutely aﬀected cats may die as a result of cardiovascular arrest with

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little preceding signs apart from fever. Table 1 lists the frequency of clinical
signs reported from an outbreak in southern California. A characteristically
aﬀected kitten is shown in Fig. 1.

Clinicopathologic abnormalities resulting from infection with nonhemor-

rhagic FCV strains are generally nonspeciﬁc, such as neutrophilia and
hyperglobulinemia. The latter may be a prominent ﬁnding in cats with
chronic stomatitis. In contrast, infection with hemorrhagic FCV strains may
result in severe clinicopathologic derangements reﬂecting damage to
multiple organ systems (Hurley et al, unpublished data, 2002). Frequently
reported serum chemistry proﬁle abnormalities include hyperbilirubinemia,
which may be mild or severe; hypoalbuminemia; hyperglycemia; elevated
creatine phosphokinase (CPK) and aspartate aminotransferase (AST); and
mildly elevated alanine aminotransferase (ALT). Results of the serum
chemistry proﬁles collected from 10 cats in the southern California outbreak
are reported in Table 2. There have been no consistent ﬁndings on complete
blood count (CBC); in some cases, the CBC has been normal. Mild-to-
marked lymphopenia is relatively common, and in some cats, mild-to-
moderate neutrophilia or thrombocytopenia has been noted. Hematocrit is
normal to mildly decreased, although more severe anemia may develop in
cats with coagulopathies. Abnormal CBC ﬁndings in 10 cats from the
southern California outbreak are reported in Table 3. Blood work results
were not available for the remaining cases.

Table 1
Clinical signs reported

for cats infected with FCV-kaos, a hemorrhagic fever-like FCV causing

an outbreak in southern California, summer, 2002

Clinical sign

Number of cases
(percentage of total)

Number of fatal cases
(percentage of total)

Fever

44 (81%)

21 (95%)

Limb or facial edema

28 (52%)

16 (73%)

Oral ulcers

25 (46%)

7 (32%)

Nasal discharge

16 (30%)

6 (27%)

Dyspnea

9 (17%)

7 (32%)

Ulcerations

/alopecia

9 (17%)

4 (18%)

Jaundice

6 (11%)

6 (40%)

Conjunctivitis

/ocular discharge

6 (11%)

4 (18%)

Diarrhea

4 (7%)

3 (14%)

Vomiting

4 (7%)

2 (9%)

Limping

3 (6%)

Total

Abbreviation:

FCV, feline calicivirus.

From review of medical records or interviews with owners if no written record was

available.

Median, 105.1

F, range, 103.0

–106.5

F (39.4

–42.4

C).

24 reports of limb edema, 13 reports of facial edema.

Data from

Glenn M, Radford AD, Turner PC, et al. Nucleotide sequence of feline calicivirus

(FCV) and phylogenetic analysis of FCVs. Vet Microbiol 1999;67:175–93.

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Although overall mortality from the systemic hemorrhagic-like febrile

disease is approximately 50% and may be even higher in adult cats,
subclinical infection or mild disease indistinguishable from URTD caused
by nonhemorrhagic FCV strains was recognized in 20% of cats in the
southern California outbreak, based on positive virus isolation and genetic
sequencing results (Hurley et al, unpublished data, 2002). Mild disease is
more common in kittens, but this also may be observed in adult cats.
Subclinically infected animals can play an important role in perpetuating an

Fig. 1. A cat suﬀering from hemorrhagic feline calicivirus infection (FCV-kaos strain) showing
characteristic signs of facial edema and sore, crusting and alopecia of the face and pinnae.

Table 2
Blood chemistry abnormalities reported in cats infected with FCV-kaos during an outbreak of
hemorrhagic fever-like FCV in southern California, summer, 2002

(abnormal values in bold)

Case #

Total bilirubin
(mg

/dL)

Albumin
(g

/dL)

AST
(IU

/L)

ALT
(IU

/L)

CK
(IU

/L)

0.1

3.8

1143

1.5

2.6

102

256

3.9

1.4

223

10,930

0.1

2.9

156

0.3

2.1

2677

1.9

1.1

169

116

1697

0.6

2.1

103

639

1.83

1.2

0.97

3.01

0.4

2.3

460

Abbreviations:

ALT, alanine aminotransferase; AST, aminotransferase; CK, creatine kinase;

FCV, feline calicivirus.

Blood work was performed at the discretion of attending veterinarians and is not

necessarily representative of all aﬀected cats.

Blood work was performed at various laboratories with diﬀerent reference ranges.

/a, not available.

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K.F. Hurley, J.E. Sykes / Vet Clin Small Anim 33 (2003) 759–772

epidemic, and asymptomatic cats have been documented to transmit fatal
disease.

Necropsy ﬁndings are variable. In addition to the cutaneous edema and

ulceration apparent grossly, the most common pathology ﬁnding has been
individual hepatocellular necrosis, although this was not present in all cats
examined. Other frequently reported ﬁndings include acute interstitial pneu-
monia and free pleural and abdominal ﬂuid. Pericardial ﬂuid was reported in
one case. Intestinal crypt lesions were reported in four cats experimentally
infected with FCV-ari, and pancreatitis was reported in several cats infected
with this strain and in one cat infected with FCV-kaos (Hurley et al,
unpublished data) [7]. Gastrointestinal ulceration has been reported
occasionally. There has been no consistent cause of death identiﬁed.

Diagnosis

The clinical signs resulting from uncomplicated FCV infection can result

from infection by other infectious agents, especially feline herpesvirus 1, but
also bacterial pathogens such as Chlamydiophilia felis and Bordetella bron-
chiseptica

. Thus, clinical signs alone are not useful for establishing a deﬁnitive

diagnosis, although the presence of severe oral ulcerations or lameness may
be suggestive of FCV infection. Mixed infections may occur, and are common
in cattery situations, further complicating diagnosis. Cats presenting with
signs of epistaxis, unilateral nasal discharge, facial deformity, fundic
abnormalities, or marked submandibular lymphadenopathy should receive
a thorough work-up to rule out other causes of URTD such as malignant
neoplasia, nasopharyngeal polyps, coagulopathy, foreign bodies, dental

Table 3
Blood count abnormalities reported in cats infected with FCV-kaos during an outbreak of
hemorrhagic fever-like FCV in southern California, summer, 2002

(abnormal values in bold)

Case

HCT (%)

WBC
(cells

/lL)

Neutrophils
(cells

/lL)

Lymphocytes
(cells

/lL)

Platelets
(cells

/lL)

13,200

11,616

1188

adequate

9000

7020

1710

adequate

7800

7254

468

adequate

12,800

n/a

30.3

8600

8084

258

adequate

3000

2790

180

adequate

17,600

5100

12,500

268,000

12,800

11,700

1000

164,000

34.3

5100

4437

459

adequate

Abbreviations:

FCV, feline calicivirus; HCT, hematocrit; WBC, white blood cell count.

Blood work was performed at the discretion of attending veterinarians and is not

necessarily representative of all aﬀected cats.

Blood work was performed at various laboratories with diﬀerent reference ranges.

/a, not available.

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K.F. Hurley, J.E. Sykes / Vet Clin Small Anim 33 (2003) 759–772

disease, and cryptococcosis. Although clinical signs of the hemorrhagic
disease syndrome are more characteristic, conditions such as cryptococcosis
and immune-mediated vasculitides represent possible diﬀerential diagnoses.

Attempts to make a diagnosis in cases of URTD are encouraged in cattery

situations, as knowledge of the causative organism can assist with manage-
ment strategies. Diagnostic work-up also is recommended for individual
cats that do not respond to conventional symptomatic treatment, or that ex-
perience recurrent episodes of URTD. The diagnostic work-up for cats with
chronic nasal discharge should include routine blood work and urinal-
ysis; serologic testing for feline immunodeﬁciency virus (FIV), feline
leukemia virus (FeLV), and Cryptococcus neoformans; skull radiographs or
a nasal CT scan; and rhinoscopy and biopsy of the nasal mucosa, in addition
to microbiologic testing for viral upper respiratory tract pathogens and
antimicrobial-resistant secondary bacterial infections. Because of the
communicability and high mortality associated with virulent FCV infection,
microbiologic testing is essential for cats suspected to have the febrile
hemorrhagic syndrome, and suspect cats should be treated and handled
immediately as if they were infected with a virulent FCV strain.

The most reliable microbiologic assay for FCV is virus isolation from

nasal, conjunctival, or oropharyngeal swabs after inoculation of cell
monolayers grown in the laboratory. Oropharyngeal swabs are most likely
to yield a diagnosis, although if possible, collection of nasal and conjunc-
tival swabs in addition to oropharyngeal swabs will increase the chance of
obtaining a positive result. Swabs should be transported on ice in a viral
transport medium containing antibiotics to prevent bacterial overgrowth;
commercial swabs are available for this purpose. Fluorescent antibody has
been used on cytology and tissue samples to diagnose FCV infection, but
this is less sensitive than virus isolation, and the presence of nonspeciﬁc
ﬂuorescing debris may be associated with false-positive results.

The sensitivity of virus isolation using oropharyngeal swabs collected

from acute cases or tissue samples obtained at necropsy was close to 90% in
the southern California outbreak. Positive culture results also were obtained
on serum samples from acutely ill cats, although the sensitivity was lower
than culture from oropharyngeal swabs. Because of the chronic carrier state,
however, positive serum cultures may be a more speciﬁc indicator of acute
infection than positive oropharyngeal cultures. Sensitivity of viral culture on
oropharyngeal swabs decreases rapidly in recovering cats, and a single
negative swab taken more than 1 week after onset of clinical signs does not
rule out infection with FCV. Results of virus isolation were only inter-
mittently positive in recovered cats sampled successively over a 10-week
period, so at least two to three negative cultures should be obtained at
weekly intervals before concluding that virus shedding has terminated.

In general, serology is not recommended for routine diagnosis of FCV

infections. In many cases without development of clinical illness, acute and
convalescent phase samples are required to demonstrate recent infection

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K.F. Hurley, J.E. Sykes / Vet Clin Small Anim 33 (2003) 759–772

because of widespread exposure to FCV infections. Titers induced by
vaccination can confound diagnosis, and because of the large numbers of
FCV strains, titers may vary depending on the degree of homology between
the infecting virus and the FCV strain used in the assay. Nevertheless, in the
case of FCV strains causing hemorrhagic disease, serology using virus
neutralization has had apparent sensitivity and speciﬁcity and was useful for
investigation and control of the southern California outbreak (Hurley et al,
unpublished data, 2002). Twenty cats from this outbreak with a known his-
tory of exposure were seropositive (at least 1:16) and two cats that were
likely unexposed were seronegative (at less than 1:4). There was no cross-
neutralization between the vaccine (F9) strain and the FCV-kaos or FCV-ari
strains, so identiﬁcation of unexposed cats even in the face of recent vaccina-
tion may be possible. Whether this will be true for other hemorrhagic FCV
strains (and thus for other outbreaks) awaits further investigation.

Ampliﬁcation of FCV nucleic acid using the polymerase chain reaction

(PCR) represents another diagnostic assay that may be considered for
diagnosis. Compared with those for DNA viruses, such as feline herpesvirus
1, PCR assays for FCV are less reliable because of the diﬃculty in designing
assays that amplify nucleic acid from a variety of diﬀerent strains and the
susceptibility of viral RNA to rapid degradation by RNase enzymes in the
environment. False-positive results associated with contamination can occur
during sample collection or in the laboratory. PCR assays for FCV are
generally only available in research laboratories, and quality control may
vary from laboratory to laboratory and even between individual staﬀ mem-
bers working in the same laboratory. Therefore, the results of PCR testing
should be interpreted with caution. In the southern California outbreak of
systemic febrile hemorrhagic-like disease, all isolates recovered in culture
were ampliﬁed successfully using PCR (Hurley et al, unpublished data).

Regardless of the method used to detect FCV infection, the results of

diagnostic testing always should be interpreted in the light of clinical signs,
because asymptomatic cats may shed FCV. The same applies for FCV
strains associated with the hemorrhagic disease syndrome, because there
have been no consistent molecular diﬀerences identiﬁed between hemor-
rhagic strains and other strains of FCV, so it is not possible to identify an
isolate as a hemorrhagic disease strain using molecular typing methods.
Positive identiﬁcation of an outbreak requires that a molecularly distinct
strain of FCV be isolated and sequenced from at least two aﬀected cats in
association with the appearance of consistent clinical signs. Once an
outbreak has been identiﬁed in this manner, nucleic acid sequencing may be
useful for determining whether additional FCV isolates obtained in the
outbreak have the potential to cause hemorrhagic disease (because of their
relatedness to the original isolates), or whether they represent unrelated ﬁeld
FCV or vaccine strains. Because subclinically infected cats are capable of
transmitting the febrile hemorrhagic syndrome, all exposed cats should be
checked by viral culture or PCR before being introduced to naı¨ve cats.

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K.F. Hurley, J.E. Sykes / Vet Clin Small Anim 33 (2003) 759–772

Ideally, a full necropsy should be performed whenever the systemic

febrile hemorrhagic syndrome is suspected. At minimum, tissues to examine
include skin (particularly footpads, nasal planum, and any areas of obvious
ulceration), tongue, lung, liver, spleen, GI tract, pancreas, and lymph nodes.

Treatment

Treatment of disease resulting from FCV infection is symptomatic.

Broad-spectrum antimicrobials (eg, amoxicillin [22 mg

/kg by mouth every

12 hours] or doxycycline [10 mg

/kg by mouth every 24 hours]) may be

required to counteract secondary bacterial infection. Fluid and nutritional
support is essential in severe cases, and airway humidiﬁcation also should be
considered. Placement of esophagostomy or gastrostomy tubes should be
considered to allow enteral nutrition of cats with inappetence. Oxygen may
be required for cats with pneumonia.

Cats with the febrile hemorrhagic syndrome should be placed in isolation

and treated aggressively with colloid-containing ﬂuids such as hetastarch, in
addition to the treatments described previously. In the cases described,
a variety of diﬀerent treatments were tried, including a wide range of broad-
spectrum antibiotics, and oral interferon-alpha. Glucocorticoids at immu-
nosuppressive doses (1 mg

/kg by mouth every 12 hours) have been used

without increasing mortality, and there is weak evidence that they may have
beneﬁcial eﬀects. In the outbreak in southern California, ﬁve severely af-
fected cats were treated with dexamethasone (in addition to other sup-
portive treatments) and survived. One cat treated with prednisone died, but
the remaining cats that failed to survive did not receive glucocorticoid treat-
ment (K.H., unpublished data, 2002). One cat in the outbreak in northern
California lived despite glucocorticoid treatment, whereas the remainder
were not treated with steroids.

Control recommendations

The outbreaks of febrile hemorrhagic caliciviral disease described in the

United States have demonstrated the importance of control measures to limit
spread of feline upper respiratory tract viruses, because of the high mortality,
poor eﬃcacy of vaccines, and lack of speciﬁc treatments for the condition.
Although outbreaks can be dramatic and devastating within an aﬀected clinic
or shelter, the disease remains extremely uncommon. Quick recognition
and implementation of eﬀective control measures, including proper
disinfection, quarantine, and testing procedures, will reduce the impact of
this disease further.

Recognizing that the hemorrhagic disease may be spread even by mildly

symptomatic cats (especially kittens), veterinarians should exercise careful
infectious disease control whenever dealing with a cat with URTD. Feline
calicivirus is inactivated reliably by sodium hypochlorite (when stored and

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K.F. Hurley, J.E. Sykes / Vet Clin Small Anim 33 (2003) 759–772

used correctly in the absence of organic matter and with adequate contact
time) but not by other compounds commonly used for disinfection in
veterinary hospitals, such as chlorhexidine or quaternary ammonium
compounds [14–17]. Several clinics involved in the southern California
outbreak were using quaternary ammonium compounds for disinfection at
the time the outbreak occurred. Therefore, routine decontamination after
exposure to a cat with upper respiratory infection should employ cleaning with
a detergent solution followed by disinfection with 5% sodium hypochlorite
diluted with water to 1:32 (1/2 cup of 5% bleach per gallon of water). In the
absence of eﬀective disinfection, feline caliciviruses reportedly can persist in
a dried state at room temperature (20

C) for up to 28 days [14]. Furthermore,

the calicivirus causing rabbit hemorrhagic disease remained infective in a dried
state at room temperature for 105 days [18]. Therefore, eﬀective cleaning and
quarantine are crucial. Speciﬁc control measures recommended when the
febrile hemorrhagic disease is diagnosed or strongly suspected are as follows:

1. Strictly isolate suspect cases. Limit the number of staﬀ handling aﬀected

cats and require full protective clothing (including jumpsuits or
protective smocks, gloves, shoe protection, and caps) for all people
entering isolation (including clients if permitted to visit). Provide
separate supplies for treatment, examination, and cleaning, in isolation.
Separate air exchange is probably not necessary, but aﬀected cats should
be at least 5 ft away from healthy cats to prevent droplet spread. Suspect
cases may be treated as outpatients if medically appropriate, and there
are no unexposed cats at home or if at-home isolation is practical.

2. Strictly isolate all exposed cats (away from symptomatic cats) for at least

2 weeks after exposure. No exposed cat should be allowed to remain in
the hospital outside of isolation, including blood donor cats and any
clinic residents, as these cats may serve to propagate the infection. These
cats should be monitored for fever, oral ulceration, anorexia, or other
signs of disease.

3. Exposed or infected cats should be monitored by viral culture of

oropharyngeal swabs before being released into environments containing
naı¨ve cats. Ideally, at least two consecutive negative viral cultures at least
1 week apart should be obtained before release. Fully recovered cats that
remain persistently culture positive have been sent home to naı¨ve cats
without apparently transmitting the disease. It is unknown, however,
whether or at what point such cats completely cease being a risk to others.

4. Thoroughly clean the entire premises with 5% bleach solution diluted

with water at 1:32, including reception areas and any other area where
animals are permitted. Surfaces should be scrubbed thoroughly to
remove any caked on organic matter before disinfection. Special care
should be taken to clean any instruments that are handled commonly,
such as telephones, keyboards, and doorknobs. Instruments may be
disinfected using heat sterilization.

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K.F. Hurley, J.E. Sykes / Vet Clin Small Anim 33 (2003) 759–772

5. For contaminated areas such as carpeting or upholstered furniture that

cannot be cleaned with bleach, mechanically clean as thoroughly as
possible with vacuum followed by steam cleaning and close such areas to
cats for a minimum of 4 weeks. The same recommendation applies to
contaminated homes.

6. All staﬀ should change clothes and shoes and wash hands immediately

before leaving the hospital, especially if going to another clinic or to
a home containing cats.

7. Contact owners of all cats seen since the suspected index case was

hospitalized. These individuals should be advised to watch for leth-
argy, anorexia, fever, oral ulcerations, signs of URTD, or cutaneous
edema.

8. Contact all veterinary clinics, shelters, and rescue groups in the area,

especially those veterinary clinics that may see clients from the aﬀected
hospital.

9. Although there is no evidence that these virulent FCV strains pose any

zoonotic threat, notiﬁcation of public health authorities should be
considered, as rumors of a hemorrhagic fever may cause public concern.

10. Clinics in which disease continues to spread despite instituting precau-

tions should consider closing to cat admissions for 1 to 2 weeks. This
was necessary in two clinics involved in the outbreak that occurred in
southern California (Hurley et al, unpublished data, 2002).

11. Although vaccination with FCV vaccines has not appeared to be pro-

tective in the outbreaks reported, it is possible that future hemorrhagic
FCV strains may arise that are not vaccine-resistant. If vaccinated cats
are aﬀected less severely in any outbreak, vaccination of naı¨ve cats with a
modiﬁed live vaccine at least 1 to 2 weeks before exposure may be
helpful. Vaccination with a modiﬁed live vaccine may interfere with
diagnosis, however, as vaccination may cause mild signs of URTD and
positive results on viral culture.

Summary

In addition to being important upper respiratory tract pathogens of cats,

FCVs are increasingly reported as a cause of a highly contagious febrile
hemorrhagic syndrome. Strains causing this syndrome are genetically
diﬀerent from the vaccine strain and other nonhemorrhagic FCV isolates.
They apparently diﬀer from one outbreak to another. The syndrome is
characterized variably by fever; cutaneous edema and ulcerative dermatitis;
upper respiratory tract signs; anorexia; occasionally icterus, vomiting, and
diarrhea; and a mortality that approaches 50%. Adult cats tend to be more
severely aﬀected than kittens, and vaccination does not appear to have
a signiﬁcant protective eﬀect. Rapid recognition of the disease through
identiﬁcation of clinical signs and appropriate testing, followed by strict

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K.F. Hurley, J.E. Sykes / Vet Clin Small Anim 33 (2003) 759–772

institution of disinfection, isolation, and quarantine measures, are essential
to prevent widespread mortality resulting from the infection.

References

[1] Glenn M, Radford AD, Turner PC, et al. Nucleotide sequence of UK and Australian

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1999;67(3):175–93.

[2] Johnson RP. Antigenic change in feline calicivirus during persistent infection. Can J Vet

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[3] Wardley RC. Feline calicivirus carrier state. A study of the host

/virus relationship. Arch

Virol 1976;52(3):243–9.

[4] Radford AD, Turner PC, Bennett M, et al. Quasispecies evolution of a hypervariable

region of the feline calicivirus capsid gene in cell culture and in persistently infected cats.
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[5] Wardley RC, Gaskell RM, Povey RC. Feline respiratory viruses–their prevalence in

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[6] Dawson S, McArdle F, Bennett M, et al. Typing of feline calicivirus isolates from diﬀerent

clinical groups by virus neutralisation tests. Vet Rec 1993;133(1):13–7.

[7] Pedersen NC, Elliott JB, Glasgow A, et al. An isolated epizootic of hemorrhagic-like fever

in cats caused by a novel and highly virulent strain of feline calicivirus. Vet Microbiol
2000;73(4):281–300.

[8] Mutze G, Cooke B, Alexander P. The initial impact of rabbit hemorrhagic disease on

European rabbit populations in south Australia. J Wildl Dis 1998;34(2):221–7.

[9] Mitro S, Krauss H. Rabbit hemorrhagic disease: a review with special reference to its

epizootiology. Eur J Epidemiol 1993;9(1):70–8.

[10] Gaskell RM, Povey RC. Transmission of feline viral rhinotracheitis. Vet Rec

1982;111(16):359–62.

[11] Johnson RP, Povey RC. Feline calicivirus infection in kittens borne by cats persistently

infected with the virus. Res Vet Sci 1984;37(1):114–9.

[12] Poulet H, Brunet S, Soulier M, et al. Comparison between acute oral

/respiratory and

chronic stomatitis

/gingivitis isolates of feline calicivirus: pathogenicity, antigenic proﬁle

and cross-neutralisation studies. Arch Virol 2000;145(2):243–61.

[13] Geissler K, Schneider K, Platzer G, et al. Genetic and antigenic heterogeneity among feline

calicivirus isolates from distinct disease manifestations. Virus Res 1997;48(2):193–206.

[14] Doultree JC, Druce JD, Birch CJ, et al. Inactivation of feline calicivirus, a Norwalk virus

surrogate. J Hosp Infect 1999;41(1):51–7.

[15] Eleraky NZ, Potgieter LN, Kennedy MA. Virucidal eﬃcacy of four new disinfectants.

J Am Anim Hosp Assoc 2002;38(3):231–4.

[16] Scott FW. Virucidal disinfectants and feline viruses. Am J Vet Res 1980;41(3):410–4.
[17] Kennedy MA, Mellon VS, Caldwell G, et al. Virucidal eﬃcacy of the newer quaternary

ammonium compounds. J Am Anim Hosp Assoc 1995;31(3):254–8.

[18] Smid B, Valicek L, Rodak L, et al. Rabbit haemorrhagic disease: an investigation of some

properties of the virus and evaluation of an inactivated vaccine. Vet Microbiol 1991;
26:77–85.

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Feline hemotropic mycoplasmosis

(feline hemobartonellosis)

Jane E. Sykes, BVSc(Hons), PhD

Department of Medicine and Epidemiology, University of California,

2108 Tupper Hall, Davis, CA 95616, USA

Feline hemobartonellosis, or feline infectious anemia, is a bacterial

disease of domestic cats that may be associated with development of severe
hemolytic anemia. The ﬁrst documentation of an epierythrocytic organism
from an anemic cat was made in South Africa in 1942 [1]. The organism was
recognized in smears of blood and spleen that had been collected
postmortem. Because of its resemblance to Eperythrozoon species, it was
named Eperythrozoon felis, although no further studies were performed.
Recognition of the infection in the United States did not occur until 1953,
when Flint and Moss described ecxperimental induction of anemia by way
of intraperitoneal injection of infected blood from an anemic cat from
Colorado [2]. In 1955, the name Haemobartonella felis was suggested for the
organisms, because unlike Eperythrozoon species, they rarely were seen free
in the plasma, and ring forms were identiﬁed uncommonly [3,4]. The
infection then was recognized in cats from other US states, including
California, Idaho, Utah, Nebraska, Kansas, Minnesota, and Massachusetts
[5,6], and subsequently was found to have a worldwide distribution [7–17].

Recently, sequence analysis of the 16S rRNA genes of Haemobartonella

and Eperythrozoon species revealed that they are in fact mycoplasmas [18].
Because these organisms are unrelated to Bartonella, and because there are
other infectious causes of anemia in cats (such as Cytauxzoon felis, Babesia
felis

, and feline leukemia virus), it is suggested that the terms ‘‘feline

hemobartonellosis’’ and ‘‘feline infectious anemia’’ be abandoned and
replaced by ‘‘feline hemotropic mycoplasmosis (FHM).’’ Hereafter, these
organisms will be referred to as hemotropic mycoplasmas. The trivial name
‘‘hemoplasmas’’ also has been suggested for these organisms [19].

Vet Clin Small Anim

33 (2003) 773–789

E-mail address:

jesykes@ucdavis.edu

0195-5616/03/$ - see front matter

doi:10.1016/S0195-5616(03)00027-5

Etiologic agent

Hemotropic mycoplasmas are small (0.3 to 0.8 lm), epierythrocytic,

gram-negative organisms that infect a variety of mammalian species,
including people [20]. The small size of the organisms led to initial proposals
that they were actually viruses, based on their ﬁlterability [21]. Subsequently,
they were classiﬁed in the family Anaplasmataceae, order Rickettsiales,
because they lacked an outer membrane, were unculturable, and were para-
sitic to erythrocytes.

Two organisms have been identiﬁed in cats, and have hitherto been

classiﬁed as diﬀerent variants of Haemobartonella felis [18,22–25]. The Ohio
strain (large form) is apparently more pathogenic, whereas the more recently
identiﬁed California strain (small form) has not been associated with disease
in immunocompetent cats. On blood smears, the large form is pleomorphic,
varying from cocci to small rings and rods, and sometimes forming short
chains of three to six organisms. In situ hybridization was used recently to
prove that these bodies were associated with Mycoplasma haemofelis DNA
[26]. The small form appears as small cocci, 0.3 lm in diameter, although
M haemofelis

and ‘‘Candidatus M haemominutum’’ may not always be

reliably distinguished by blood smear alone [27].

Analysis of the 16S rRNA genes of these strains has showed them to be

only 83% similar, with the small form of the organism being more closely
related to Eperythrozoon species than to the large form. With the
reclassiﬁcation of Haemobartonella and Eperythrozoon species as mycoplas-
mas, the large form was renamed M haemofelis and the small form
‘‘Candidatus M haemominutum’’ [24,28]. The Candidatus status for newly
identiﬁed hemoplasmas reﬂects a provisional classiﬁcation, which will be
maintained until more information becomes available that supports the
classiﬁcation.

The 16S rRNA gene of M haemofelis is greater than 99% homologous

with that of M haemocanis (previously H canis), which causes anemia in
splenectomized dogs. In fact, using this method of classiﬁcation, some
M haemofelis

isolates are more similar to M haemocanis than they are to

other M haemofelis isolates, so it has been suggested that M haemofelis and
M haemocanis

may be the same organism infecting diﬀerent species of

animals.[29,30]. Cats inoculated with blood from a dog infected with
M haemocanis

, however, have remained asymptomatic, but blood from

these cats, when inoculated into dogs, reproduced the disease [31]. Blood
from M haemofelis-infected cats, when inoculated into splenectomized and
nonsplenectomized dogs, did not result in anemia or cytologically detectable
bacteremia, and inoculation of susceptible cats with this blood also did not
result in anemia or bacteremia [6,31]. Because of these results, and mor-
phology diﬀerences between these organisms (M haemocanis tends to form
long chains on the surface of erythrocytes), 16S rRNA gene sequencing may
not be adequate to discriminate these species [32]. Birkenheuer et al recently

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J.E. Sykes / Vet Clin Small Anim 33 (2003) 773–789

compared the RNase P genes of these two species, which accumulate
mutations more rapidly than the 16S rRNA genes [30]. Using this method,
sequence homology between M haemofelis and M haemocanis was lower
(94.3% to 95.5%), compared with that noted within strains of M haemofelis
(99.4%) and M haemocanis (97.7% to 100%). Further studies on the
relationships of these organisms are required.

To add further confusion, another canine hemotropic mycoplasma that

shares similar morphology and 93% 16S rRNA gene sequence homology
with Candidatus M haemominutum has been discovered (Sykes et al,
unpublished data, 2003). Whether this is the same or a closely related
organism awaits further study. As Eperythrozoon ovis (originally named
after its sheep host) also has been documented in goats, extension of the host
range of other hemotropic mycoplasmas seems possible.

The hemotropic mycoplasmas form a cluster of organisms within the

mycoplasmas that is most closely related to the fastidiosum cluster.
The latter cluster contains M fastidiosum and M cavipharyngis, which are
fastidious, glucose-fermenting mycoplasmas that have been isolated from
horses [33] and guinea pigs, respectively [34]. The hemotropic mycoplasma
and fastidiosum clusters fall within the pneumoniae group, which includes
the human mycoplasmal pathogens M pneumoniae and M genitalium [35].
More detailed phylogenetic relationships between hemotropic mycoplasmas
and other related organisms are represented in Fig. 1. The genome of M
haemofelis

has been sequenced partially [36]. With further sequencing of

the genomes of other organisms and the inevitable discovery of new
organisms, the hemoplasmas likely will form a large cluster within the
mycoplasmas.

The reclassiﬁcation of these organisms as mycoplasmas may seem

inappropriate initially based on their association with erythrocytes; other
mycoplasmal pathogens generally associate with mucosal surfaces of the
respiratory and urogenital tracts, occasionally causing arthritic disease.
Even 35 years ago, however, it was noted that Haemobartonella and
Eperythrozoon

resembled mycoplasmas through their lack of cell walls and

susceptibility to tetracyclines [37]. Several additional similarities between
hemotropic mollicutes and other members of the pneumoniae group have
been identiﬁed, including a metabolic pattern resembling that of the
fermentative mollicutes and the possession of bleb structures involved in
host cell attachment. Like M haemocanis, M haemofelis, and M haemosuis,
the human pathogen M pneumoniae may induce cold agglutinin formation
in the infected host [19], and elevated reticulocyte counts have been detected
in 64% of patients infected with M pneumoniae [38,39]. Cold agglutinins
induced by M pneumoniae usually are directed against the carbohydrate
antigen I, which also may be the case for M haemofelis. In addition, M
pneumoniae

can enter the blood and bind to this antigen on erythrocytes

[40]. Reclassiﬁcation of Haemobartonella and Eperythrozoon species as
mycoplasmas undoubtedly will assist in future studies of these organisms.

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Epidemiology

Because of the poor sensitivity of available diagnostic tests for hemo-

tropic mycoplasmas, few accurate epidemiologic studies of feline infection
were reported before 2000. Organisms were detected cytologically in 374
of 43,514 (0.9%) cats presenting to veterinary teaching hospitals in the
United States from March 1964 to October 1971 [41], and 5 of 97 (5.2%) cats
in the United Kingdom in 1960 [7]. Another study in the United Kingdom
found a prevalence of 28% (15/57) when acridine orange stain was used, but
only 5% when Giemsa was used [42]. Using a combination of four diﬀerent
stains, the prevalence in Scotland was reported as 23.2% in 155 cats
examined between November 1979 and July 1980 [43]. Finally, Grindem et al
detected the organism in Giemsa-stained blood smears collected from 6 of
123 (4.9%) cats from Wake County, NC. This included 3 of 83 (3.6%)
healthy cats and 3 of 40 (7.5%) ill cats, although a few of the healthy cats
were anemic or tested positive for feline leukemia virus (FeLV) [44].

Fig. 1. Phylogenetic tree based on sequence analysis of 16S rRNA genes showing relationships
between the hemotropic mycoplasmas and other members of the class Mollicutes. Boostrap
percentage values are given at the nodes of the tree; Asteroleplasma anaerobium was used as the
outgroup.

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Subsequently, Jensen et al used the more sensitive polymerase chain

reaction (PCR) to study the epidemiology of FHM [45]. Positive results were
obtained in 23 of 82 (28%) cats suspected to have FHM (based on the
presence of fever, anemia, or visible organisms on blood smears), and 20 of
138 (14.5%) control cats (from which complete blood cell counts [CBCs]
had been obtained for other reasons). The suspect cat samples originated
from various parts of the United States, whereas most of the control cats in
this study came from Colorado. Signiﬁcantly greater numbers of suspect
cats were infected with M haemofelis (9 of 82 cats, 12.2%) or both M
haemofelis

and ‘‘Candidatus M haemominutum’’ (4 of 82 cats, 4.9%) than

control cats (0 of 138 cats and 1 of 138 cats, respectively). Also, anemic cats
(packed cell volume [PCV] of less than 25%) were more likely to be infected
with M haemofelis (4 of 28 cats, 14.3%) or both species (2 of 28 cats, 7.1%)
than nonanemic cats (3 of 128 cats [2.3%] and 0.8% [1 of 128 cats],
respectively). Suspect (9 of 82 cats [11.0%]) and anemic cats (2 of 28 cats
[7.1%]) were equally likely to be infected with ‘‘Candidatus M haemominu-
tum’’ as control (19 of 128 cats [13.8%]) and nonanemic (17 of 128 cats
[13.3%]) cats. The same PCR assay was used in for an epidemiologic study
of 426 cats in the United Kingdom, which found a similar overall prevalence
of ‘‘Candidatus M haemominutum’’ (16.9%), but few cats were infected with
M haemofelis

(1.4%) or dually infected (0.2%). There was no association

between infection with ‘‘Candidatus M haemominutum’’ and anemia.
Preliminary results of studies performed at the University of California,
Veterinary Medical Teaching Hospital (using the same PCR assay) have
revealed a prevalence of 11.5% (11 of 96 cats) in cats presenting with a PCV
of less than 29%, but only ‘‘Candidatus M haemominutum’’ has been
identiﬁed to date (Sykes et al, unpublished data, 2003) These cats were not
necessarily suspected to have FHM.

Several risk factors have been identiﬁed for FHM (Table 1). A male

predisposition has been apparent in most studies [5,7,44,46,47]. The relative
risk associated with being male was estimated as 2.5 in a study of 374 cats

Table 1
Risk factors associated with infection by feline hemotropic mycoplasmas

Risk factor

Reference(s)

Male gender

[5,7,45,47,48]

Age less than 4–6 years

[42,45,47]

Presentation during the spring and summer months

[42]

Positive FeLV status

[45]

History of cat bite abscesses

[45]

Absence of current vaccinations

[45]

Outdoor status

[45]

Presence of anemia

[45]

History of anemia

[45]

Abbreviations:

FeLV, feline leukemia virus.

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J.E. Sykes / Vet Clin Small Anim 33 (2003) 773–789

from 11 diﬀerent institutions in the United States [41]. Cats younger than 4
to 6 years of age are also at higher risk of developing FHM, with the risk
declining in older cats [41,44,46]. The frequency of FHM also may be higher
in the spring and summer months, presumably as a consequence of in-
creased roaming of cats outdoors [41]. A positive FeLV status also has been
identiﬁed as a risk factor [44]. Additionally, a history of cat bite abscesses,
absence of current vaccinations, and outdoor roaming has been identiﬁed as
risk factors [44].

The mode of transmission of feline hemotropic mycoplasmas remains

enigmatic. Cats housed together for prolonged periods do not transmit the
disease [6]. Transmission has been accomplished experimentally by par-
enteral injection and oral administration of infected blood. As kittens
have been found to be infected as early as 3 hours after birth, vertical
transmission (by way of placental or lactogenic spread) seems likely [48].
Ticks, lice, and ﬂeas, have been suggested as vectors, as is known for many
other hemotropic mycoplasmas [8,19,49]. Some epidemiologic studies have
incriminated the presence of ectoparasites as risk factors [43], whereas in
most studies, there has been little correlation between the presence of
ectoparasites and infection with the organism [4,8,41,44,45,50,51]. Nearly
half (10 of 23 of positive samples) of the positive samples from suspect cats
in one study were from areas where ﬂea infestations are uncommon [45].
Mosquito-borne transmission remains a possibility, as documented for
M haemosuis

[52]. Because of the large number of cats with a recent history

of cat bite abscesses, spread by biting also has been suggested [8].

Clinical signs and laboratory abnormalities

Infection with M haemofelis can result in disease even in immunocom-

petent cats. Depression, inappetence, and dehydration are common signs,
and some cats also may present with weight loss. Anemia results in signs of
weakness, pallor of the mucous membranes, tachypnea, tachycardia, and
occasionally syncope. Some owners may report their cat eats dirt or litter or
licks cement. Other physical examination abnormalities may include cardiac
murmurs, sometimes splenomegaly, and less commonly icterus. Some cats
may be febrile, and moribund cats may be hypothermic.

Autoagglutination may be noted in blood smears from some infected

cats. The most characteristic abnormality on the CBC is regenerative ane-
mia, with anisocytosis, reticulocytosis, polychromasia, Howell-Jolly bodies,
and sometimes marked normoblastemia (increased circulating nucleated
red blood cells). Erythrophagocytosis occasionally is observed in periph-
eral blood [13,53]. Nonregenerative anemia also may be noted. In some
cases this is because suﬃcient time for a regenerative response has not
elapsed. In others, anemia is nonregenerative as a result of concurrent FeLV
infection, although macrocytic, normocytic nonregenerative anemias have

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been documented in FeLV-negative cats with FHM [54,55]. White blood cell
counts may be normal, elevated, or low. Platelet counts may be increased
[47]. The serum chemistry proﬁle may reveal elevated alanine aminotrans-
ferase (ALT) as a result of hypoxia, hyperbilirubinemia, and prerenal azo-
temia. Hyperproteinemia may be seen in some cats.

Infection with ‘‘Candidatus M haemominutum’’ appears to result in mild

or absent clinical signs in immunocompetent cats [23,45]. Transient fever or
hypothermia may occur after infection [22]. Experimental infections have
shown a decline in the hematocrit and leukocyte counts, but they usually
remain within reference ranges [23]. Cats concurrently infected with FeLV
may develop more severe anemia, with signs resembling those described for
infection with M haemofelis [56].

Pathogenesis

The pathogenesis of FHM has been divided into preparasitic, acute,

recovery, and carrier phases [57]. After inoculation of experimental cats with
M haemofelis

, there is a variable delay of 2 to 34 days (most commonly about

2 to 3 weeks) before acute onset of clinical signs, which corresponds to the
preparasitic phase [22,23]. The preparasitic phase tends to be longer if the
organism is administered orally. Experimentally, anemia and parasitemia
then occur and generally persist for about 18 to 30 days (acute phase).
Mortality is highest during this phase. In the ﬁeld, onset is reported more
commonly as gradual, with associated signs of weight loss, progressive
lethargy, and dehydration. In surviving cats, the hematocrit then returns to
normal or near-normal (recovery phase), and organisms disappear from
blood smears. Despite organism disappearance, recovered cats remain
persistently infected for years (carrier phase), the organism somehow evading
the host immune system [53,58]. Intermittent reappearance of organisms in
chronically infected cats, with relapse of anemia, has been documented in
some studies [12,23,44,58].

Inoculation of cats with ‘‘Candidatus M haemominutum’’ is associated

with mild or subclinical infection that, as with M haemofelis, persists in the
face of the immune response [23]. After experimental infection, organisms
are visible for 1 to 3 weeks, beginning around day 14 after inoculation. As
described for M haemofelis, cycles of recrudescence have been documented
in chronically infected cats [23].

In some cats infected with M haemofelis, cyclical changes in the he-

matocrit and numbers of infected erythrocytes occur, with sharp declines in
the hematocrit correlating with appearance of large numbers of organ-
isms in blood smears [23,53,58]. The number of infected erythrocytes
may decline from 90% to less than 1% in less than 3 hours [53,57].
Sequestration of organisms in splenic and pulmonary macrophages has been
suggested to explain their disappearance from blood smears.

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It has been suggested that recrudescence of severe disease may follow stress,

pregnancy, intercurrent infection, or neoplasia. Unfortunately, attempts to
reproduce disease reactivation experimentally through abscesses, glucocorti-
coids, cyclophosphamide, and splenectomy have been disappointing [58].
Administration of dexamethasone or methylprednisolone caused a decline in
the hematocrit and reappearance of organisms on blood smears, although
these were mild [58,59]. Also, FeLV-infected cats concurrently infected with
‘‘Candidatus M haemominutum’’ failed to develop repeat episodes of
hemolytic anemia despite development of severe FeLV-related illnesses,
including peritonitis and lymphoma [56]. In contrast to hemotropic
mycoplasma infections of other host species, splenectomy has a variable
eﬀect on the course of FHM [2]. Recrudescence of anemia and parasitemia has
been documented in some chronically infected cats [4,7], although other
studies suggest splenectomy increases the number of visible organisms in
blood smears without causing signiﬁcant anemia [57,58].

Anemia results predominantly from extravascular hemolysis; intravas-

cular hemolysis is not a prominent feature of the disease. The organism
appears to locate itself in an indentation on the erythrocyte surface, which
persists after the organism detaches; ﬁne ﬁbrils connect the organism with
the erythrocyte [60,61]. Positive direct Coombs tests may be noted in cats
with FHM [62,63]. Cold agglutinins have been documented in cats, dogs,
and pigs infected with hemotropic mycoplasmas [63–66] and were shown in
one study of FHM to correlate roughly with development of severe anemia
[63]. These antibodies only agglutinated parasitized erythrocytes and
neuraminidase-treated erythrocytes; normal erythrocytes were unaﬀected,
suggesting the antibodies may be directed against altered I-membrane
antigen complex. Increased osmotic fragility of erythrocytes has been
demonstrated in cats with FHM [62,67], and infected cells take on a spherical
appearance when examined using electron microscopy [61]. Red blood cell
lifespan also is decreased [68]. Ultrastructural studies have revealed an
increased proportion of erythrocytes containing crystalloid inclusions in
infected cats, the percentage of cells containing inclusions increasing with
worsening bacteremia [69]. These inclusions may reduce the deformability of
erythrocytes and promote their removal from the blood by the spleen. The
organisms are also capable of ‘‘bridging’’ adjacent erythrocytes, which also
might promote splenic trapping and removal of red blood cells.

Several authors have suggested that feline hemotropic mycoplasmas may

lower cats’ resistance to concurrent infections [48,70]. Such a phenomenon
has been documented in other host species infected with hemoplasmas,
particularly laboratory animals, in which underlying hemotropic myco-
plasma infections confounded studies of other pathogens [65,71,72]. For
example, infection of mice with E coccoides leads to enhanced susceptibility
to other infections, including mouse hepatitis virus (MHV) [72]. The as-
sociation between M haemofelis and FeLV ﬁrst was recognized in the
1970s [73]. In some studies, concurrent infection with FeLV worsened

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expression of FHM. Severe macrocytic, hypochromic anemia; splenomeg-
aly; marked parasitemia; and resistance to tetracycline therapy were more
likely in coinfected cats than in cats infected with FeLV or hemoplasmas
alone [46,55]. In contrast, another study described development of severe
anemia even in the absence of concurrent FeLV infection [54]. Strain
diﬀerences may explain these discrepancies. Nevertheless, in several studies,
the prevalence of FeLV antigenemia in cats infected with hemotropic
mycoplasmas has been high (up to 48%) [43,44]. Concurrent occult infection
with hemoplasmas should be considered in any FeLV-positive cat with
macrocytosis, even in the absence of reticulocytosis [74]. Infection of FeLV
positive cats with hemoplasmas may enhance the virulence of FeLV. When
4-month old cats were infected with FeLV alone or FeLV and hemotropic
Mycoplasma

species, only coinfected cats developed viremia followed by

erythroid aplasia [75]. Furthermore, cats coinfected with FeLV and
‘‘Candidatus M haemominutum’’ were more likely to develop clinically
signiﬁcant anemia than cats infected with ‘‘Candidatus M haemominutum’’
alone, and myeloproliferative disease developed 6 to 7 weeks after
inoculation of FeLV-infected experimental cats with ‘‘Candidatus M
haemominutum’’ [56]. Induction of myeloproliferative disease with the
strain of FeLV used in that study had not previously been possible. In
another case-control study of 297 cats, cats with FHM were 12-fold more
likely to develop hematopoietic neoplasia than age- and gender-matched
cats without FHM, although those cats were not speciﬁcally tested for FeLV
[73]. Proposed mechanisms for accelerated neoplastic transformation have
included stimulation of erythroid mitosis, leading to a greater chance of
neoplastic transformation in erythroid precursor cells, or decreased
resistance to neoplasia secondary to immune stimulation.

Conversely, feline immunodeﬁciency virus (FIV) infection does not

appear to enhance the pathogenicity of M haemofelis, nor does M haemofelis
infection accelerate the progression of FIV infection [46,76]. In a study of
feral farm cats, a negative correlation was noted between FIV seropositivity
and infection with hemotropic mycoplasmas [77].

Finally, it is possible that infection with one hemoplasma species may

exacerbate disease caused by another. For example, cats chronically infected
with M haemominutum, when subsequently inoculated with M haemofelis,
appeared to experience more severe and prolonged disease than cats inoc-
ulated with M haemofelis alone [22]. Inoculation of one M haemofelis carrier
cat with M haemominutum did not precipitate disease. Unfortunately, inability
to standardize organism numbers in experimental inocula confounds com-
parisons of disease severity, which simply may reﬂect dose-dependent eﬀects.

Immune response

Much remains unknown regarding immunity to hemotropic mycoplasma

infections in cats, but recovered cats seem to be susceptible to reinfection [4].

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Cats infected with M haemofelis failed to produce antibodies that cross-
reacted with ‘‘Candidatus M haemominutum’’ [23], whereas cats infected
with M haemominutum developed antibodies that were somewhat cross-
reactive to M haemofelis. In one study, infection with M haemominutum
protected cats against challenge with M haemofelis [24]. Nevertheless, dual
infections with these two species have been documented experimentally and
naturally [22,45].

The immune response to M haemofelis has been evaluated [57]. Five

major antigens with molecular masses of 14, 45, 47, 52 and 150 kDa were
identiﬁed. The most immunodominant antigens were the 14, 52 and 150 kDa
antigens, which were designated major antigens 1, 2, and 3, respectively. The
potential usefulness of these antigens for diagnosis or immunization awaits
investigation.

Diagnosis

Attempts to isolate feline hemotropic mycoplasmas in the laboratory

have been unsuccessful, including those using mycoplasma media and fresh
cat blood agar [24]. As a result, until recently, diagnosis has relied on
detection of the organism following cytologic examination of blood smears.
Unfortunately, because of the cyclical nature of infection, organisms are
often not visible on blood smears, and the sensitivity of this method
has been estimated to be less than 50% [22]. Careful examination of
Romanowsky-stained blood smears is required, and the organisms fre-
quently are confused with Howell-Jolly bodies, Heinz bodies, stain
precipitate, Pappenheimer bodies, and moisture artifacts, resulting in over
diagnosis of the disease. Optimum methods of preparation, staining, and
examination of smears to facilitate accurate identiﬁcation of hemotropic
mycoplasmas have been described [78,79]. Ideally, blood smears should be
made immediately after sample collection. Some have recommended making
fresh smears or collecting blood into citrate tubes, as organisms tend to
detach from erythrocytes after addition of EDTA. The low concentration of
EDTA in blood collection tubes might not aﬀect organism attachment,
however [27]. Acridine orange stain, which binds nucleic acid and can be
visualized using a ﬂuorescence microscope, has been shown to improve
sensitivity [42,80]. Unfortunately, this requires ﬁxation of blood samples in
formalin before blood smear preparation, and the stain and ﬂuorescence
microscopies are not routinely available. Other studies have not shown
improved sensitivity using this stain [59].

The advent of PCR technology has improved the ability to detect H felis

substantially. Several PCR assays have been described, all of which are
based on detection of the 16S rRNA gene [23,24,45,59,81]. These assays
detect as few as 130 organisms/mL of blood and have been shown to be
signiﬁcantly more sensitive than blood smear evaluation, although it may
not detect the organism in asymptomatic carrier cats consistently

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[22,23,45,59,81]. These assays are available in some laboratories in the
United States. In experimental models, PCR becomes positive well before
clinical signs of infection appear, and before seroconversion [23]. Not all
assays detect both M haemofelis and M haemominutum [24]. The assay
described by Jensen et al is capable of detecting and diﬀerentiating between
M haemofelis

and M haemominutum, generating PCR products that diﬀer in

size for each organism [45]. Accurate identiﬁcation of the infecting species is
important, because PCR results correlate least well with clinical illness in
cats infected with M haemominutum. Although some asymptomatic cats test
positive for M haemofelis, in naturally infected cats, there is a strong
correlation between PCR positivity for M haemofelis and anemia. Because
of lack of standardization of PCR assays between laboratories, together
with variable quality control, the results of PCR testing for feline hemo-
plasmas should be interpreted with caution and in light of the clinical ﬁnd-
ings. Samples should be submitted before antimicrobial therapy, which will
cause PCR results to become negative. A real-time, or quantitative PCR
assay has been developed for feline hemotropic mycoplasmas, which allows
determination of the numbers of organisms in the blood [82]. Further
studies are required to determine whether results of this assay can predict
clinical disease and be used to assess response to antimicrobials.

Despite the inability to culture these organisms, serologic assays for hemo-

tropic mycoplasmas using immunoﬂuorescent antibody technology have
been reported. These assays have been used to detect antibodies directed
against feline hemotropic mycoplasmas and against other hemotropic
mycoplasmas such as E coccoides and H muris. Blood smears from acutely
infected cats were used as a substrate for the feline assays [23]. Seroconver-
sion occurred in most cats by day 21 after inoculation. The maximum
titer (400) was reached 1 week later. Antigens such as major antigen 1,
which is recognized consistently throughout the course of infection [57],
represent potential diagnostic antigens for future serologic assays. The re-
sponse to this antigen occurs as early as 14 days postinoculation and persists
for at least 2 months.

Treatment

Treatment is indicated for cats with clinical signs and laboratory

abnormalities consistent with FHM. Treatment of PCR-positive, healthy
cats is not recommended, as no regime has been identiﬁed that completely
eliminates the organism [27]. Furthermore, reinfection of these cats may
occur after treatment.

Tetracycline administration is associated with rapid clinical improvement

with minimal adverse eﬀects; therefore tetracyclines have been the drugs of
choice for treatment of FHM. Because of a low incidence of adverse eﬀects
and infrequent administration, doxycycline is recommended. A dose of 5
to10 mg/kg by mouth every 24 hours for 2 to 3 weeks has been suggested.

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Doxycycline liquid is preferred over tablets, as tablet administration has been
associated with esophageal stricture formation [83,84]. Despite clinical
improvement with this therapy, complete elimination of the organism does
not seem to occur. Studies using PCR have shown reappearance of
hemoplasma DNA in the blood 3 days to 5 weeks after discontinuation
of doxycycline therapy (2.5 to 5 mg/kg by mouth every 12 hours for 10 to
21 days) [23,59]. The macrolide antibiotic azithromycin, which also has
antimycoplasmal properties, was ineﬀective for treatment of FHM using
an experimental model [22]. The treatment regime evaluated was 15 mg/kg by
mouth every 12 hours for 7 days. Trials with imidocarb diproprionate also
have had disappointing results [85]. In contrast, oral administration of enro-
ﬂoxacin appears to be an eﬀective alternative treatment to doxycycline [86].

Other drugs investigated include penicillin and chloramphenicol.

Although some response to penicillin has been reported [2], the absence of
a cell wall in these organisms implies resistance to penicillin. The antimy-
coplasmal drug chloramphenicol also has been suggested [2], although some
cats have shown a more prolonged response to tetracycline than to chlor-
amphenicol, and FHM has developed in some cats within days of initiating
chloramphenicol treatment for upper respiratory infections [8,87]. Chlor-
amphenicol also has bone marrow suppressive eﬀects, which has potential to
interfere with erythroid regeneration.

Some authors have recommended the concurrent use of prednisone to

suppress secondary immune-mediated hemolytic anemia. The recommended
dose is 1 mg/kg by mouth every 12 hours, which subsequently is tapered over
3 weeks. In many cases, this has been necessary because of the diﬃculty
diﬀerentiating between FHM and primary immune mediated hemolyhcane-
mia (IMHA) in the absence of sensitive PCR assays. As glucocorticoid
treatment can increase organism numbers in the blood, and because they can
exacerbate unidentiﬁed concurrent infections, controlled clinical trials are
required to investigate whether addition of glucocorticoids improves
outcome in cats with FHM. Treatment with antimicrobials alone has led to
successful resolution of FHM in many cases.

Finally, transfusions with packed red blood cells may be required for cats

with severe, rapidly developing anemia with accompanying signs of weak-
ness, lethargy, or inappetence. Because of the existence of natural auto-
antibodies in cats, cross-matching always should be performed before
transfusion. Fluids and blood should be administered judiciously to avoid
ﬂuid overload, especially in cats presenting with cardiac murmurs. If possi-
ble, thoracic radiography and echocardiography should be performed in
these cats before aggressive ﬂuid therapy.

Prevention

Inadvertent transmission by blood transfusion of blood from carrier cats

has been documented [2]; therefore blood donors should be tested for

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hemoplasmas using PCR assays. Keeping cats indoors also is likely to
prevent infection, as outdoor status has been identiﬁed as a risk factor.
Control of ﬂeas and ticks using products such as ﬁpronil is recommended.

Public health signiﬁcance

Organisms resembling hemotropic mycoplasmas occasionally have been

documented in people, including anemic patients with AIDS and systemic
lupus erythematosus [20,88]. Archer et al also reported on an epierythrocytic
organism in people, but its ultrastructural appearance diﬀered from that
reported for the hemotropic mycoplasmas [89]. As blood-feeding vectors
such as mosquitoes and ticks have the potential to transmit hemotropic
mycoplasmas to multiple host species, and closely related hemoplasmas have
been identiﬁed in more than one host species, these organisms should be
treated as potential zoonoses until more is known regarding their host ranges.

Summary

Hemotropic mycoplasmas represent an important cause of anemia in cats

worldwide. Previously known as Haemobartonella species, sequencing of the
16S rRNA genes of these organisms has led to their reclassiﬁcation as
mycoplasmas. Two species have been identiﬁed in cats, M haemofelis and
‘‘Candidatus M haemominutum.’’ The latter organism alone has not been
associated with disease in naturally infected cats but may cause anemia in
FeLV-infected cats and accelerate development of FeLV-induced myelo-
proliferative disease. The mode of transmission of these organisms re-
mains enigmatic. Nevertheless, development of sensitive DNA-based tests
for these unculturable organisms has improved the understanding of the epi-
demiology and pathogenesis of FHM. Cats with clinical signs and laboratory
abnormalities consistent with FHM should be treated with doxycycline;
enroﬂoxacin may represent an eﬀective alternative. Transfusion with packed
red blood cells after cross-matching may be required for severely anemia cats,
and addition of prednisone may be required if the diagnosis of FHM is
uncertain, or response to antimicrobials alone is insuﬃcient. Aﬀected cats
should be tested for FeLV, the most common concurrent infection in cats
with FHM.

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Leptospirosis

A re-emerging zoonotic disease

Cathy E. Langston, DVM

*, Kerry J. Heuter, DVM

Renal Medicine Service, Animal Medical Center, 510 East 62nd Street,

New York, NY 10021, USA

Animal Medical Center, 510 East 62nd Street,

New York, NY 10021, USA

Leptospirosis is a zoonosis of worldwide signiﬁcance, caused by infection

with pathogenic Leptospira species. The taxonomy and classiﬁcation of
these organisms are varied and complex. Before 1989, the genus Leptospira
was divided into one of two species: Leptospira interrogans and L biﬂexa.
L interrogans

is the only pathogenic species, and it is distributed worldwide

in approximately 160 mammalian species. Within L interrogans alone, there
have been over 200 serovars recognized.

In dogs and cats, infection is caused by at least eight distinct serovars

(Table 1). Each serovar is maintained in nature by one or more subclinically
infected wild or domestic animal reservoir hosts. These hosts then serve as
a potential source of infection and illness for people and other incidental
animal hosts. When infected, incidental hosts will develop a more severe
clinical illness and shed organisms for shorter period of time [1].

Leptospires are thin, ﬂexible, ﬁlamentous (0.1 to 0.2 lm wide

6 to 12 lm

long) bacteria made up of ﬁne spirals with hook-shaped ends (Fig. 1). They
are composed of a protoplasmic cylinder that is wound around a straight
central axial ﬁlament. The outer envelope is composed of lipopolysaccharide
and antigenic mucopeptide. The composition of the lipopolysaccharide is
similar to that of other gram-negative bacteria, but it has lower endotoxic
activity. Leptospires are motile, making writhing and ﬂexing movements
while rotating along their long axis. Leptospires are unique among
spirochetes, because they have terminal hooks. A helically shaped cell cylin-
der and two periplasmic ﬂagella enable the organisms to burrow into tissue.

Vet Clin Small Anim

33 (2003) 791–807

* Corresponding author.
E-mail address:

cathy.langston@amcny.org (C.E. Langston).

0195-5616/03/$ - see front matter

doi:10.1016/S0195-5616(03)00026-3

The leptospires are aerobic bacteria that require special media and

conditions for growth. Strains can be cultivated in media containing 10%
rabbit serum or 1% bovine serum albumin plus long-chain fatty acids.
Media must be kept at a pH of 6.8 to 7.4 and a temperature of 28

to 30

An incubation period of a few days to 4 weeks is required. Leptospires
cannot be seen on gram-stained smears and are faintly colored by Giemsa or
Wright’s stain. They can be seen on dark-ﬁeld microscopy, however [2].

The most common serovars seen in cases of canine leptospirosis in the

United States and Canada have been canicola, icterohaemorrhagiae,
grippotyphosa

, pomona, and bratislava. The overall prevalence of leptospi-

rosis cases declined substantially from the early 1970s to the early 1980s, after
introduction of a bivalent vaccine for serovars canicola and icterohaemor-
rhagiae

in the early 1970s [3]. Although the number of cases of clinical disease

had declined, serologic studies over the past 50 years had found a signiﬁcant
number of seropositive dogs, ranging from 10% to 38%, with diﬀerent

Table 1
Common serovars of Leptospira interrogans sensu lato infecting animals

Serovar

Primary reservoir hosts

Canicola

Dog

Icterohaemorrhagiae

Rat

Grippotyphosa

Vole, raccoon, skunk, opossum

Pomona

Cow, pig, skunk, opossum

Hardjo

Cow

Bratislava

Rat, pig, horse?

Autumnalis

Mouse

Bataviae

Dog, rat, mouse

Fig. 1. Leptospires (arrows) in renal tubule (Warthin-Starry stain

1000). (From Shelley

Newman, DVM, DACVP, Animal Medical Center, New York, NY.)

792

C.E. Langston, K.J. Heuter / Vet Clin Small Anim 33 (2003) 791–807

predominating serovars [4–10]. Direct comparisons of these surveys are
diﬃcult because of varied cut-oﬀ values of positive titers and inconsistencies
in serovars tested. The prevalence of clinical disease began increasing again in
the mid 1990s [3]. The incidence of disease attributed to serovars canicola and
icterohaemorrhagiae

has decreased, whereas infections with serovars grip-

potyphosa

, pomona, and bratislava have increased [1,4–6,11–16]. The serovar

autumnalis

may be emerging as a source of clinical disease [17]. Seropositivity

and clinical disease from serovars other than icterohaemorrhagiae and
canicola

, however, have been reported in dogs for over 40 years [8,18].

Cats appear to be less susceptible to spontaneous and experimental

infections, and clinical reports of leptospirosis in cats are rare [1,19–22].
Cats will develop antibody titers, and they are probably exposed to lep-
tospires excreted by wildlife or in urine of cohabitating dogs. Transmission
from rodents carrying serovars ballum or icterohaemorrhagiae also is
suspected. Outdoor cats have the highest seroprevalence. Serovars canicola,
grippotyphosa

, and pomona have been isolated from cats, and antibodies to

serovars grippotyphosa, hardjo, autumnalis, and icterohaemorrhagiae have
been detected in naturally exposed cats [1].

Epidemiology

The disease is maintained in nature by chronic infection of the renal

tubules of maintenance hosts. Infected mammalian hosts excrete leptospires
in their urine. Infection is endemic in the maintenance host and usually is
transferred from animal to animal by direct contact. Maintenance hosts
typically are infected at an early age and do not develop clinical disease.

Leptospires are transmitted to incidental hosts by direct or indirect

contact. Direct transmission occurs through contact with infected urine,
venereal and placental transfer, bite wounds, or ingestion of infected tissues.
Direct spread of infection is enhanced by crowding of animals as may occur
in a kennel. Indirect transmission occurs through exposure of susceptible
animals to contaminated water sources, soil, food, or bedding. There is
evidence that spirochetes survive in insects and other invertebrate hosts, but
the signiﬁcance of this ﬁnding with regard to disease transmission is
unknown. Once outside the host, leptospires do not replicate [1]. Depending
on the conditions, however, leptospires may remain viable for months when
environmental factors are optimal [1,23].

The optimal survival condition for leptospires is a warm, wet environment

with neutral to slightly alkaline stagnant or slow-moving water. Hence, the
disease tends to be seasonal in temperate climates, with peak incidence
occurring when weather permits, and year-round in tropical climates.
Disease outbreaks often increase after periods of ﬂooding or increased
rainfall [14,24]. In arid areas or during drought conditions, infections of
accidental hosts are more common around water sources. Raw sewage or
other similar contamination will decrease the survival time of leptospires.

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C.E. Langston, K.J. Heuter / Vet Clin Small Anim 33 (2003) 791–807

Optimum survival in soil is favored by a neutral or slightly alkaline pH.
Spirochetes survive only transiently in undiluted acidic urine (pH 5.0 to 5.5),
whereas the opposite conditions provide more suitable habitats. Ambient
temperatures between 0

C and 25

C favor the survival and replication of

leptospires, whereas freezing markedly decreases survival. There is an
increased incidence of canine leptospirosis during late summer and early fall
and in the southern semitropical belt of the United States [1,11–13,24].

Incidental hosts that survive the acute infection will have long-term renal

colonization with shedding in urine for months to years. Although dogs are
known to be persistent renal carriers of serovar canicola, the duration of
shedding of other serovars has not been determined.

Pathogenesis

Leptospires penetrate mucous membranes or abraded skin and multiply

rapidly upon entering the blood. They then spread and further replicate in
many tissues, including the kidney, liver, spleen, central nervous system
(CNS), eyes, and genital tract. Increases in serum antibodies thereafter clear
the spirochetes from most organs, but organisms may persist in the kidney
and be shed in urine for weeks to months. The extent of damage to internal
organs varies with virulence of the organism and host susceptibility. Certain
serovars have the tendency to produce acute hemorrhagic, hepatic, or most
commonly, renal dysfunction. More than one form may occur in a given
animal, and the clinical manifestations can vary among outbreaks and
geographic areas with a given serovar.

The mechanisms by which leptospires cause disease are not well

understood. Toxins and enzymes produced by leptospires may contribute
to their pathogenicity. The clinical and pathological features of infection
have suggested the presence of an endotoxin. Leptospiral lipopolysaccharide
stimulates neutrophil adherence and platelet activation, which may be
involved in inﬂammatory and coagulatory abnormalities that occur. Several
laboratories recently have isolated a lipopolysaccharide-like substance, but
this has not been shown to contribute to the pathogenesis of leptospirosis
[2]. Toxins that may be hemolysins have been isolated in some strains [23].

Leptospirosis causes a severe vasculitis with endothelial damage, re-

sulting in injury to capillaries, tissue edema, hemorrhagic diathesis, and po-
tentially disseminated intravascular coagulation (DIC). Renal insuﬃciency
and failure are the result of tubular damage associated with colonization
and replication of the organisms in renal tubular epithelial cells [2]. Acute
impairment of renal function also may result from decreased glomerular
ﬁltration and hypoxia caused by kidney swelling that impairs renal blood
perfusion [1]. Hypovolemia and hypotension may occur in severe cases as
a result of dehydration, massive hemorrhage, or vasculitis. Myocarditis,
pericarditis, and cardiac dysrhythmias are well-documented manifestations
of leptospirosis that can lead to hypoperfusion [2].

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Hepatic involvement manifests most commonly as icterus though

decreased serum albumin levels, increased globulin levels, and impaired
production of vitamin K-dependent clotting factors also are seen. Focal
hepatocellular necrosis is the most common histopathological change.
Results of histochemical studies suggest that the fundamental hepatic lesion
is caused by subcellular eﬀect on enzyme systems [2]. The degree of icterus in
canine and human leptospirosis usually corresponds to the severity of
hepatic necrosis.

Serovars icterohaemorrhagiae and pomona have been associated most

commonly with severe hepatic dysfunction, whereas canicola and grippoty-
phosa

typically are associated with minimal liver involvement. Younger dogs

(younger than 6 months) seem to develop more signs of hepatic dysfunction
in any disease outbreak. Initial hepatic injury and persistence of the
organism in the liver can result in altered hepatic circulation, ﬁbrosis, and
immunologic disturbances that bring about a chronic inﬂammatory re-
sponse. Chronic active hepatitis has been a sequela to serovar grippotyphosa
infection in dogs. Extensive hepatic ﬁbrosis and failure may result from this
process [1].

Central nervous system involvement in people most commonly manifests

as aseptic meningitis [25]. Leptospires enter the cerebrospinal ﬂuid (CSF) in
the early septicemic phase of the illness. The meningeal signs often appear in
the second week of illness, when the leptospires are being cleared from the
CSF and antigen–antibody complex-induced inﬂammation may be respon-
sible for the symptoms [23,25]. Other than stiﬀness, disorientation, and
posterior paresis, symptoms of CNS involvement (ie, altered sensorium,
headache, neck pain, psychosis, or seizures) are reported rarely in dogs
[11,13].

Leptospires may enter the aqueous humor during the septicemic phase

and may persist for several months. Uveitis typically develops during the
second week of the illness. The persistence of the leptospires in the aqueous
humor may lead to a chronic, recurrent latent uveitis. It has been postulated
that the production of antibodies to leptospires cross-react with ocular
tissues [23]. Myalgia is a common ﬁnding in leptospirosis. Abortion or
infertility resulting from transplacental transmission of leptospires associ-
ated with serovar batavaiae infection has occurred in a dog [1].

Clinical ﬁndings

Leptospirosis may present as peracute, acute, subacute, or chronic

disease. Clinical signs in canine leptospirosis depend on the age and
immunity of the host, environmental factors aﬀecting the organisms, and
virulence of the serovar. Large breed (heavier than 15 kg), male, outdoor,
middle age dogs are aﬀected most commonly [1,3,11–14,28]. Although
historically thought of as a rural disease, dogs in urban environments are at

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C.E. Langston, K.J. Heuter / Vet Clin Small Anim 33 (2003) 791–807

risk. Peracute leptospiral infections can be manifested by massive
leptospiremia and death with very few clinical signs. The initial signs of an
acute infection include pyrexia (39.5

C to 40

C), shivering, and muscle

tenderness. Vomiting, dehydration, and shock then follow. Tachypnea,
rapid and irregular pulse, and poor capillary perfusion also have been noted.
Coagulation defects and vascular injury are apparent with hematemesis,
hematochezia, melena, epistaxis, and widespread petechiae. Terminally ill
dogs become depressed and hypothermic, while renal and hepatic failure
does not have time to develop.

Subacute infections are the most commonly recognized form of

leptospirosis. This form manifests with fever, anorexia, vomiting, de-
hydration, and increased thirst. Reluctance to move and paraspinal hyper-
esthesia in dogs may result from muscular, meningeal, or renal inﬂammation.
Mucous membranes appear injected, and petechial and ecchymotic hemor-
rhages are widespread. Conjunctivitis, rhinitis, and tonsillitis usually are
accompanied by coughing and dyspnea. Progressive deterioration in renal
function is manifested ﬁrst by polyuria and polydipsia, which may progress
to oliguria or anuria. Renal function in some dogs surviving subacute
infections may return to normal within 2 to 3 weeks, or chronic compensated
polyuric renal failure may develop.

In addition to causing icterus, intrahepatic cholestasis and necrosis from

hepatic inﬂammation may be so complete that acholic feces may develop.
Chronic active hepatitis or ﬁbrosis may develop as a sequela to leptospirosis.
Overt signs of liver failure may include chronic inappetence, weight loss,
ascites, icterus, or hepatoencephalopathy.

Gastrointestinal [GI] signs tend to be more severe and persistent in dogs

with leptospirosis compared with other causes of acute renal failure. GI
inﬂammation and ileus may be severe enough for intestinal intussusceptions
to occur. Careful abdominal palpation should be performed in dogs that
develop persistent vomiting and diarrhea. Feces will become scanty in such
cases, and hematochezia or melena will be apparent.

Respiratory signs such as labored breathing and cough occur in a small

percentage (3% to 20%) of dogs with leptospirosis [11,13,26]. Thoracic
radiographs frequently reveal a patchy alveolar or nodular pattern or
generalized interstitial pattern, predominating in the caudodorsal lung ﬁelds
(Fig. 2). Dogs with no clinical signs of respiratory involvement can have
signiﬁcant radiographic changes [11,12,26]. Pulmonary hemorrhage, edema,
acute respiratory distress syndrome, and interstitial pneumonitis are
pulmonary complications in people [27]. Hemorrhage, congestion, edema,
neutrophilic and lymphocytic inﬁltration, and bronchopneumonia have
been noted histologically in dogs [12,13] (Fig. 3).

Leukocytosis with or without a left shift is common, with total white

blood cell counts generally ranging from 16,500 to 45,000 cells/lL.
Leukopenia may be present in the leptospiremic phase. Thrombocytopenia
is common and may be present in conjunction with DIC, although

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hemorrhagic complications are thought to be caused primarily by
endothelial damage from vasculitis [1,11–14,28].

Most dogs (83% to 100%) with leptospirosis present in renal failure [11–

14]. The degree of elevation of serum urea nitrogen (BUN) and creatinine
concentrations vary with the severity of disease. Hyperphosphatemia and
hypocalcemia are encountered frequently. Electrolyte disturbances are
caused by renal and GI involvement, with hyponatremia, hypochloremia,
and hypokalemia occurring frequently. In some cases with terminal anuria,
hyperkalemia is present. Metabolic acidosis is present, caused by renal
failure and dehydration [1,11–14,28]. Urinalysis shows a typical picture of
acute renal failure, with glucosuria, proteinuria, and an active urine sedi-
ment (pyuria, hematuria) and granular casts [1,11–14,28].

Fig. 2. Thoracic radiograph of dog with leptospirosis showing typical interstitial inﬁltrate.

Fig. 3. Photomicrograph of pulmonary hemorrhage and cellular inﬁltrate in lung of dog with
leptospirosis (H&E

400).

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Abdominal ultrasonography is consistent with acute renal failure. The

kidneys may appear normal or have renomegaly. There may be increased
cortical echogenicity. A bright demarcation between the cortex and medulla
(‘‘medullary rim sign’’) and perinephric eﬀusion have been reported in dogs
with leptospirosis [12,14,29].

The hepatic involvement is usually initially less severe than the renal

involvement, and involvement includes elevations in alkaline phosphatase,
alanine transferase, aspartame transferase, and bilirubin. The peak increase
is usually 6 to 8 days after the onset of disease. Creatine kinase increases
with skeletal muscle inﬂammation. Increased concentrations of amylase and
lipase are caused by inﬂamed hepatic and small intestinal tissue and
decreased renal clearance [1,11–14,28].

Many leptospiral infections in dogs are chronic or subclinical.

Leptospirosis should be tested for when dogs present with fever of unknown
origin, unexplained renal or hepatic disease, or anterior uveitis. In addition,
healthy dogs in kennels, multidog households, neighborhoods, or other
environs where infection in other members has been documented should be
screened for leptospirosis.

Speciﬁc diagnostics

The microscopic agglutination test (MAT) is the standard serologic test

used for diagnosing leptospirosis. Serial dilutions of sera are mixed with
leptospiral organisms, and the highest dilution that agglutinates 50% of the
organisms is recorded. Most laboratories start with a dilution of 1:100, and
further twofold dilutions are performed to the end-point. This test is
somewhat serovar speciﬁc. The highest titer is considered the infecting
serovar, with lower positive titers considered to be cross-reactivity.
Commonly tested serovars for dogs include icterohaemorrhagiae, canicola,
pomona

, grippotyphosa, and hardjo. The test requires dark-ﬁeld microscopy,

and the laboratory must maintain cultures of the various serovars tested.
Thus, this test usually is referred to a commercial laboratory.

The classic conﬁrmation of leptospiral infection is based on a fourfold

rise in the MAT titer. Acute titers drawn within the ﬁrst 7 to 10 days of
infection may be negative. Convalescent titers are drawn 2 to 4 weeks later.
Ideally, paired samples should be submitted together to avoid variation in
test batches. A single titer of greater than 1:800 in an unvaccinated dog with
classic signs of leptospirosis provides presumptive evidence of leptospirosis,
although some would suggest using a cut-oﬀ of 1:3200 or higher [28,30].
Vaccination causes serovar-speciﬁc titers that are rarely greater than 1:300,
although titers as high as 1:3200 have been reported. High vaccinal titers
rarely persist for more than 3 months after vaccination [1,30].

Early antibiotic therapy will decrease the magnitude of rise in titer, such

that infected dogs may not show the classic fourfold rise on the convalescent

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sample. Titers in naturally infected dogs will decrease over several weeks to
many months. Dogs are well adapted to serovar canicola and may actively
shed leptospires with titers under 100.

Several ELISA tests have been developed for rapid diagnosis in people

[23,31]. ELISA tests have been evaluated for dogs also but have limited
availability [32–34]. This test is not serovar-speciﬁc, and a positive test
should be conﬁrmed with the serovar-speciﬁc MAT. The ELISA test
distinguishes between an IgG and IgM response. IgM titers rise within 1
week of infection and peak at 14 days after infection, whereas IgG titers are
not present until 2 to 3 weeks after infection and peak at 1 month. A high
IgM titer would suggest acute infection. Because IgM and IgG antibodies
will agglutinate leptospires, the MAT tests for both, but the MAT more
closely follows IgM titers. MAT and ELISA tests are based on detection of
an immunologic response by the host. Eﬀective tests that would allow
detection of organisms could provide a diagnosis earlier in the course of
disease but are less reliable than MAT.

Darkﬁeld microscopy can detect live leptospires in a wet mount of fresh

urine. A minimum of 10

organisms/mL must be present for detection.

Other organisms or debris can be mistaken for the characteristic shape and
writhing movements of leptospires by inexperienced operators. Because of
false positive and false negative results, this test adds presumptive evidence
of leptospiral infection but must be conﬁrmed by other methods.

Leptospires are fastidious organisms that require special growth medium,

and culture rarely is attempted for routine diagnosis. Samples for culture
should be obtained before antibiotic therapy. Leptospiremia (and presence
of leptospires in the CSF) is present during the ﬁrst week of infection, but
numbers of organisms decrease as the antibody titer rises. Blood samples for
culture should be anticoagulated with preservative-free heparin or sodium
polyethylene sulfonate (blood culture bottles), but not with citrate, which
inhibits leptospires. Urine is the ideal ﬂuid to be cultured after the ﬁrst week
of infection. Administration of a low dose of furosemide (0.5 mg/kg) just
before urine collection facilitates recovery. Fluid samples should be diluted
1:10 (vol/vol) with saline, bovine serum albumin, or culture medium to
minimize eﬀects of inhibiting substances such as antibodies. Urine should be
alkalinized to a pH of 8 or greater during transport, since leptospires cannot
survive for more than a few hours in an acidic environment. Tissue samples
can be cultured also, with liver and kidney being the preferred samples.
Postmortem tissues may contain tissue contaminants that overgrow the
fastidious leptospiral organisms; antemortem samples are preferred,
however [1].

Fluorescent antibody techniques have been developed to test for

leptospires in body ﬂuids such as blood and urine, or in tissue imprints of
liver and kidney [1,15,30]. These techniques generally do not distinguish
between serovars. Polymerase chain reaction (PCR) techniques have been
used in urine, blood, CSF, and aqueous humor to detect leptospires; urine is

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the preferred sample, however [35]. Because of the exquisite sensitivity of
this technique to contamination in the laboratory, good quality control
protocols are necessary to provide meaningful results. Neither ﬂuorescent
antibody nor PCR techniques are widely used clinically.

Typical histopathological ﬁndings in the kidney include a lymphoplasma-

cytic tubulointerstitial nephritis (Fig. 4). Leptospires can be seen in the
tubules with special stains (Warthin-Starry staining, silver impregnation,
Giemsa staining), although antibiotic therapy decreases the number of
organisms, making detection unlikely.

Treatment-speciﬁc therapy

There has been much debate over the relative merit of speciﬁc antibiotic

regimens for treatment of this disease in people. Randomized prospective
evaluations of diﬀerent antibiotics for treatment of leptospirosis in dogs are
limited. Studies in other species have varied results, which may be because of
diﬀerences in host adaptation to the serovar used, species-speciﬁc metab-
olism of antibiotic, serovar susceptibility, timing of initiation of therapy,
and methods of detection of infection [36]. Antibiotic therapy is directed
initially at clearing the leptospiremic phase and subsequently at clearing the
leptospiruric (carrier) phase.

High doses of penicillin, ampicillin, and amoxicillin can clear the

leptospiremia phase. Bacteremia usually is reduced within hours of drug
administration. Early administration inhibits bacterial multiplication, which
lessens the damage to organs like the liver and kidney. Vomiting animals
should receive parenteral therapy. Penicillin and its derivatives are the
antibiotics of choice for initial therapy [1], but doxycycline is also eﬀective at

Fig. 4. Photomicrograph of kidney from a dog with leptospirosis, showing lymphoplasmacytic
tubulointerstitial inﬁltrate (H&E

200).

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clearing leptospiremia. Doxycycline can be used in patients with renal
failure, as it is predominantly excreted in the feces. Tetracycline is neph-
rotoxic and should not be used in azotemic animals [37].

Clearing the leptospires from tissues, particularly the kidney, is more

problematic. In a study of hamsters infected with icterohaemorrhagiae,
leptospires were detected by PCR in the kidney after treatment with
ampicillin (40 and 100 mg/kg intramuscularly) and oﬂoxacin (15 and 30
mg/kg intramuscularly) for 4 days [36]. Enroﬂoxacin was not eﬀective in
hamsters in one study [38], but ﬁve of six infected hamsters survived with
treatment with ciproﬂoxacin [39]. Doxycycline (10 mg/kg intramuscularly
daily for 4 days) cleared tissues of leptospires in a hamster model [36] and is
eﬀective in people [40]. Oxytetracycline at high doses for 3 to 5 days was
eﬀective in swine [38], and a single injection was eﬀective in cattle [41].
Erythromycin and aminoglycoside antibiotics are eﬀective, but aminoglyco-
sides should not be used while renal function is impaired. Cephalosporins,
chloramphenicols, and sulfonamides are ineﬀective. The recommendation to
clear the carrier state in dogs is a 2-week course of doxycycline.

Doxycycline also has been evaluated as chemoprophylaxis against

leptospirosis in people who have a high-risk of exposure (ie, military
personnel deployed to endemic areas). Once weekly administration (200 mg
orally) markedly decreased the infection rate [42]. No information about
chemoprophylaxis in dogs is available. Because of the risk of developing
bacterial resistance, this therapy is not recommended [1].

Supportive treatment

Fluid therapy is one of the ﬁrst considerations for treatment of acute

renal failure caused by leptospirosis. Rehydration should occur over 4 to 24
hours, depending on the cardiovascular status. Most patients should be
rehydrated over a short time (4 to 6 hours). If the patient appears hydrated,
a ﬂuid volume equal to 3% to 5% of body weight should be administered to
account for clinically undetectable dehydration. Urine output should be
assessed after rehydration, and ﬂuid therapy should be tailored to ﬂuid
output (ins and outs). This method is appropriate for oliguric/anuric
patients, to avoid overhydration, and for polyuric patients who frequently
have volumes of urine exceeding the clinician’s estimate.

If urine output remains low (below 0.25 mL/kg per hour) in a hydrated

patient with blood pressure adequate to perfuse the kidneys (greater than 80
mm Hg mean arterial pressure), diuretics are indicated. Mannitol (0.5 gm/kg
administered intravenously over 20 minutes) is an osmotic diuretic, but it is
contraindicated in dehydrated or overhydrated patients. Furosemide, a loop
diuretic, may induce urine ﬂow in 20 to 30 minutes after an initial intra-
venous (IV) bolus of 2.2 mg/kg. If not, the dose can be doubled serially, up to
10 mg/kg. Higher doses may lead to ototoxicity. Furosemide and mannitol
can be given together.

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Once hydration is restored, treatment of metabolic acidosis is recom-

mended when the blood pH persists below 7.2, or the serum bicarbonate
concentration is less then 16 mEq/L. Sodium bicarbonate is dosed according
to the formula: body weight (kg)

0.3 (20–patient bicarbonate

concentration). One quarter to one third of the calculated dose can be
administered as an IV bolus, and an additional one quarter to one third of
the dose administered over the next 4 to 8 hours. Rapid administration,
excessive dosing, or administration to a patient with impaired respiratory
function can lead to paradoxical CNS acidosis.

Hyperkalemia may cause cardiac or ECG abnormalities, characterized by

bradycardia, wide, ﬂattened or absent P waves, peaked T waves, wide QRS
complex, atrial standstill, idioventricular rhythm, ventricular ﬁbrillation, or
asystole. Regular insulin (0.1 to 0.25 U/kg intravenously) with dextrose to
prevent hypoglycemia (1 to 2 gm/U of insulin as an IV bolus, followed by 1
to 2 gm/U over the next 4 to 8 hours) or bicarbonate (0.5 to 2 mEq/kg
intravenously) temporarily shifts potassium intracellularly, with an eﬀect
occurring within about 20 to 30 minutes. If a more immediate cardiopro-
tective eﬀect is needed, 10% calcium gluconate (0.5 to 1.0 mL/kg intra-
venously) can be administered as a slow IV bolus, although it does not
decrease plasma potassium concentration.

Histamine (H

receptor) blockers are used commonly to treat uremic

gastritis and include famotidine (0.5 to 1 mg/kg intravenously every 24
hours), ranitidine (2.2 mg/kg intravenously every 24 hours), or cimetidine
(2.5–5 mg/kg IV q 12 hours). Sucralfate (0.25 to 1 gm by mouth every 6 to 8
hours) aids in healing of uremic ulcers. It should be given at least 30 to 60
minutes before oral antacids. Metoclopramide, a centrally acting antiemetic,
is sometimes used (0.2 to 0.5 mg/kg subcutaneously or intramuscularly every
6 to 8 hours or 1 to 2 mg/kg per day as IV constant infusion). Meto-
clopramide is a dopamine receptor antagonist and should not be adminis-
tered concurrently with dopamine. Other antiemetics such as chlorpromazine
can cause hypotension, if the patient is not adequately hydrated. There is
little information on the use of 5-HT

serotonic receptor antagonists, like

ondansetron or dolasetron, in treating vomiting associated with renal failure
in animals.

Hypertension may accompany acute renal failure, and excessive volume

expansion (particularly in the face of oliguria or anuria) can exacerbate the
condition. Therapy may not be necessary, if the hypertension is mild, and
the renal failure is resolving rapidly. Emergency therapy may be necessary,
however, to prevent catastrophic eﬀects in more severe cases.

Because of the highly catabolic state associated with acute renal failure,

nutritional support should be started early in the course of the illness. If
vomiting can be adequately controlled pharmacologically, enteral feeding
with a feeding tube should be started. If enteral feeding is not possible,
partial or total parenteral nutrition should be instituted. Calories should
be predominantly carbohydrates with restricted protein content. Ideally,

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protein should be supplied as essential amino acids. Oral phosphate binders
(ie, aluminum hydroxide) to treat hyperphosphatemia should be used once
the patient is being fed enterally.

Adequate monitoring of patients with leptospirosis is essential. These are

acutely, severely ill patients whose status can change rapidly and dra-
matically. Urine output should be monitored in all patients with renal
failure. This is accomplished most accurately with an indwelling urinary
catheter with a closed collection system. Changes in body weight over the
course of hospitalization primarily reﬂect changes in body water content
and hydration. Other parameters, such as blood pressure, central venous
pressure, and packed cell volume, should be monitored as needed.

If conventional medical management fails to induce diuresis in an oliguric

or anuric patient, if life-threatening complications are present (ie, hyper-
kalemia, volume overload), or if azotemia fails to improve after 24 hours of
therapy, dialytic therapy (hemodialysis or peritoneal dialysis) should be
considered.

Outcome

Survival rates for dogs with leptospirosis range from 78% to 88% [11–

14], which exceeds survival rates for acute renal failure in general (43%) [43].
Many dogs will recover completely, although some will be left with residual
chronic renal failure. Patients that fail medical management and require
hemodialysis also have a favorable a survival rate of 86% [14].

Zoonosis/public health considerations

Leptospirosis is a presumed to be the most widespread zoonotic disease in

the world [44]. It is a sporadic disease in people in the United States, with
outbreaks occurring in occupational or recreational settings. Veterinarians
are considered at risk, along with farmers, workers in slaughter facilities,
animal caretakers, animal researchers, and sewer system workers [45].
Recreational activities associated with transmission of leptospirosis include
freshwater swimming, canoeing, kayaking, trail biking, hunting, and
increasingly, adventure travel and competitions involving water sports in
tropical locations [2,46]. Exposure during daily activities occurs in people
with contact with pet dogs or domestic livestock and in areas with rodent
infestation. A recent outbreak in Baltimore was associated with potential
exposure to rat urine in inner city alleys. Nineteen of 21 rats trapped in the
region were PCR positive for leptospiral organisms [47]. Cat ownership has
been associated with a lower incidence of a positive IgG ELISA test in people
in Baltimore [48].

Leptospirosis in people ranges from asymptomatic disease to fulminant

fatal disease and can be confused with common viral ailments. Mild
leptospiral disease in people is a self-limiting disease characterized by fever,

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myalgia, headache, anorexia, nausea, vomiting, and abdominal pain.
Defervescence occurs 4 to 7 days after onset of signs. The second stage of
this disease may feature aseptic meningitis and ocular manifestations. Icteric
leptospirosis (Weil’s syndrome) is characterized by fever, jaundice, and
azotemia. Supportive therapy has reduced mortality to 5% to 10% with this
form of disease [2].

Prevention

Vaccinating pet and kenneled dogs can help decrease the incidence and

severity of infection, and newer vaccines may help decrease development of
a carrier state. Vaccines against leptospirosis confer immunity only to the
serovars contained in the vaccine. The early bivalent bacterins were
produced from chemically inactivated whole cultures, which make the
vaccine relatively allergenic. A newer subunit vaccine was released in 2000.
Proteins from the outer envelope of the leptospiral organism are extracted
and puriﬁed, discarding the cellular debris. This method of manufacture
theoretically creates a vaccine that induces a protective antibody response
and is free of other antigenic material. This vaccine protects against serovars
icterohaemorrhagiae

, canicola, pomona, and grippotyphosa.

The initial series of vaccination should include a minimum of three

injections 3 to 4 weeks apart. Booster vaccinations are recommended annu-
ally (and sometimes biannually) in animals in an endemic region.

Elimination of rodent infestation is recommended to decrease exposure

to infected rat urine. Avoidance of areas with a high risk of transmission (ie,
drainage from areas with infected cattle or deer, especially if water is warm,
slow-moving, and alkaline) theoretically would be advisable.

In kennel or hospital settings, certain precautions can help decrease

transmission to other dogs. Infected dogs should be isolated. If the dog is
critically ill and needs to remain in the intensive care unit, urine should be
contained. This can be accomplished by maintenance of an indwelling closed
urinary collection system or placing absorbent pads in the cage to collect
voided urine. Urine contaminated articles should be handled as a biological
hazard. Caution is advised when cleaning cages to avoid aerosalization of
leptospires by high-pressure water jets. In the authors’ hospital, cages of
patients with suspected or conﬁrmed leptospirosis (and areas of urine
contamination outside the cage) are cleaned with bleach and disposable
paper towels; hosing is not performed. Leptospires are susceptible to deter-
gents, iodophor disinfectants, and drying [1]. Ambulatory dogs with suspec-
ted or conﬁrmed leptospirosis should not be allowed to urinate in areas
frequented by other dogs.

Because leptospirosis is a zoonotic disease, owners and all hospital

personnel in contact with a conﬁrmed or suspected case should be advised of
appropriate precautions. Gloves should be worn when handling the dog.
Goggles and face mask should be worn if urine or potentially contaminated

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material (eg, blood, saliva, or CSF) may be encountered (ie, during cage
cleaning or emptying a urinary collection system).

Summary

Leptospirosis is a re-emerging infectious disease that occurs in dogs in

urban and rural environments. It is caused by a ﬁlamentous spiral bacterium
that has a predilection for renal tubules. Acute renal failure, hepatic
dysfunction, and hemorrhagic diathesis are the most common clinical signs.
Treatment with antibiotics and supportive care can manage a high
percentage of cases successfully. Newer vaccines developed in response to
the change in frequency of certain serovars may decrease the incidence of
clinical disease. Leptospirosis aﬀects a wide variety of species and is zoonotic.

Acknowledgment

The authors gratefully acknowledge the assistance of Nyssa Reine in

preparation of this manuscript.

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Bartonellosis

Lynn Guptill, DVM, PhD

Department of Veterinary Clinical Sciences, Purdue University,

625 Harrison Street, West Lafayette, IN 47907, USA

Species of Bartonella are fastidious, gram-negative bacteria in the family

Bartonellaceae

of the a-2 subgroup of the Proteobacteria [1]. The Bartonella

species includes organisms that originally comprised the genera Bartonella,
Rochalimaea

, and Grahamella [1,2]. Bartonella is phylogenetically close to the

Rickettsia

and bacteria of the species Brucella, Agrobacterium, and Aﬁpia

[3–5]. The type species is Bartonella bacilliformis, an intracellular parasite
of human erythrocytes and endothelial cells that causes severe hemolytic
anemia and cutaneous angioproliferative lesions in people. It is endemic to
some countries in South America and is transmitted among people by
Lutzomyia

species sand ﬂies [1,6]. B (Rochalimaea) quintana caused trench

fever in World War I and is known to cause endocarditis, bacillary
angiomatosis, bacillary peliosis, and chronic lymphadenomegaly in people. It
is transmitted among people by the human body louse [7,8]. B (Rochalimaea)
henselae

causes bacteremia in healthy cats and was detected by polymerase

chain reaction (PCR) in tissues of ﬁve sick dogs [9–12]. It causes cat scratch
disease (CSD), bacillary angiomatosis, bacillary peliosis, relapsing fever with
bacteremia, meningitis, encephalitis, neuroretinitis, and endocarditis in
people [11,13–17]. B. clarridgeiae causes bacteremia in healthy cats [18–21].
It was serologically associated with a CSD-like condition in two people
[22,23], was associated with aortic valve endocarditis in a dog using culture
and PCR testing [24], and was detected by PCR in the liver of a dog with
hepatopathy [12]. B koehlerae [25] and B bovis (formerly B weissii) [26] were
isolated from two healthy cats each, and their pathogenic signiﬁcance in cats
or other species is not known. B (Rochalimaea) elizabethae was isolated from
a person coinfected with HIV and endocarditis [27] and was detected by PCR
in the blood of one sick dog [10]. B vinsonii subspecies vinsonii has been
isolated from voles. B vinsonii subspecies berkhoﬃi has been isolated from the
blood of healthy and diseased dogs, and from a person with endocarditis
[28,29]. B vinsonii subspecies arupensis was isolated from a cattle rancher with

Vet Clin Small Anim

33 (2003) 809–825

E-mail address:

guptillc@purdue.edu

0195-5616/03/$ - see front matter

doi:10.1016/S0195-5616(03)00024-X

fever and bacteremia [30], and B grahamii was associated with neuroretinitis
in one person [31]. Other Bartonella species isolated from woodland
mammals and for which the pathogenic potential in any species is unknown
include B alsatica, B birtlesii, B doshiae, B (Grahamella) peromysci, B
schoenbuchii

, B taylorii, and B tribocorum [32–35]. Additional species are

being characterized and will be added to the genus Bartonella in the future.

Epidemiology

Feline B henselae infection ﬁrst was reported in 1992 [36], and canine B

vinsonii

subspecies berkhoﬃi infection ﬁrst was reported in 1994 [37]. Since

then, natural infection of cats with four Bartonella species (B henselae,
B clarridgeiae

, B koehlerae, and B bovis) [11,25,23,26,38], and of dogs

with four Bartonella species (B vinsonii subspecies berkhoﬃi, B henselae, B
clarridgeiae

and B elizabethae) [5,9,24,39], has been reported. Seroepidemio-

logic studies of cats indicate exposure to Bartonella species, most frequently
B henselae

, occurs worldwide. Seroprevalence is greatest in warm, humid

climates, particularly in older cats, feral cats, and cats infested with ﬂeas
[21,40–48]. B henselae bacteremia occurs in approximately 5% to 40% of
cats in the United States, also with a higher prevalence in warmer, more
humid regions [21,18,49,44]. Approximately 90% of cats belonging to
people with CSD had B henselae bacteremia [50]. Approximately 10% of
cats with Bartonella bacteremia in the United States were infected with B
clarridgeiae.

Between 16% and 31% of cats in two studies in France and

31% of cats in the Philippines [21,18,48] with Bartonella bacteremia were
infected with B clarridgeiae. B koehlerae was isolated from two cats (from
one household in California), and B bovis was isolated from a cat in Utah
and one in Illinois [25,26]. Domestic cats are considered the major reservoir
and vector for infections in people with B henselae. Cats may be the reser-
voir for B clarridgeiae, and cattle may be the reservoir for B bovis. The
reservoir for B koehlerae is unknown.

Wild felids also are exposed to Bartonella infection. Eighteen percent of

panthers in Florida, 28% of mountain lions in Texas, and 30% to 53% of
free-ranging and captive wild felids in California had serum antibodies to
B henselae

[51,52].

Bartonella henselae

are genetically diverse. There are two recognized 16S

rRNA types of B henselae and at least two subgroups within each type [53].
Coinfection of cats with B henselae 16S rRNA types I and II and with B
henselae

and B clarridgeiae [19,54,55] is reported. There are regional

diﬀerences in prevalence of infection of cats with diﬀerent rRNA types of
B henselae

[10,48,43]. There may be genomic variation in B henselae during

the course of infection in cats [56]. Such variation may enhance the ability of
B henselae

to persist in the cat for prolonged periods.

Bartonella henselae

seroreactivity was reported in 3 (3%) of 100 dogs

whose serum was submitted for autoimmune screening in the United

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L. Guptill / Vet Clin Small Anim 33 (2003) 809–825

Kingdom [57], in 2 (6.4%) of 31 dogs in a convenience sample in Hawaii [58],
and in 4 (7.7%) of 52 dogs in a convenience sample in Japan [59]. B vinsonii
subspecies berkhoﬃi seroreactivity was reported in 69 (3.6%) of 1920 sick
dogs from North Carolina and Virginia, 4 (10%) of 40 sick dogs in Israel [60],
163 (8.7%) of 1872 healthy military working dogs in the United States [61],
68 (14%) of 483 healthy dogs in Southern France, Africa, French Guyana,
and Martinique [62], and 19 (38%) of 49 sick dogs in Thailand [63]. There
were no culture-positive blood samples in a study testing 211 dogs in the
United Kingdom [55].

Seroprevalence and bacteremia were evaluated in coyotes in California.

Three hundred eight (35%) of a convenience sample of 869 coyotes were
seropositive for B vinsonii subspecies berkhoﬃi [64]. Thirty-one (28%) of
another sample of 109 coyotes were bacteremic with B vinsonii subspecies
berkhoﬃi

and 83 (76%) of the 109 coyotes were seropositive [65]. This

information suggests that coyotes may be reservoirs for B vinsonii subspecies
berkhoﬃi

Pathogenesis

Cats

Bartonella henselae

is believed to be transmitted naturally among cats

by cat ﬂeas (Ctenocephalides felis felis). The exact role of the ﬂea in
transmission has not been determined. B henselae was transmitted among
cats by transferring ﬂeas fed on infected cats to speciﬁc pathogen free cats
and by intradermal inoculation of excrement from infected ﬂeas [66,67].
Cats fed on by Bartonella-infected ﬂeas that were enclosed in capsules that
prevented contamination of cats with ﬂea excrement did not become
infected with B henselae [67]. This suggests that transmission does not occur
by way of ﬂea saliva. Ticks also may have a role in transmission.

In laboratory studies, cats were experimentally infected with B henselae

through intravenous (IV) or intramuscular (IM) [54] inoculation with
infected cat blood, and by IV, subcutaneous, intradermal or oral inoculation
with laboratory-grown bacteria [68–70]. B henselae transmission does not
occur when infected cats cohabit with uninfected cats in a ﬂea-free environ-
ment [68,70], indicating that transmission among cats does not occur directly
through cat bites, scratches, grooming, or sharing of food dishes and litter
boxes when ﬂeas are absent. Transmission did not occur when cats were
inoculated IM with urine of bacteremic cats [71]. There was no transmission
between ﬂea-free B henselae-bacteremic female cats and uninfected males
during mating, nor was there transplacental or transmammary transmission
to kittens [68,72].

Bacteremia with B henselae and B clarridgeiae is commonly chronic and

recurrent. Experimentally infected cats kept in arthropod-free environments
maintained relapsing B henselae or B clarridgeiae bacteremia for as long as

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L. Guptill / Vet Clin Small Anim 33 (2003) 809–825

454 days [54]. Relapsing bacteremia was reported in naturally infected cats
for 3 years; however, reinfection of cats living in private homes may occur
through exposure to ﬂeas [50].

Cats appear to be protected from reinfection with homologous strains of

B henselae

, but not in all cases against heterologous challenge. Cats

previously infected with B henselae 16S rRNA type II were not protected
from infection with B henselae 16S rRNA type I [54], but they were protected
against homologous challenge. Cats infected with B henselae type I or II were
susceptible to challenge infection with B clarridgeiae, and cats infected with
B koehlerae

or B clarridgeiae were susceptible to challenge infection with

B henselae

type I or II. In contrast, cats infected with B henselae type I were

protected partially or completely against challenge infection with B henselae
type II [73].

The localization of Bartonella in cats has not been completely deter-

mined. Bartonella are intracellular bacteria, and B henselae has been de-
tected within erythrocytes of infected cats [74]. Bartonella also may be
located intracellularly in vascular endothelial cells of infected cats as has
been suggested for rodents [75]. Extracellular B henselae also are detected
in blood and other tissues of infected cats [76].

Dogs

In an epidemiologic study of dogs in Virginia and North Carolina, dogs

reported by owners to have moderate to heavy tick infestation or heavy ﬂea
exposure were 14.2 times or 5.6 times more likely, respectively, than dogs
without tick or ﬂea infestation, to be seropositive for B vinsonii subspecies
berkhoﬃi

. This suggests that ticks and ﬂeas may be vectors for transmission

of B vinsonii subspecies berkhoﬃi [77] to dogs. Questing Ixodes paciﬁcus
ticks collected in California were PCR positive for several species of
Bartonella

[55,78], further supporting the hypothesis that ticks are vectors

for Bartonella transmission.

Dogs experimentally inoculated with B vinsonii subspecies berkhoﬃi were

infected persistently for up to 247 days but did not show clinical signs of
disease other than transient fever in some dogs. There was some evidence of
immunosuppression. Dogs in two studies had temporary cyclic decreases or
mild sustained decreases in peripheral blood CD8 + lymphocyte numbers
[79,80]. There was decreased phagocytosis by monocytes of infected dogs,
but there was no diﬀerence in oxidative burst activity of the monocytes
compared with those of control dogs. Neutrophil phagocytosis was normal
[80].

Experimentally infected dogs produced serum IgG antibodies to B vinsonii

subspecies berkhoﬃi that peaked within the ﬁrst 2 to 4 weeks following
inoculation, then declined, but remained increased above a reciprocal titer of
64 through at least 184 days following inoculation in one study, and through
132 days in another study [79,80].

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L. Guptill / Vet Clin Small Anim 33 (2003) 809–825

Blood cultures in experimentally infected dogs were positive for B vinsonii

subspecies berkhoﬃi through 42 days following inoculation in eight dogs and
for as long as 76 days in one dog. Organisms were detected in peripheral
lymph nodes by PCR at up to 142 days following inoculation in one dog and
in the liver of another dog at 247 days following inoculation, suggesting that
B vinsonii

subspecies berkhoﬃi may persist in healthy dogs for prolonged

periods [79,80].

Clinical ﬁndings

Cats

Clinical signs are rare in naturally infected cats. Four cats developed fever

following elective surgical procedures [81]. One cat with uveitis was sero-
logically positive for B henselae infection with evidence of ocular production
of anti-Bartonella IgG antibodies [82]. Seven (14%) of 49 animals in
a convenience sample of cats with uveitis had evidence of ocular production
of anti-Bartonella IgG antibodies [83]. There were no clinical signs reported in
65 naturally infected cats in another study [84]. It is not known if argyrophilic
bacteria described in lymph nodes of cats with persistent lymphadenomegaly
[85] were Bartonella or if peliosis hepatis of cats [86] was caused by Bartonella.

Clinical signs in cats experimentally infected with B henselae are mild,

transient, and vary with the strain of B henselae used for inoculation
[54,70,87]. Cats inoculated intradermally developed areas of induration or
abscess at inoculation sites between approximately 2 and 21 days after
inoculation [54,70,71,87–89]. Pure cultures of B henselae were obtained from
these lesions in some cats [70]. Other transient clinical ﬁndings included
generalized or localized peripheral lymphadenomegaly (lasting for about
6 weeks following inoculation), short periods of fever (greater than 103

39.4

C) during the ﬁrst 48 to 96 hours following inoculation and again for

24 to 48 hours at approximately 2 weeks following inoculation. Some cats
were lethargic and anorexic when febrile. Mild neurologic signs (nystagmus,
whole body tremors, focal motor seizures, either decreased, or exaggerated
responses to external stimuli, behavior changes), and epaxial muscle pain
also were reported [54,70,71,87,88]. Reproductive failure occurred in some
cats experimentally infected with B henselae [72]. Cats experimentally
infected with B koehlerae exhibited no clinical signs [90].

Most experimentally infected cats had no abnormalities on complete

blood counts, serum biochemical tests, or urine analysis [54,70,71,87,88].
A few had transient anemia early in the course of infection, and some
had persistent eosinophilia [54]. Mature neutrophilia occurred in some cats
during periods of skin inﬂammation [70]. Cats had hyperplasia of lymphoid
organs, small foci of lymphocytic, pyogranulomatous, or neutrophilic in-
ﬂammation in multiple tissues (lung, liver, spleen, kidney, and heart), and
small foci of necrosis in the liver [54,70].

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L. Guptill / Vet Clin Small Anim 33 (2003) 809–825

Because bacteremia can persist for long periods in cats, a potential

causative role for Bartonella species in chronic diseases of cats has been
proposed. One study suggested that cats seropositive for B henselae and
feline immunodeﬁciency virus (FIV) were more likely to have gingivitis or
lymphadenomegaly than cats seropositive for either infection alone [91].
Results of another study suggested possible associations between B henselae
seropositivity and stomatitis or various urinary tract disorders [92].

No contribution of Bartonella infections to development of chronic

illnesses in domestic cats has been veriﬁed.

Because of the high prevalence of Bartonella infection in domestic cats,

substantiation of a causative association between any disease or syndrome
and Bartonella infection will require large, carefully controlled epidemio-
logic studies.

Dogs

Dogs experimentally infected with B vinsonii subspecies berkhoﬃi

exhibited no clinical signs except for transient fever in some dogs [79,80].
Many dogs may be infected naturally with B vinsonii subspecies berkhoﬃi
without clinical signs; in two surveys, 8.7% and 14% of healthy dogs were
seropositive [61,62]. Bartonella species have been detected in tissues of sick
dogs with a variety of clinical presentations, however. B vinsonii subspe-
cies berkhoﬃi has been associated with endocarditis, cardiac arrhythmias,
granulomatous rhinitis, and granulomatous lymphadenopathy in dogs
[39,93,94]. B henselae was detected using PCR in the liver of a dog with
peliosis hepatis [9], the liver of another dog with granulomatous hepatitis
[12], in the blood of a dog with septic peritonitis following penetration of the
intestinal wall by a foreign body [10], and in the blood of two dogs with
systemic disease that had been treated with corticosteroids [10]. B elizabethae
was detected using PCR in the blood of a moribund dog with chronic renal
failure [10], and B clarridgeiae was detected in the liver of a dog with
hepatopathy [12] and in the heart valve of a dog with endocarditis [24].
B vinsonii

subspecies berkhoﬃi has been detected in heart valve tissue of

multiple dogs with endocarditis and appears to be an important cause of
culture-negative endocarditis of dogs. Whether other species of Bartonella
are signiﬁcant opportunistic pathogens of dogs is not clear.

Diagnosis

Diagnosis of Bartonella infection is not straightforward. Clinical signs are

transient and variable; therefore determining which sick animals are likely to
have Bartonella infection is diﬃcult. In addition to the need to test sick pets,
veterinarians may be asked to test healthy pets belonging to clients with
Bartonella

-related illnesses, or to screen healthy cats that are being con-

sidered as pets for immunocompromised people.

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L. Guptill / Vet Clin Small Anim 33 (2003) 809–825

The use of serologic testing alone as a diagnostic tool is problematic, in

that there are not clear criteria for establishing a deﬁnitive diagnosis through
serology. Five percent to 12% of cats with B henselae bacteremia are
seronegative [46,84]. Serum IgG antibodies persist in experimentally infected
cats and dogs for prolonged periods, and how long antibodies persist fol-
lowing clearance of an infection is unknown.

Immunoﬂuorescent antibody (IFA) and enzyme immunoassay (EIA)

tests are available for serologic testing. Infections with some strains or
species of Bartonella may be missed using either method, depending on the
antigen preparations used [95]. The positive predictive value of a positive
serologic test for bacteremia in cats is low. Positive predictive values of IFA
and EIA tests for anti-B henselae serum IgG in cats are 39% to 46%. The
utility of a negative serologic test is greater, as the negative predictive values
for these tests in cats are high, at 89% to 97% [20,43,46,84]. Therefore,
a negative serologic test may be a useful tool for screening cats as potential
pets. Predictive values for serologic tests for dogs have not been reported.
For coyotes, the positive predictive value of a positive IFA test for bacter-
emia was 29%, and the negative predictive value was 73% [65].

The use of Western blot tests has been advocated for serodiagnosis of

feline B henselae infections, but the diagnostic accuracy of Western blot tests
awaits further investigation. In human medicine, Western blot testing re-
mains problematic, in that serologic responses of people as evaluated by
immunoblot vary among patients [96].

Blood culture is indicated for sick pets with positive serologic test results

whose history and clinical presentation suggest possible Bartonella infection.
A positive blood culture or culture of other tissue is the most reliable test for
deﬁnitive diagnosis of active Bartonella infection. Because of the relapsing
nature of feline Bartonella bacteremia, a single blood culture is not a sensitive
diagnostic tool for bacteremia, and multiple blood cultures may be necessary
[55]. Blood culture is considered insensitive for detecting canine Bartonella
infection; most dogs with Bartonella endocarditis are blood culture negative,
and experimentally infected dogs have relatively short periods of bacteremia.

Blood for culture should be obtained using sterile technique, and the

blood should be placed in ethylenediamine tetra-acetic acid (EDTA)-
containing tubes or lysis centrifugation blood culture tubes (Isolator tubes,
Wampole, Cranbury, NJ). If blood is collected into EDTA tubes, it should be
frozen, and ideally, plastic EDTA tubes used. Blood should be sent to labo-
ratories familiar with these fastidious organisms, and laboratories should be
contacted for speciﬁc instructions for sample collection and submission.

Standard PCR testing for Bartonella DNA in blood may be no more

sensitive than blood culture for detection of active Bartonella infection.
Nested PCR testing may increase sensitivity over standard PCR for detecting
Bartonella

DNA in cat blood [97] and may be available through some

research laboratories. An advantage of PCR testing is that the results are
often available more quickly than those of blood culture. Another beneﬁt is

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L. Guptill / Vet Clin Small Anim 33 (2003) 809–825

that the DNA products of PCR may be sequenced, and the species or strain
of Bartonella therefore identiﬁed. Detecting DNA does not always equate
to detection of living organisms, however. Samples for PCR testing should
be obtained using sterile technique, and individual laboratories should be
contacted for collection and submission guidelines.

Pets may be coinfected with multiple pathogens. For example, dogs with

a variety of clinical signs from a North Carolina kennel were found (using
PCR) to be coinfected with from two to six tick-transmitted pathogens.
Pathogens found coinfecting some of the dogs in addition to B vinsonii
subspecies berkhoﬃi included Ehrlichia canis, E chaﬀeensis, E equi (now
Anaplasma phagocytophila

), E (now Anaplasma) platys, Rickettsia species, or

Babesia canis

[98]. In another study, cats that were seropositive for B henselae

and FIV reportedly had more severe gingivitis than cats infected with FIV
alone [91]. Such coinfections make attributing clinical signs of disease to
infection with a particular organism diﬃcult, and also have important
implications for therapy.

Treatment

People with Bartonella infections causing bacillary angiomatosis-peliosis,

or endocarditis are treated with a variety of antibiotics, including doxy-
cycline, erythromycin, ciproﬂoxacin, rifampin, gentamicin, trimethoprim-
sulfamethoxazole, clarithromycin, and azithromycin [99,100]. Antibiotic
treatment is eﬀective for people with bacillary angiomatosis, but not for
people with cat scratch disease or endocarditis. Most people with Bartonella
endocarditis require valve replacement.

Cats

Documenting clearance of feline Bartonella infections through antibiotic

treatment is diﬃcult because of the prolonged nature of the infections and the
relapsing bacteremia. Treatment of Bartonella infections appears to require
long-term (at least 4 to 6 weeks) antibiotic administration. No regimen of
antibiotic treatment has been proven eﬀective for deﬁnitively eliminating
Bartonella

infections in cats [18,89,101]. Enroﬂoxacin (5.4 to 7.6 mg/kg by

mouth every 12 hours) treatment for 28 days appeared to clear B henselae or
B clarridgeiae

infection in ﬁve of six treated cats that were followed for

12 weeks after treatment [18]. Recent studies, however, show that enroﬂoxa-
cin causes retinal degeneration and blindness in some cats, and use of more
than 2.5 mg/kg every 12 hours is contraindicated [102]. Doxycycline (6.9 to
12.8 mg/kg by mouth every 12 hours) appeared to clear B henselae or B
clarridgeiae

infection in one of six cats treated for 2 weeks. Doxycycline (4 to

10.4 mg/kg by mouth every 12 hours) for 4 weeks appeared to clear infection
in one of two cats [18]. Antibiotics tested in other studies (erythromycin,
amoxicillin, amoxicillin-clavulanic acid, and tetracycline) decreased the level

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L. Guptill / Vet Clin Small Anim 33 (2003) 809–825

of bacteremia in treated cats. The cats, however, either were not followed for
a prolonged period after treatment [89], or the treatment could not be
deemed successful compared with no treatment, because untreated cats
became blood culture negative after the same length of time as did cats that
were treated [101]. Rifampin used alone or in combination with doxycycline
has been recommended [103]. Azithromycin has been recommended for
treatment of infected cats, but data from controlled eﬃcacy studies with
long-term follow-up are lacking.

Because of the uncertainty of antibiotic eﬃcacy and the concern that

routine treatment for asymptomatic bartonellosis may induce resistant
strains, treatment can be recommended only for animals showing clinical
signs of disease. In some cases, treatment of a healthy pet may be rec-
ommended if it is the only alternative to euthanasia. Although treatment
decreases the level of bacteremia in cats, there is no concrete evidence that
treatment of the cat will decrease the probability of transmission of
Bartonella

infection to an owner.

Dogs

Dogs with clinical signs of Bartonella infection and positive serologic test

results should be treated with antibiotics. Information regarding treatment
of dogs for Bartonella infection is limited, but there is a consensus that
treatment should be prolonged, at least 4 to 6 weeks [104]. Macrolide
antibiotics may be the best option, based on results reported in the hu-
man literature. Azithromycin has been used to treat dogs, as have several
other antibiotics at varying doses, including doxycycline and enroﬂoxacin
[10,12,39,94,104]. There are no published controlled studies evaluating the
eﬃcacy of any antibiotic regimen for treatment of dogs.

Therapeutic monitoring

The possibility of coinfection with multiple pathogens should be inves-

tigated in all animals suspected of Bartonella infection, and the animals
should be evaluated and treated accordingly.

When animals are treated with antibiotics, several follow-up cultures

performed at 4- to 8-week intervals should be obtained from treated animals
to try to assess eﬃcacy [18,89,101]. Client education regarding the uncer-
tainty of antibiotic eﬃcacy, the importance of prolonged follow-up after
antibiotic treatment, and the importance of vector control and other means
of preventing transmission is essential.

Prevention

Cats

Prevention of Bartonella infections in cats is accomplished best by

avoiding exposure to infected animals and their ﬂeas. Because B henselae

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L. Guptill / Vet Clin Small Anim 33 (2003) 809–825

and B clarridgeiae have been transmitted through inoculation of infected cat
blood [71], cats that are seropositive for Bartonella should not be used as
blood donors. Development of a vaccine to prevent Bartonella infection in
cats has not been successful.

Dogs

Avoiding infestation with ticks, ﬂeas or other arthropod vectors is the

best strategy for preventing Bartonella infection in dogs. Although dogs do
not appear to develop chronic bacteremias as do cats, the use of seropositive
dogs as blood donors should be avoided.

Public health

Bartonella

species cause many clinical syndromes in people. including

CSD (typical and atypical forms, including encephalopathies in children),
bacillary angiomatosis, parenchymal bacillary peliosis, relapsing fever with
bacteremia, endocarditis, optic neuritis, pulmonary, hepatic, or splenic gran-
ulomas, and osteomyelitis [7,14,17,28,30,99,105,106]. Immunocompetent
individuals tend to have more localized infections, whereas infections that
occur in immunocompromised individuals are more often systemic and can
be fatal. Veterinarians, veterinary staﬀ, groomers, and others with extensive
companion animal contact are at a greater risk for Bartonella infection than
members of the general public [107,108].

Transmission of B henselae from cats to people probably occurs through

contamination of cat scratches with ﬂea excrement. Transmission may occur
through cat bites if cat blood or ﬂea excrement contaminate the bite site [66].
Ticks are believed to be vectors for transmission of some Bartonella infections
to dogs. The role of arthropod vectors in transmission of Bartonella infec-
tions to people remains undeﬁned, but people should take precautions
against ﬂea and tick exposure. Some people with Bartonella infections have
reported exposure to dogs and not cats [59,109], and others report no animal
contact at all.

The US Public Health Service/Infectious Diseases Society of America

(USPHS/IDSA) Guidelines for Preventing Opportunistic Infections Among
HIV-Infected Persons [110] recommend the following when acquiring a new
cat: adopt a cat over 1 year of age that is in good health, avoid rough play
with cats, maintain ﬂea control, wash any cat-associated wounds promptly,
and do not allow a cat to lick wounds or cuts. There is no evidence that
declawing cats decreases the probability of transmission of B henselae from
cats to people. The USPHS/IDSA Guidelines state that there is no evidence
of any beneﬁt to cats or their owners from routine culture or serologic testing
of cats for Bartonella infections. Because the negative predictive value of B
henselae

serology for feline bacteremia is very good, however, serology may

818

L. Guptill / Vet Clin Small Anim 33 (2003) 809–825

be an appropriate screening test for cats that immunocompromised persons
are considering acquiring as pets.

Summary

The role of Bartonella species as pathogens in dogs and cats is being

deﬁned. Diagnosis and treatment of Bartonella infections of dogs and cats
remain challenging. As new information regarding Bartonella infections of
companion animals becomes available, the understanding of the pathogen-
esis of these infections will improve.

Most Bartonella species infecting dogs and cats are zoonotic, with B

henselae

the most important zoonotic species. B henselae bacteremia is

common in domestic cats, and cats transmit B henselae to people. Trans-
mission of Bartonella infections among cats and dogs is believed to occur
primarily by way of arthropod vectors. Control of arthropod vectors and
avoiding interactions with pets that result in scratches or bites are the most
eﬀective means to prevent transmission between animals and people.

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[100] Bass JW, Freitas BC, Freitas AD, Sisler CL, Chan DS, Vincent JM, et al. Prospective

randomized double-blind, placebo-controlled evaluation of azithromycin for treatment of
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[101] Regnery RL, Rooney JA, Johnson AM, Nesby SL, Manzewitsch P, Beaver K, et al.

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[102] Wiebe V. Fluoroquinolone-induced retinal degeneration in cats. J Am Vet Med Assoc

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the dog and cat. Philadelphia: WB Saunders; 1998. p. 337–43.

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[107] Noah DL, Kramer CM, Verbsky MP, Rooney JA, Smith KA, Childs JE. Survey of

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[108] Kumasaka K, Arashima Y, Yanai M, Hosokawa N, Kawano K. Survey of veterinary

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825

L. Guptill / Vet Clin Small Anim 33 (2003) 809–825

Canine borreliosis

Meryl P. Littman, VMD

University of Pennsylvania School of Veterinary Medicine, Department of Clinical

Studies–Philadelphia, 3900 Delancey Street, Philadelphia, PA 19104-6010

Borreliosis is not a new disease. The DNA footprints of Borrelia

burgdorferi

are in museum specimens of ticks from New England mice of

1894 [1]. Earlier, when colonists came to North America, forests were
cleared for farms, thereby decreasing the deer population in the area. As
farms were abandoned, they were replaced by suburbia, including its wild
vegetation and private wooded areas, which encouraged increased density of
deer. Migratory birds helped bring more ticks and their organisms back
from the woods and the coastal islands’ nature preserves.

The ﬁrst documented case of borreliosis in North America was in 1969,

when a physician in Spooner, WI, went grouse hunting, got bitten by a tick,
and later had a rash. But the disease was not named ‘‘Spooner’’ disease [2].
Instead the disease was named for a Connecticut neighborhood with
a cluster of children who were diagnosed with juvenile rheumatoid arthritis,
where two assertive mothers, Judith Mensch and Polly Murray [3],
complained to health authorities in 1975. Allen Steere, a young Yale
rheumatologist with Centers for Disease Control and Prevention (CDC)
training, interviewed patients (39 children, 12 adults) previously diagnosed
with rheumatoid arthritis, ringworm, multiple sclerosis, lupus, and other
ailments. Only a few remembered a rash, thought to be from an insect bite,
weeks to months before other signs. First thought to be virally caused, the
disease got more mysterious each year. Within days of a tick bite, many had
an expanding rash at the site with an inﬂuenza-like illness and severe
headache. Weeks later, some people had arthritis or neurologic or cardiac
signs, and weeks to months later, intermittent, sometimes shifting mono-
oligoarthritis occurred, most often involving the knee (Table 1). The rash
resembled the expanding bull’s eye rash called ‘‘erythema chronicum
migrans’’ (ECM) associated with Ixodid tick bites, ﬁrst described in Europe
in 1909 in a patient who later developed neurologic signs [9]. The European

Vet Clin Small Anim

33 (2003) 827–862

E-mail address:

merylitt@vet.upenn.edu

0195-5616/03/$ - see front matter

doi:10.1016/S0195-5616(03)00037-8

ble

ompariso

huma

and

canine

Nort

erican

borre

liosis

Huma

[4–6]

Dogs

[1,7,8

]

linical

sign

Asym

ptoma

tic

infections

10%

asymp

tom

atic

[

95%

asymp

tomat

rly,

localized

(with

day

80%

untrea

ted

have

ECM

rash

Common:

infuenza

-like

signs:

feve

head

ache,

myalgia,

arthralgia

regi

onal

lym

phad

enopath

signs.

rly,

dissem

inated

(un

treated)

eeks

after

exposur

Com

mon:

mono-

olig

oarthrit

15%

have

neuro

logic

sign

(eg,

facia

palsy,

lymphoc

ytic

meningiti

radiculon

europath

ies).

have

cardiac

sign

(eg,

A–V

block

Multip

rashes.

signs

gener

ally.

Unkn

own

whe

‘‘Lyme

phro

pathy’’

rare

cardiac

neurolo

gic

signs

mig

occur.

Rare

hepa

tic,

ocula

signs.

dissem

inated

(untr

eated)

wee

months

ter

expo

sure

60%

have

inte

rmitte

attacks

no-arth

ritis

mig

rator

polyar

thropa

thy.

Mon

lyarthro

pathy

feve

anorexia.

have

ronic

neurolo

gic

man

ifestatio

ns.

Intermittent

chronic

Chro

nic

axo

nal

polyn

europat

hy,

can

mimic

multiple

sclerosis.

Cogn

itive

disturb

ances

Chro

nic

mild

encephalo

pathy.

Possible

shiftin

leg

lameness.

Self-limiting

exper

imental

beagle

studies,

onl

seen

you

pupp

ies.

‘‘Ly

neph

ropath

y’’

needs

udy.

Incide

nce

unkno

wn.

Time

aft

exp

osure

unkno

wn.

animal

model.

Pat

hogene

sis

may

cause

persist

ence

infection

som

patie

nts

and

immunolog

mechan

isms

oth

ers.

May

invo

lve

oth

agents

such

Ehrlich

spp,

besia

spp,

Bartone

lla

spp,

ckettsia

spp

Lepto

spirosis.

Rare

cardiac

urolo

gic

involve

ment.

828

M.P. Littman / Vet Clin Small Anim 33 (2003) 827–862

Jarisch

–Herxhe

imer

reaction

10%

15%

hav

transien

wors

ening

signs

soon

aft

arting

treatmen

because

inc

reased

cyto

kine

release

Not

rep

orted.

Treatm

ent-r

esista

chronic

Lyme

arthr

itis

10%,

associa

ted

ith

HLA-D

and

molec

ular

mim

icry

Osp

with

LFA-

Unkno

wn,

needs

study.

Recom

mended

treatmen

Doxyc

ycline

(100

2–3

wks)

Amoxicillin

(500

2–3

wks)

Eryt

hromycin

(250

2–3

wks)

Neurolo

gic

chronic

cases:

Ceft

riaxon

(2g

2–4

wks)

Doxyc

ycline

(10

mg/kg

12–2

2–4

wks).

Amoxicillin

(20

mg/kg

8–12

2–4

wks).

Azithro

mycin

(25

mg/k

2–4

wks).

Ceftriaxone

arely

use

clinically)

mg/

2–4

wks.

‘‘Lyme

nephro

pathy’’

treated

with

antib

iotics

lon

ger

than

others

(if

surv

ives);

ACE

inhibiter,

low-dose

asp

irin,

lloids,

treatm

ent

for

renal

failur

Avai

lable

vac

cines

LYM

Erix

(mar

keted

Dece

mber

1998

bruary

2002

)

SmithK

line

Beecha

Biolo

gicals

Philad

elphia

longer

available

Lyme

Vax—ba

cter

in,

novalen

Dodg

Animal

Healt

Ove

rland

Park

KS.

Galaxy

me—bac

terin

bivalen

Sche

ring-

Plough

Animal

Healt

Kenilworth

NJ.

Recomb

itek—su

bunit

recom

binant

OspA,

nona

djuvan

ted,

Meria

Limited

Atlan

ta,

ProLym

e—subu

nit

Osp

adjuva

nted

Inte

rvet,

Millsboro,

DE.

Abbre

viatio

ns:

ECM,

eryth

ema

chroni

cum

migran

IV,

intra

veno

us.

829

M.P. Littman / Vet Clin Small Anim 33 (2003) 827–862

rash responded to penicillin, but the organism responsible was unidentiﬁed.
Serendipitously, in 1981 Willy Burgdorfer, who trained earlier in Europe
and knew about ECM rashes, was examining Shelter Island ticks for
Rickettsia rickettsia

. He found instead that 50% of the deer ticks were

infected with odd, spiral-shaped bacteria, which he named B burgdorferi
[10]. Serum samples from patients with Lyme disease showed high antibody
titers directed against the spirochetes grown in culture, and animal models
proved Koch’s postulates. Serologic tests were developed; clinical signs were
described; outer surface proteins were characterized; the DNA of B
burgdorferi

was sequenced, and vaccines were produced.

In the United States, more than 132,000 cases of Lyme disease have been

reported since 1982, with more than 17,000 reported in 2000 (Fig. 1).
Reporting is hampered by under-reporting and overdiagnosis. Roughly 89%
of the cases were reported from the northeastern and Mid-Atlantic States, 7%
from the upper Midwest, and 1% from the West Coast (Fig. 2). The other 3%
were cases from all other states combined and probably represent infection
picked up by people who had visited endemic areas. Twelve states account for
95% of the cases. They are, in order of incidence, Connecticut, Rhode Island,
New Jersey, New York, Delaware, Pennsylvania, Massachusetts, Maryland,
Wisconsin, Minnesota, New Hampshire, and Vermont.

On May 22, 1989, Newsweek’s cover showed an Ixodes tick, magniﬁed 98

times, and stated ‘‘A tiny tick is spreading a mysterious illness in 43 states.’’
Such media attention fed a growing panic, and Lyme disease became known
as ‘‘the great mimicker.’’ Controversy ensued about overdiagnosis and

Fig. 1. Annual reports of Lyme disease, 1982 to 2002. (Data from the Centers for Disease
Control and Prevention. Available at www.cdc.gov/mmwr/preview/mmwrhtml/mm5102a3.htm.)

830

M.P. Littman / Vet Clin Small Anim 33 (2003) 827–862

overtreatment of this trendy disease. Lyme disease was deemed a most
important infectious disease in North America, second only to HIV/AIDS.
Internet search engines such as Google yield more than 200,000 sites
concerning Lyme disease, many of which are anecdotal, poorly sub-
stantiated accounts (only 55 sites were found helpful to the research
community) [11]. People in Lyme-endemic areas naturally are concerned
about Lyme disease in their loved ones, including their dogs, and
veterinarians play an important role in educating owners about the disease
and public health.

Much has been learned about the life cycle of the vector tick and the

causative organism, including its full DNA sequence. A new serologic test
with high sensitivity and speciﬁcity has helped to recognize those exposed
and infected, but the clinician may not be certain that clinical signs are
caused by that infection. There is still much speculation and controversy
about the interpretation of laboratory test results, about which individuals
need to be treated, how they should be treated, and how prevention is best
accomplished. Although clinicians feel more conﬁdent about which animals
are exposed and infected, the diagnosis of Lyme disease remains
problematic. The diagnosis of Lyme disease has relied upon a combination
of (1) exposure to the organism and Ixodid ticks in geographic areas
endemic for borreliosis, (2) clinical signs of illness that have been associated
with borreliosis, (3) ﬁnding the organism or antibody directed against the
organism, (4) diagnostic work-up ruling out other diﬀerentials, and (5)
response to therapy.

Fig. 2. National Lyme disease risk map with four categories of risk. (Data from the Centers for
Disease Control and Prevention. Available at www.cdc.gov/ncidod/dvbid/lyme/index.htm.)

831

M.P. Littman / Vet Clin Small Anim 33 (2003) 827–862

This article discusses these ﬁve criteria and what concerns we still have.

The diagnosis of Lyme disease is not as easy as it looks. Diagnosis is
confounded by the realization that the clinical signs from Anaplasma
phagocytophila

are similar, and that in areas where seroprevalence of Lyme

disease is high, seroprevalence of A phagocytophila is also relatively high.
Other agents in these areas, including Ehrlichia, Rickettsia, Bartonella,
Babesia

, and even Leptospira species add to the complexity. Clinicians might

not be sure which infection, if any, is causing clinical signs, or whether
coinfection increases the likelihood of disease, and they often settle for
a diagnosis of ‘‘a doxycycline-responsive disease.’’

Exposure: the tick and the organism

Seroprevalence studies in various hosts help plot the risks in endemic

areas and note changes over the years (Table 2). In New England, for
instance, 54% of the Ixodid ticks tested positive for B burgdorferi. Even
within endemic states, tick prevalence and percentage of tick infectivity can
be quite focal, and some counties are much more aﬀected than others. In
south coastal Maine, 100% of the dogs were seropositive for Lyme within
a 0.8 km area studied, but only 2% were seropositive 1.5 km beyond there
[30]. Dogs living in endemic areas can be sentinels for focal areas of high
exposure. Exposure to B burgdorferi is mostly dependent on exposure to
Ixodes

ticks (aka, deer ticks, black-legged ticks, or bear ticks). These hard

ticks have a recognizable U-shaped groove just anterior to the anus. Their
mouth parts extend outward. A dorsal hard plate (scutum) covers the entire
back of the adult male, but only part of the female’s back. Ixodes scapularis,
once called I dammini, is the main vector in the East and Midwest; I paciﬁcus
is in the West. Since lizard, with a blood-borne borreliacidal factor [31], is
a preferred host of I paciﬁcus nymphs, Lyme disease is not as prevalent
along the West Coast as in the East or Midwest, where I scapularis nymphs
feed on small mammals and birds.

Deer ticks are small hard three-host ticks (Fig. 3) that feed on each host

for 3 to 5 days, drop oﬀ to molt, and ﬁnd the next host by questing on
vegetation. A fascinating 2-year life cycle and trans-stadial transmission
allow the ticks to pick up B burgdorferi from recently infected hosts. The six-
legged larvae hatch and feed in the summer; they molt into eight-legged
nymphs who feed in the spring, and they molt to adults who breed and feed
on their hosts in the fall. Larvae and nymphs feed on small mammals
(especially white-footed mice in the East and Midwest) and migratory birds
such as robins [32,33], and nymphs and adults feed on white-tailed deer and
other large mammals, such as dogs and people. Because larvae often feed on
the same kinds of small animals and birds as nymphs do, the noninfected
larvae pick up B burgdorferi in the summer from a host that recently was
infected by the bite of an infected nymph in the past spring.

832

M.P. Littman / Vet Clin Small Anim 33 (2003) 827–862

Table 2
Prevalence of agents carried by Ixodes ticks in various species and geographic areas

Agent sought

Host

State

% positive

Reference

Borrelia

burgdorferi

Dogs, healthy

NY, CT

89.6%

Magnarelli 1990 [12]

‘‘

Dogs, lame

NY, CT

92.9%

Magnarelli 1990 [12]

‘‘

Dogs

52%

Hinrichsen 2001 [13]

‘‘

40%

Guerra 2001 [14]

‘‘

Horses

Northeast US

45.1%

Magnarelli 2000 [15]

‘‘

Mice

83.3%

Staﬀord 1999 [16]

‘‘

Ixodes

ticks

Russia

38%

Morozova 2002 [17]

‘‘

54%

Chang 1998 [18]

Borrelia

spp

‘‘

The Netherlands

13%

Schouls 1999 [19]

Bb + Ap

Deer

49%

Magnarelli 1999 [20]

Bb + Ap

Ixodes

ticks

Chang 1998 [18]

Dogs

14.4%

Hinrichsen 2001 [13]

‘‘

Northern CA

9% to 47%

Foley 2001 [21]

‘‘

Dogs,

Lyme negative

CT, NY

0% in 1985

Levy 2002 [22]

‘‘

6.6% in 2001

Levy 2002 [22]

‘‘

Dogs, Lyme +

CT, NY

15.6% in 1985

Levy 2002 [22]

‘‘

40% in 2001

Levy 2002 [22]

‘‘

Humans,

healthy

14.9%

Bakken 1998 [23]

‘‘

Humans,

blood donors

11.3%

Aguero 2002 [24]

‘‘

Humans,

Lyme +

35.6%

Aguero 2002 [24]

‘‘

20%

Ravyn 1998 [25]

‘‘

Horses

MN, WI

17.6%

Bullock 2000 [26]

‘‘

Northeast US

15.9%

Magnarelli 2000 [15]

‘‘

Deer

60%

Walls 1998 [27]

‘‘

25%

Walls 1998 [27]

‘‘

Wood rats

Northern CA

37.6%

Castro 2001 [28]

‘‘

Mice

Northern CA

8.5%

Castro 2001 [28]

‘‘

23.5%

Staﬀord 1999 [16]

‘‘

Ixodes

ticks

Chang 1998 [18]

‘‘

25%

Chang 1998 [18]

‘‘

Russia

Morozova 2002 [17]

Ehrlichia

spp.

‘‘

The Netherlands

45%

Schouls 1999 [19]

Babesia microti

Mice

76.9%

Staﬀord 1999 [16]

Bartonella

spp.

Ixodes

ticks

The Netherlands

70%

Schouls 1999 [19]

‘‘

Present near

patients

Eskow 2001 [29]

TBE virus

‘‘

Russia

46%

Morozova 2002 [17]

Abbreviations:

Bb, Borrelia burgdorferi; Ap, Anaplasma phagocytophila (aka Ehrlichia

phagocytophila

, agent of HGE, E equi); TBE, tick-borne encephalitis virus, a variant of

Powassan virus.

833

M.P. Littman / Vet Clin Small Anim 33 (2003) 827–862

Borrelia burgdorferi

are helical-shaped unicellular spirochetes, sized 10–

0.18–0.25 lm, with ﬂagellar projections [10]. They live in the tick’s mid-

gut and are transferred along with the blood meal mostly between 48 to 72
hours of attachment to the host [34]. These gram-negative bacteria require
special silver stains, Giemsa, or acridine orange and dark-ﬁeld microscopy
to visualize in wet preparations. They are microaerophilic and have special
culture requirements. They are not freely living organisms like their cousin,
Leptospira

, and they quickly die outside the body. The organism may be

transmitted by way of blood products [35], urine [36], milk [37], or
transplacentally [1]. In the dog, B burgdorferi spreads mainly by tissue
migration [38] rather than hematogenously. The organism is associated
especially with collagen and connective tissue. Although positive cultures or
polymerase chain reaction (PCR) testing may be obtained sometimes from
urine, control dogs housed with carrier dogs for 1 year showed no
seroconversion [39].

Lyme disease is caused by organisms of the B burgdorferi senso lato species,

which includes four subgroups: B burgdorferi senso stricto (in North America
and Europe), B garinii (Eurasia), B afzelii (Eurasia), and B japonica (Japan).
The predominant North American strain of B burgdorferi senso stricto is
strain B31. Other borrelias exist, some of which appear to be nonpathogenic,
and some which cause relapsing fever. A distinct Borrelia isolate from Florida
dogs, whose sera showed cross-reactivity with Lyme antigens, shows that
inaccurate diagnosis of Lyme disease could occur at least in that area [1]. B
lonestari

, transmitted by lone star ticks (Amblylomma americanum) in the

southeastern and south-central states causes Southern Tick-Associated Rash
Illness (STARI), similar to the rash of Lyme disease, but patients’ sera do not
cross-react with B burgdorferi antigens (www.cdc.gov).

The DNA of B burgdorferi B31 has been sequenced (www.tigr.org/

tigr-scripts/CMR2/GenomePage3.spl?database

¼ gbb) including 1.44 10

Fig. 3. From left to right: the deer tick (Ixodes scapularis) adult female, adult male, nymph, and
larva on a centimeter scale. (Data from the Centers for Disease Control and Prevention.
Available at www.cdc.gov/ncidod/dvbid/lyme/index.htm.)

834

M.P. Littman / Vet Clin Small Anim 33 (2003) 827–862

base pairs (bp), or about 1700 genes. The functions of the genes are being
investigated. There is a linear chromosome and 21 plasmids (linear and
circular) that encode many outer surface proteins (Osps). The plasmids of B
burgdorferi

are diﬀerent than those of other bacteria, in that these hold

essential elements of the B burgdorferi genome. Similar to Mycoplasma, the
genes do not encode biosynthetic pathways, but mostly are for transport and
binding proteins. B burgdorferi binds to glycosaminoglycans (GAGs, such as
heparin, heparan sulfate, and dermatan sulfate), to platelets and endothelial
cells by means of integrins, to collagen-associated extracellular matrix by
means of decorins, and to ﬁbrocytes by residing in deep invaginations on the
cell surface [40]. The genome includes ‘‘runaway paralogy’’ (duplications), 161
gene families with up to 41 members (probably important for immune
evasion), and a high number of pseudogenes [41].

Many genes encode various surface proteins (there are at least 150

diﬀerent proteins) [42]. Some have been identiﬁed and characterized. OspA
appears to be important for attachment of B burgdorferi to the tick mid-gut,
and is a predominant surface lipoprotein in the unfed tick and culture in
vitro. During tick feeding, temperature and pH changes occurring with the
blood meal induce down-regulation of OspA expression and up-regulation
of expression of OspC, which is not expressed when B burgdorferi is in the
tick or in vitro culture. After about 2 days of tick feeding, a heterogenous
group of B burgdorferi within the feeding tick exists, including OspA

OspC

, OspA/OspC+, OspA+/OspC, and OspA+/OspC+ variants.

Organisms most successful in entering the host are those with less expression
of OspA. The time until successful transmission of B burgdorferi from the
tick to the mouse host was found to be exactly between 52 to 53 hours of
attachment [43], which probably is related to the time it takes for down-
regulation of OspA. A complicated up-regulation involving 116 lipoproteins
occurs in the ﬁrst 10 days of infection in the mouse, but most are then down-
regulated (perhaps until later) between days 17 and 30 [44]. These changes
appear to allow B burgdorferi to proceed to chronic infection. Within 17
days of infection in the mouse, as anti-OspC antibodies emerge, B
burgdorferi

that are not expressing OspC are selected, and the antigenic

variation game continues. Animals with anti-OspC antibody will eliminate
OspC-expressing B burgdorferi but will not clear infection. This is a
mechanism of immune evasion [45]. Culture of B burgdorferi from these
mice shows that they can express OspC again.

The Erp family (OspEF-related proteins) includes OspE, expressed in

early infection, and OspF, expressed in later infections [42]. The Erp surface
proteins bind complement inhibitor factor H, and help with the persistence
of infection [46]. The organism is able to change parts of its ‘‘coat’’ many
times. The diversity of antigens seen is found interspecies (among the B
burgdorferi senso lato

group) and intraspecies, depending on the isolate. The

changes in gene expression of some surface proteins is planned and expected
in response to the tick’s blood meal. Later, in response to the host’s immune

835

M.P. Littman / Vet Clin Small Anim 33 (2003) 827–862

system, random changes may occur as a result of the spontaneous mutations
and recombinations among expressed and silent cassettes in the gene
families. There appears to be recombination between OspA and OspB genes,
and resulting deletions and chimeric fusions of gene segments cause diversity
in a decorin binding protein A (DbpA) [42]. Many other Osps likewise are
changed, and the antigens presented to the host, especially in early infection,
are diﬀerent than those expressed by B burgdorferi within the tick or from in
vitro cultures. Thus, antibodies react to diﬀerent antigens in earlier infection
compared with later infection, when the immune system will have cleared
those B burgdorferi expressing antigens it has seen before.

An important gene allowing for antigenic variation in early disseminated

infection (but not in the tick or in vitro [47–49]) is the VlsE site, named
‘‘variable major protein-like sequence, expressed’’ because of its similarity to
the vmp (variable major protein) gene of B hermsii, agent of relapsing fever
[48]. The VlsE site, along with 15 additional silent vls cassettes (partial gene
fragments), was found on one of the linear plasmids (lp28-1). In response to
the host’s inﬂammatory mediator, interferon (IFN)–c [50]. Promiscuous
recombination of the gene locus with silent cassettes further down the
chromosome occurs unidirectionally [49] and leads to new antigenic
variants, ensuring B burgdorferi survival and adaptation to the host. It
appeared that promiscuous recombination could produce millions of
antigenic variants [48], but the gene changes are localized to variable
regions of VlsE, and some areas are stable or nonvariable, suggesting that
the number of possible variations will level oﬀ in time [51]. The functions of
the nonvariable areas of VlsE are unknown, but one of these is useful for
diagnostic serologic testing. New diagnostic tests have been able to
overcome the problems of heterogeneity of antigens expressed by ﬁnding
immunodominant epitopes that are constant and nonvariable among the
pathogenic species of Borrelia and during the length of infection. Future
vaccine technology also will be directed by these ﬁndings.

Besides antigenic variation, there are several other ways that B

burgdorferi

can persist in its host. The host defends against many other

types of bacterial infections by limiting iron availability to the microbe;
however, B burgdorferi has no need for iron [52]. B burgdorferi can hide
in folds of cellular membranes, where it is less exposed to the host’s immu-
nity or to antibiotics [53]. Morphologically, in hostile environments, B
burgdorferi

can transform into granular and cystic spherical structures,

called spheroplasts, starvation- or L-forms (named after Lister Institute)
and blebs, which still may be antigenic [54]. Some variants can be converted
back to motile spirochetes in vivo or in vitro under the right conditions [55–
62]. B garinii cysts are resistant to freeze-thawing, and may allow the
organism to stay dormant in the host until conditions improve [55].

Motile B burgdorferi converted to cystic forms within one minute when

placed in distilled water and when transferred to growth medium [BSK-H],
sprouted one to ﬁve ﬁlaments that grew to normal motile B burgdorferi [56].

836

M.P. Littman / Vet Clin Small Anim 33 (2003) 827–862

B burgdorferi

convert to L-forms in cerebrospinal ﬂuid (CSF) of patients

with neuroborreliosis, thereby making isolation more diﬃcult [57]. L-forms
may be more likely to develop in media containing certain antibiotics [54,59]
and appear to be degraded by hydroxychloroquine [60] and metronidazole
[61] (to which motile B burgdorferi are resistant). Future treatment of Lyme
disease may entail a combination of medications to eradicate the motile and
cystic forms of B burgdorferi.

Clinical signs of illness associated with borreliosis in people

People exposed to B burgdorferi are much more likely than dogs to show

clinical signs (Table 1). Even without treatment, signs usually resolve, and
only one case of human mortality has been attributed to Lyme disease.
People usually have a rash (ECM) at the site of the tick bite within days,
which can expand to 15 cm or more, within days. The rash may have
a ‘‘bull’s eye’’ appearance as it clears. An early inﬂuenza-like illness often
occurs for about 1 week. Weeks later, arthritis, neurologic, or cardiac
manifestations may occur, and months later, possible chronic or in-
termittent arthritis, neurologic or dermatologic signs. Clinical signs depend
on the Borrelia subtype and the patient’s genetic make-up. Most patients
respond readily to treatment. Doxycycline usually is given, since human
granulocytic ehrlichiosis (HGE) can mimic Lyme disease, has caused
mortality, and does not respond to penicillins. Some people need retreat-
ment, and some appear to have an immune-mediated chronic form of Lyme
disease that continues despite clearance of the organism. Post-treatment
Lyme disease may occur in 10% of treated patients, usually those with HLA
haplotype DRB1*0401 (DR-4), whose chronic arthritis is nonresponsive to
antibiotics, and whose cultures and PCR tests are negative. It is thought that
this is caused by immune-mediated sequelae, secondary to molecular
mimicry of OspA with a self-antigen called LFA-1. These patients have high
anti-OspA antibody, even though they have not been vaccinated with an
OspA containing product. Thus OspA can be expressed in chronic
borreliosis in mammals, as antigenic variation exposes whatever antigens
are in the agent’s repertoire. There is debate about whether persistent Lyme
disease is caused by L-forms or fragments that are diﬃcult to culture,
persistence of viable spirochetes, or whether the signs are indeed caused by
immune-mediated disease.

Spontaneous canine borreliosis

First described in 1984 and 1985 [63,64], the most common presentation

of Lyme disease in dogs is that of acute or intermittent lameness with
swelling or pain of one or a few joints (often carpi), fever, lethargy,
anorexia, and perhaps local lymphadenopathy. Joint taps from inﬂamed

837

M.P. Littman / Vet Clin Small Anim 33 (2003) 827–862

joints generally show neutrophilic inﬁltration, but these are the same
changes expected in joints aﬀected with immune-mediated disease, other
tick-borne arthritis, and other diseases. Complete blood cell count (CBC)
and chemical screening are nonremarkable. Rarely, neurologic signs have
been attributed to Lyme disease [65–68], sometimes demonstrating high
CSF/serum Lyme titer ratios [65,66]. One case report of complete heart
block and myocarditis was attributed to Lyme disease [69]. Rashes have not
been described. At the Matthew J. Ryan Veterinary Hospital of the
University of Pennsylvania (MJRVHUP), dogs with polyarthropathy had
positive Lyme titers more often than the general hospital population (59%
versus 37%, M.P. Rondeau, personal communication, 2003). Western blots
on sera from the polyarthropathy dogs showed 53% had antibody to
natural exposure, 35% to natural exposure and vaccine, and 12% to vaccine
only [70]. In comparison, Western blots on sera from dogs with ‘‘post-
vaccinal Lyme-like illness’’ showed 53% with vaccinal antibody only and
33% with antibodies to natural exposure and vaccine [71].

Single titers suﬃce to look for borreliosis exposure, since clinical signs in

experimental dogs did not commence until after seroconversion. Dogs
respond quickly, usually within 1 to 2 days, after starting antibiotic therapy.
Antibiotics generally are continued for 2 to 4 weeks. If signs do not abate
within 1 to 2 days, steroids are added, to which most dogs respond.

Studies show that up to 50% to 90% of healthy, asymptomatic dogs

living in endemic areas are seropositive for Lyme disease (Table 2).
Although 4.8% of seropositive dogs demonstrated lameness, fever, anorexia
or lethargy, those signs were seen in 4.6% of seronegative dogs in those
areas [72]. In Connecticut, positive Lyme titers were seen in 68% of dogs
with lameness and 70% of nonlame dogs [73,74]. It is diﬃcult to know if B
burgdorferi

can be blamed for clinical signs when a dog is sick and has

a positive titer.

In dogs, a severe protein-losing nephropathy with renal failure has been

putatively associated with Lyme disease, but the incidence is not known.
Renal lesions in dogs with positive Lyme titers have been described in Lyme-
endemic areas [74–78], but no animal model exists to study it. Putatively
called ‘‘Lyme nephritis’’ or ‘‘Lyme nephropathy,’’ the disease is a unique
combination of immune-mediated glomerulonephritis, diﬀuse tubular
necrosis with regeneration, and lymphocytic-plasmacytic interstitial nephri-
tis. It is associated with positive Lyme titers. In some cases, special stains
showed spirochetes in the renal tissue [74–76] and urine culture isolated B
burgdorferi

in one case [75]. Aﬀected dogs were younger than dogs with

other glomerulopathies, and they were more often Labradors, [76] Golden
retrievers [76], or Shetland sheepdogs (N.A. Sanders, personal communi-
cation, 2002) when compared with the author’s hospital population. Illness
was noted for 1 day or up to 8 weeks, but presentation was generally because
of acute vomiting, anorexia, lethargy, and weight loss in the summer to late
fall months. Clinically, the dogs showed protein-losing nephropathy (PLN)

838

M.P. Littman / Vet Clin Small Anim 33 (2003) 827–862

and renal failure with azotemia, hypoalbuminemia, hypercholesterolemia,
hyperphosphatemia, proteinuria, and possible hemoglobulinuria, hematu-
ria, and glycosuria. Remarkable CBC changes included a nonregenerative
anemia, thrombocytopenia, and stress leukogram. Many of the cases were
hypertensive, and many became edematous during aggressive treatment.
Most dogs rapidly progressed and died within 1 to 2 weeks, but three dogs
lived for several months. About 30% had a history of lameness, usually
within 3 to 6 weeks of presentation, and some had been treated for Lyme
disease with clinical response. Almost 30% had been vaccinated for Lyme
disease. One dog had a positive Ehrlichia canis titer, but A phagocytophila
(Ehrlichia equi) or Babesia species infections have not been ruled out.
Neurologic signs (seizure, nystagmus) were noted in about 15% of dogs,
which might have been because of thromboembolic events, hypertension,
infection involving the CNS, or uremic encephalopathy.

The author monitors Lyme positive dogs for proteinuria, and if found,

a diagnostic work-up and treatment for PLN (doxycycline, angiotensin-
converting enzyme [ACE] inhibitor, low-dose aspirin) is initiated early,
before the dog shows outward signs of illness. The dogs seem stable for
longer times, but whether this is because of the intervention or to the natural
course of this disease is not known. Studies need to be done to characterize
this syndrome further and to study the possible protective or additive eﬀect
of vaccination. Because Lyme nephropathy is not seen in any model of
Lyme disease, it may be that Lyme seropositivity is merely a marker for tick
exposure, and ‘‘Lyme nephropathy’’ may really be caused by Ehrlichia,
Babesia

, Bartonella species, mixed-infections, or something undiscovered.

The Midwest seems to see less PLN in their Lyme-positive dogs than is seen
in the East, perhaps because of strain diﬀerences, genetic diﬀerences for
susceptibility in dogs, or the possibility of more mixed infections in certain
areas. Because thrombocytopenia, glomerulonephritis, and neurologic signs
have been noted in dogs with Ehrlichia or Babesia infections, the possibility
that these may be involved with ‘‘Lyme nephropathy’’ is intriguing.

Experimental canine borreliosis

Dogs injected with B burgdorferi grown in culture show seroconversion,

but not illness [39,79,80]. Cultured organisms do not simulate real exposure
because their surface proteins are not those from natural tick feeding. Many
elegant studies were done next by using natural tick challenges with about
15 to 20 ticks per dog, using ticks collected from highly endemic areas where
more than 60% to 80% of the ticks harbor B burgdorferi. Adult beagles
challenged by way of the tick model seroconvert but show no signs of illness
[39]. That is not unexpected since more than 95% of seropositive dogs in the
ﬁeld are asymptomatic [72]. Young beagle puppies (6 to 12 weeks old) were
used mostly in subsequent studies, because they often show clinical signs
[38,39,81–86]. About 2 to 5 months after exposure, young pups may have

839

M.P. Littman / Vet Clin Small Anim 33 (2003) 827–862

a 4-day self-limiting illness with anorexia, fever, lameness with joint swelling
in the limb closest to where the ticks were placed [38], and perhaps local
lymphadenopathy. Some dogs had a few recurrences, 2 to 4 weeks apart, but
these were also self-limiting without treatment, and the dogs were normal in
between and thereafter. A study with older pups (12 to 26 weeks old)
showed 77% developed transient lameness, but only for 1 to 2 days, and
none had recurrences [87]. None of the tick challenged dogs developed a rash
or cardiac, neurologic, or renal signs of illness, although histologically some
had pericarditis, perineuritis, meningitis, or rarely encephalitis or neuritis
[88]. Joint taps from joints with acute arthritis showed neutrophilic
inﬂammation [39]. There were no CBC or serum chemistry changes.
Necropsy of asymptomatic dogs (the carrier puppies later in life) showed
mild nonsuppurative inﬁltration of the synovia [39], but similar ﬁndings in
seronegative and vaccinated dogs make the clinical relevance of this
doubtful. [89] B burgdorferi has a predilection for connective tissue, and was
more often cultured in the dog model from skin, synovia, peritoneum,
pericardium, fascias, skeletal muscle, lymph node, and meninges (not from
blood, spleen, kidneys, urinary bladder, urine, brain, or CSF) [90]. Skin
biopsies at the site of the tick bite showed perivascular inﬁltration of
mononuclear cells, and B burgdorferi could be cultured from the skin more
than 1 year after exposure. Even after high doses of antibiotics for 30 days
(doxycycline 10 mg/kg every12 hours; amoxicillin 20 mg/kg every 8 hours,
azithromycin 25 mg/kg every 24 hours, or ceftriaxone 25 mg/kg intra-
venously every 24 hours), some dogs showed persistent infection, mostly
based on positive PCR tests, but a few had positive cultures of B burgdorferi
from skin biopsies [38,82–84]. There is much debate about persistence of
infection after treatment (Table 3).

Ticks were obtained from Lyme-endemic areas for these challenge

studies, and although serology rarely was done on experimental dogs, some
coinfections likely occurred. Coinfections may alter host immune response
and the severity of arthritis [96,97]. When looked for, seropositivity in
experimental dogs was found (four dogs had positive B microti titers [39],
and 9 of 20 dogs [81] and 7 of 20 dogs [88] had positive E equi
(A phagocytophila) titers). Although not statistically signiﬁcant, a trend
appeared, as B burgdorferi/A phagocytophila coinfected dogs comprised 7 of
13 (53%) dogs with more severe lameness compared with two of seven
(29%) dogs with less severe lameness [81]. And in a study showing more
neurologic involvement histologically than previously seen, the seven B
burgdorferi

/A phagocytophila coinfected dogs were not identiﬁed [88]. A few

studies found negative titers for Powassan virus or E canis antibodies, but E
canis

tests do not test for E equi (A phagocytophila) antibodies.

Another problem in the understanding of Lyme disease in dogs is that

‘‘Lyme nephropathy’’ has no animal model (Koch’s postulates have not
been satisﬁed). Primarily beagles have been used in these experimental
studies, and they have not shown renal changes described in some

840

M.P. Littman / Vet Clin Small Anim 33 (2003) 827–862

ble

Deba

conc

erning

persistence

infection

after

treatm

ent

for

dise

ase

idence

for

persistence

infe

ction

despite

treatm

ent

Reb

uttal

Some

infected

peop

and

dogs

are

ill

culture

sitive

aft

treatm

ent

[4–6,38,82

sma

perce

ntage,

and

retreatment,

leas

eople,

for

longe

wks)

usu

ally

clears

them

[4].

Onl

dogs

had

positive

culture

aft

treatm

ent

[38].

Findings

the

same

are

rep

orted

than

one

pape

[38,8

2].

Man

tick

-challen

ged

studies

show

that

PCR

tests

are

still

sitive

after

treatm

ent

[38,8

2–84

ese

studies

repea

ﬁ

dings

the

sam

gs.

Althou

man

had

positive

PCR

tests,

PCR

testin

does

not

diﬀe

rentiate

tween

viable

organi

sms

and

non

viable

fragm

ents

impote

spir

ochet

es.

PCR

correlates

well

with

cult

ure

result

[91].

the

PCR

positive

then

viable

org

anisms

are

probabl

still

there

were

very

recent

there,

within

few

wee

ks.

the

PCR

prob

was

for

plas

midial

DNA

(eg,

Osp

gene)

and

chromo

somal

geno

me,

are

done,

and

the

rat

imbalanced,

this

may

ind

icate

non

viable

damag

[92].

Studie

dogs

with

persistent

PCR

tests

but

negative

ltures

after

treatm

ent

used

only

OspA

gene

prob

[83,8

did

state

targe

ratio

thei

resu

lts

[38,8

2],

the

ﬁn

dings

are

diﬃcu

interpret.

Fragm

ents,

blebs,

cysts

mig

not

cleared

readily

they

are

intra

cellular

hid

ing

conn

ective

tissue

Mice

treate

ith

antib

iotics

have

damag

spiroc

hetes

that

may

picked

PCR

for

month

but

eventu

ally

are

cle

ared

and

are

infective

[93].

can

invade

synovi

cells

[53],

and

intracellular

organi

sms

uld

evade

eatment

and

the

immune

syste

Fib

roblasts

infected

intra

cellularly

with

were

damag

and

did

live

long,

organi

sms

ould

free

[94].

Some

infected

dogs

treate

with

antibio

tics

had

ELIS

titers

that

initially

droppe

then

arted

rise

again

months

lat

er,

presuma

bly

from

regrowth

org

anisms

left

behind

[82].

any

probabl

are

not

cle

ared

with

treatm

ent.

wer

VlsE

testing

sho

wed

that

eated

dogs’

titers

drop

ped

within

weeks

treatm

ent

and

staye

low

least

wks

thereafte

[95].

Most

dogs

prob

ably

are

cle

ared

infe

ction

with

eatment

Abbre

viatio

ns:

Bb,

rrelia

burgdorfe

;

PCR

polym

erase

chain

react

ion.

841

M.P. Littman / Vet Clin Small Anim 33 (2003) 827–862

spontaneous cases with Lyme exposure, mostly in retrievers. Beagles may
respond diﬀerently than other breeds (eg., beagles exposed to E canis
developed only mild chronic disease, whereas infected German shepherds
developed severe chronic ehrlichiosis). Beagles had an increased cell-
mediated response to the infection [98]. Perhaps if Labradors (a breed
predisposed to the putative Lyme nephropathy) were used to study Lyme
disease, diﬀerent results might be seen.

Pathogenesis and host immune responses can be studied in the dog

model. Many inﬂammatory cytokines are up-regulated in the presence of
live B burgdorferi [81,99], and some even are triggered by Osps or fragments
of nonviable spirochetes [86,99]. Because B burgdorferi can leave blebs
(fragments of membrane bound outer surface proteins, some of which
stimulate inﬂammation), chronic signs may be caused by these nonviable
remnants. Some of these fragments may contain plasmid DNA, and they
may be recognized by PCR with primers for the plasmidial OspA gene. If
other primers are used, however (eg, PCR with 23S RNA primers), they may
be negative or weak, because the full genome is not there. For instance, in
patients with late manifestations of Lyme disease, a discrepancy was proven
to exist during PCR testing of synovial ﬂuid; that is, DNA sequences of
plasmid-encoded genes OspA and OspB were detected easily, while that of
the B burgdorferi genome were not detected [92]. Perhaps fractions of B
burgdorferi

might include plasmid DNA encoding inﬂammatory Osps but

not DNA encoding whole viable organisms. Because B burgdorferi is very
slow growing (especially variants), there is still debate about this (Table 3).
Treatment of some carrier dogs with immunosuppressive doses of cortico-
steroids (prednisone 2 mg/kg every 12 hours for 2 weeks) showed that some
dogs that had not received antibiotics had reactivation of latent B
burgdorferi

infection with severe polyarthritis [38,89]. Signs of recurrence

were not seen during the 2 weeks of immunosuppressive therapy, but 4 to 6
days after steroids were stopped, and signs cleared without treatment after
6 to 8 days [38,83]. Signs were not seen in study dogs that had received
antibiotics for infection and then were immunosuppressed, even though
some had positive PCR tests. Because most treated dogs did not have
positive cultures (only positive PCR tests), the organisms might be in an
altered form, such as variants, fragments, or impotent spirochetes, and
thereby not able to be cultured [93].

Find the organism or antibody directed against the organism

Isolation and culture of B burgdorferi are diﬃcult because of the

organism’s slow growth and microaerophilic requirements. Blood samples
may yield very few organisms during illness [100], but sometimes positive
cultures are found even in asymptomatic cases [101]. Modiﬁed BSK
(Barbour-Stoenner-Kelly) media are generally used [10]. Staining techniques

842

M.P. Littman / Vet Clin Small Anim 33 (2003) 827–862

to see organisms require dark-ﬁeld microscopy, silver, acridine orange,
Giemsa, or direct ﬂuorescent antibody stains. The PCR can target speciﬁc
DNA of B burgdorferi (eg, by using primers for the OspA gene or 23S RNA
gene) [88]. If PCR testing for OspA is stronger than for genomic targets,
target imbalance may indicate that only fragments, remnants, or blebs of B
burgdorferi

persist [92]. These are nonviable, but they may be immunogenic.

Perhaps if these remnants are intracellular or hiding in connective tissue
cellular invaginations, they may show positive PCR for quite some time.
Ears and bladders of infected mice treated with antibiotics showed
concordance of culture and PCR; however, synovia or ﬁbrous tissue was
not studied [91]. Because PCR testing is not readily available to the clinician,
the question is mainly one for investigators.

Titers

Indirect evidence of exposure is found by serologic tests for antibody

directed against B burgdorferi antigens. Interpretation is diﬃcult, since
many healthy dogs in endemic regions are seropositive. Having a positive
titer was not predictive for development of limb or joint disorder; 4.8% and
4.6% of seropositive and seronegative dogs, respectively, developed limb or
joint disorder over 20 months of observation [72]. Indirect ﬂuorescent
antibody (IFA), ELISA, and kinetic enzyme-linked immunosorbent assay
(KELA) antibody tests are available, using whole cell cultured B burgdorferi.
Testing for IgM versus IgG is unnecessary in dogs, since dogs do not show
signs of illness before IgG rises. Because new antigens are expressed over
time, IgM antibody may be found in chronic cases, giving rise to confusion as
to when the dog was exposed. Titers remain high for years. Testing is not
standardized, and interlaboratory results are inconsistent [102]. Concerns
about cross-reactivity, especially from Leptospira antibodies [103], decreased
sensitivity [104] or speciﬁcity [105], and because IFA or ELISA titers do not
distinguish between vaccinal and natural exposure antibodies, Western blot
has been necessary in the past. Because bacterins are derived from cultured
organisms that display antigens as in the unfed tick (OspA, OspB), and
subunit OspA vaccines only would generate antibody to OspA (band at p31),
the Western blot pattern would appear diﬀerently in dogs that were
vaccinated, naturally exposed, or both (Table 4). Dogs vaccinated with
OspA subunit vaccines would have only OspA antibody bands, but dogs
vaccinated with bacterin would have many more bands. Interpretation is
complicated by cross-reacting antibodies for some bands (p41 ﬂagellin),
diﬀerent bands being important at diﬀerent times in infection, and over time,
anti-OspA and anti-OspB antibodies may appear in nonvaccinates. In-house
kits for people make use of recombinant antigens known to be most
commonly seen with natural exposure. Other tests include bactericidal assays
[87], T-cell reactivity to B burgdorferi [114], and an assay that releases bound
Lyme antigen from antibody complexes [115].

843

M.P. Littman / Vet Clin Small Anim 33 (2003) 827–862

ble

Vario

author

s’

ﬁndings

conc

erning

Weste

Blot

band

seen

with

infe

ction

versus

vac

cination

Natural

exp

osure

Borrelia

burgdorf

eri

Lyme

vaccinat

ion

(bac

terin)

Weste

Blot

[106]

[107]

[108]

[109]

(%)

[71]

[110]

[111]

[112]

(%)

[4]

[113]

[108]

[71]

[112]

(%)

[113]

P93

IgG

P83

P66

IgG

P61

P60

P58

IgG

P45

IgG

P41

ﬂag

ellin

þþ

IgM

IgG

P39

þþ

IgM

P37

P35

P34

Osp

þþ

P31

Osp

þþ

P30

IgG

P28

Osp

IgG

P26

P25

P23

Osp

IgM

IgG

P22

P21

844

M.P. Littman / Vet Clin Small Anim 33 (2003) 827–862

P19

Osp

P18

IgG

P17

P15

P14

For

[106]

tho

bands

marke

were

intensely

and

consis

tently

tected

using

all

rains

burgdorfe

;

other

band

varie

especially

regions

and

kilo

dalton

[71]

req

uires

bands

for

natu

ral

expo

sure;

cross-

reaction

ith

too

many

oth

infe

ctions/vaccina

tion

07]

requir

comm

IgM

band

earl

infection

and

comm

IgG

band

lat

infe

ction.

[4]

req

uires

mmon

IgM

band

early

infe

ction

and

mmon

IgG

band

later

infection,

but

they

are

diﬀerent

than

[107]

bbrevia

tions:

canin

earl

infe

ction;

huma

later

infe

ction;

band

most

pronou

nced

and

expected

The

cases

show

ing

that

band,

ind

icating

antib

ody

that

antig

en,

their

Western

Blot

49%

natu

rally

exposed

dogs

did

hav

low

titers

Osp

ose

band

marke

ere

inte

nsely

and

nsistently

detecte

845

M.P. Littman / Vet Clin Small Anim 33 (2003) 827–862

Veterinarians now have a revolutionary in-house test, the SNAP-3 Dx

Test (IDEXX Laboratories, Inc., Westbrook, ME). This ELISA assay for
antibody directed against B burgdorferi uses a 26 amino acid recombinant
C6 peptide as its antigen. The C6 peptide is named for the immunodominant
invariable region 6 (IR-6) within the VlsE (variable major protein-like
sequence, expressed) surface antigen, which is expressed in vivo during early
and late infection, even as antigenic variation changes other parts of the
VlsE antigen. IR-6 is immunodominant in dogs and people and conserved
among strains of B burgdorferi. Because VlsE is not expressed much in vitro
and is masked by comigrating bands, there is no analogous Western blot
band of antibody [116]. Study dogs with natural challenge with B
burgdorferi

all showed a positive C6 test within 3 to 5 weeks, before they

showed any clinical signs, and they maintained a positive test for at least 69
weeks [85]. None of the vaccinates with bacterin or subunit OspA vaccine
showed positive test results, and the test appeared to be 94% to 100%
sensitive and greater than 99% speciﬁc [85,117]. Because the test is able to
distinguish between natural exposure and vaccinal antibody, the more
arduous, expensive, and sometimes unclear Western blot test is not needed.
Studies on experimentally infected beagles showed that C6 peptide
antibodies generally wane by 12 weeks after antibiotic therapy and stayed
low during the study thereafter (35 weeks) [95]. Although the in-house test is
qualitative, samples can be sent to a reference laboratory for quantiﬁed C6
antibody testing. The in-house test also tests for heartworm and E canis
antibodies, but the E canis test is not as accurate (89.8%) as another test, the
recombinant major antigenic protein 2 (rMAP2) ELISA for E canis
antibodies, with 97% accuracy [118]. Although E ewingii and E chaﬀeensis
antibodies cross-react with E canis tests, E equi (A phagocytophila) anti-
bodies do not, and separate titers need to be done (or PCR testing for
Ehrlichia

species on a pretreatment sample).

Controversy exists involving how to interpret the information from this

revolutionary new test. Because the SNAP-3 includes a heartworm test, it
likely will be run on many healthy dogs. The ﬁnding of many healthy Lyme-
positive dogs in one’s practice has started much debate about what to do
with these asymptomatic positive dogs. Should they be treated? Which
antibiotic should be used, at what dose, and for how long? Should these
dogs be vaccinated? Because most dogs with serologic evidence of exposure
will be asymptomatic, this would comprise a large population of dogs. In
some areas, 50% to 90% of the healthy dogs will be seropositive (see Table
1), and only less than 5% will be symptomatic. There are no prospective
studies that show how many of these dogs will ever show illness caused by B
burgdorferi

during their lives; how many of those that become ill will have

self-limited or easily treatable disease; or how many will have more major
problems, such as ‘‘Lyme nephropathy.’’ How many will spontaneously
clear their infection, and how many will continue to be asymptomatic
carriers? Is their subclinical carrier status protecting them from reinfection,

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M.P. Littman / Vet Clin Small Anim 33 (2003) 827–862

or will they possibly have reactivation of latent illness if immunocompro-
mised or stressed? Although experimental carriers treated with steroids
developed ‘‘reactivation’’ of clinical signs [83], coinfections were not ruled
out, and the dogs were not weaned from high-dose steroids as usually is
done. The author has not recognized clinical evidence of any Lyme disease
signs in VHUP patients after treating them with immunosuppressive
medications for various diseases.

Historically, dogs with acute Lyme arthritis were treated successfully with

only a 2-week course of antibiotics. This probably put them into a
premunitive state. The author feels an asymptomatic Lyme positive dog
probably does not need to be treated, but it does need to be monitored for
proteinuria (early Lyme nephropathy), and the owners need to be aware of
good tick control for their pets and the implications of living in an area
where Ixodes ticks are carrying B burgdorferi and probably other organisms.
If signs of illness (lameness or proteinuria) occur, the dog is, of course,
treated, and a more thorough work-up can be done (Fig. 4).

Others agree that asymptomatic dogs do not need to be treated [8]. But

one author [22] recommends treating all asymptomatic Lyme-positive dogs
with high doses of antibiotics, doses that were found ineﬀective of clearing
dogs [38,82–84]. That same author also recommends giving Lyme vaccine
(bacterin) to that asymptomatic carrier dog on day 0 and day 14 of
doxycycline.

The author feels there is no rationale for doxycycline or Lyme

vaccination, other than to sell these products. First of all, dogs that have
titers without signs of illness are probably not genetically predisposed to
have severe forms of Lyme disease. They are either already in a premunitive
state, or they may be having occult proteinuria from early Lyme
nephropathy. In the ﬁrst situation, dogs do not need treatment unless they
become symptomatic. The dose recommended is the same dose that left
most dogs with positive PCR tests and some with positive cultures [38,82–
84]. If the infection cannot be cleared, the only expectation with treatment
is to knock them into a subclinical, premunitive state, where they probably
are already. Think of the premunitive state as in the carrier cat with
Mycoplasma

(Haemobartonella) that is PCR-positive for Mycoplasma before

and after treatment. It is likely that B burgdorferi is quiet in collagen and
areas that are treated less easily with antibiotics. Treating a lot of
asymptomatic carriers with high doses of doxycycline (10 mg/kg twice daily
for 30 days) will cause adverse eﬀects in some and microbial resistance at
large. In the second situation, if the dog is genetically predisposed to having
an immune-mediated glomerulonephritis from exposure to Lyme antigens
and is starting to deposit immune complexes within the kidney, adding more
Lyme antigens by vaccinating would be unwise. Vaccines are intended only
to protect a dog from infection before it is exposed, and even then, Lyme
vaccination is controversial.

847

M.P. Littman / Vet Clin Small Anim 33 (2003) 827–862

Fig

Flow

chart

for

lam

eness

anorexia

and

fever.

Flo

chart

for

Lyme

and

protein

uria.

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M.P. Littman / Vet Clin Small Anim 33 (2003) 827–862

Diﬀerential diagnosis, diagnostic work-up

History, signalment, and physical examination will help reveal if lameness

is caused by painful arthritis (Fig. 4). Inﬂamed joints may be caused by septic
arthritis, tick-borne arthritis (including Lyme disease, ehrlichiosis, Rocky
Mountain spotted fever, or bartonellosis), immune-mediated polyarthritis,
lupus, rheumatoid arthritis, degenerative joint disease, or trauma. A common
presentation of Lyme disease in the dog includes fever, anorexia, and joint
swelling of one or several joints, often the carpi and hocks. A thorough
diagnostic work-up often includes CBC, chemical screening, urinalysis,
testing for infectious agents such as Lyme, E canis, E equi, Rocky Mountain
spotted fever, and Bartonella species, rheumatoid factor, antinuclear antibody
(ANA), joint taps for cytology or culture, and radiographs of one or more
joints. To rule out neoplasia, chest radiographs and abdominal ultrasound are
recommended. Aspirates of lymph node or bone marrow are helpful in cases
with lymphadenopathy or CBC abnormalities. Because acute cases of many
infectious diseases will be seronegative, a complete work-up should include
convalescent titers for those diseases.

Pain or swelling close to a joint may be caused by myositis, panosteitis,

neoplasia, osteomyelitis, hypertrophic osteodystrophy, or hypertrophic
osteopathy. Radiographs, serum creatine kinase, Neospora, Toxoplasma or
Leptospira

serology, and ANA may be helpful in those cases. Diﬃculty in

rising also may be caused by weakness from metabolic, cardiopulmonary,
neurologic, or vascular diseases, which entail specialized work-ups for those
problems.

If PLN is suspected, many of the tests recommended for polyarthropathy

are done, since infectious, immune-mediated, and neoplastic diseases may
cause vasculitis and proteinuria. Familial glomerular disease or a pre-
disposition may exist in Labradors, Golden retrievers, and Shelties,
triggered by exposure to B burgdorferi. In addition, blood pressure
measurements, urine protein/creatinine ratio, urine culture, heartworm
testing, Leptospira titers, and Coombs’ tests are useful. Renal biopsy may be
done to diﬀerentiate glomerulonephritis, glomerulosclerosis, or amyloidosis.

Although other causes of polyarthropathy or PLN may be ruled out, E

equi

(A phagocytophila) or B microti have not been ruled out, and perhaps

there are other infectious diseases yet undiscovered. In Lyme-endemic areas,
coinfection with other agents of the guild in Ixodes ticks (see Table 2) and
other infectious diseases (eg, Leptospira) may cause confusion as to what
symptomatology is caused by B burgdorferi. Luckily, most of these diseases
are responsive to doxycycline, and ‘‘doxycycline-responsive disease’’ may be
diagnosed. One agent that is probably not as responsive is B microti.
Coinfections of B burgdorferi and B microti have increased symptomatology
in people [119] and mice [97]. In southern New England, 10% of Lyme
disease patients [119] and 76.9% of mice [16] carried B microti. More study
is needed concerning B microti and dogs.

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M.P. Littman / Vet Clin Small Anim 33 (2003) 827–862

Anaplasma phagocytophila

(E equi, HGE) is now recognized as causing

illness in people [120] and dogs [121] living in Lyme-endemic areas. Evidence
of A phagocytophila was found in 20% to 35.6% of Lyme disease patients
[24,25] and in 9% to 25% of Ixodes ticks [18]. In one veterinary practice,
seroprevalence for A phagocytophila in Lyme positive dogs rose from 15.6%
in 1985 to 40% in 2001, proving that coinfections are quite common in that
area. Coinfections may induce more serious disease or complicate the
presentation. It is unclear how many prior cases diagnosed as Lyme disease
may really have had A phagocytophila or coinfections. Because HGE signs
in people may mimic Lyme disease (but with much more mortality),
doxycycline is recommended as ﬁrst choice for patients with acute fever,
inﬂuenza-like signs, possible rash, and myalgia/arthralgia. Thrombocyto-
penia is more likely with A phagocytophila than with B burgdorferi, and
neutrophils may contain morulae. In some cases at MJRVHUP, morulae
were seen in neutrophils of peripheral smears or joint taps. Response is
generally rapid (within 1 to 2 days) with doxycycline. PCR testing on
a sample taken before starting treatment, or paired titers for E equi are
recommended, since acute titers are most likely negative. Antibodies
directed against E equi do not cross-react with E canis antigens, so the E
canis

tests commonly available would not be helpful. Another granulocytic

Ehrlichia

agent, E ewingii, is seen mostly in the South and can cause fever or

arthritis signs. Antibodies against E ewingii will be picked up on E canis
tests, however. A positive Lyme titer is a marker of exposure to Ixodes ticks,
and coinfections with other guild members (E equi, B microti, Bartonella,
even TBE virus) should be considered. Some of these (E equi, Barto-
nella

) may cause polyarthropathy and perhaps some (E equi, B microti,

Bartonella

) may cause protein-losing nephropathy. Bartonella has been

associated with refractory neuroborreliosis in people in New Jersey. [29]

In Europe, Western blot tests on Lyme-positive samples showed more

intense bands at p30 and p56 in symptomatic dogs compared with
asymptomatic dogs [122]. More studies may help clinicians understand if
B burgdorferi

is the cause of illness at presentation.

Response to therapy—‘‘a doxycycline-responsive disease’’

Clinically, dogs with signs of Lyme arthritis generally respond within 1 to

2 days of antibiotic therapy. Historically, clinicians generally treated acute
cases for 2 weeks and chronic intermittent arthritis for 4 weeks. No one
knows how long is long enough, and there is debate about dosing regimens.
Doses of doxycycline at 10 mg/kg every 24 hours or amoxicillin at 11 mg/kg
every 12 hours have yielded good clinical results (B burgdorferi is not
sensitive to quinolones or cephalexin). Experimental beagle models receiving
higher doses of antibiotics (doxycycline 10 mg/kg every12 hours, amoxicillin
20 mg/kg every 8 hours, azithromycin 25 mg/kg every 24 hours, or

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M.P. Littman / Vet Clin Small Anim 33 (2003) 827–862

ceftriaxone 25 mg/kg intravenously every 24 hours) for 30 days, however,
showed that not all dogs were cleared of infection. Three of 24 dogs were
documented by isolation or culture, but the others were by PCR alone (see
Table 3). Although their titers dropped initially after 6 months, titers rose
again presumably because of their carrier status and the recrudescence of
slow-growing Borrelia [82]. Follow-up was not long enough to know if they
might develop arthritic signs again or perhaps PLN. Clinically, if a dog
responds to treatment for Lyme arthritis and shows similar signs in the
future, it is unknown if the signs are caused by relapse, reinfection,
misdiagnosis with other doxycycline-responsive illnesses, auto-immune
disease, or something else.

Response to doxycycline may be because of occult treatment of other

infections or coinfections, including Ehrlichiosis, Rocky Mountain spotted
fever, Bartonella, Mycoplasma, and others. Although Lyme titers are
expected to be positive at the time of clinical signs (as in experimental
models), acute illness caused by other tick-borne diseases that mimic Lyme
disease and may be doxycycline-responsive might go undetected, because
acute titers would be seronegative, and convalescent tick titers often are not
done because of the owner’s ﬁnancial concerns. The positive Lyme titer may
be a sentinel of tick exposure, and the clinical signs might in some cases
have been wrongly attributed to Lyme disease. And since doxycycline and
the tetracycline family have anti-inﬂammatory, antiarthritic properties
[123,124], response to treatment is not as helpful in making a diagnosis
of Lyme disease. Degenerative joint disease or mild immune-mediated
polyarthropathy might respond to these eﬀects. Injuries may appear to have
responded to therapy, but time was the healer. Reactive arthritis caused by
gastrointestinal (GI) infections might respond to antibiotics [125].

In dogs that respond quickly to a short course of antibiotics, there is no

reason to double the dose, or increase the length of time treatment is given,
because there are no guidelines as to how long is long enough. Most dogs
with arthritis signs caused by B burgdorferi respond quickly to antibiotics
and are not referred to teaching hospitals. Most dogs diagnosed with
‘‘chronic Lyme arthritis’’ that has not responded to antibiotics are worked
up and found to have immune-mediated polyarthropathy. They generally
respond well to steroids. In people, there is debate concerning whether
‘‘chronic Lyme disease’’ is caused by persistent infection (see Table 3),
immune-mediated sequelae, or something else. Aggressive, long-term
antibiotic treatment in people with ‘‘chronic Lyme disease’’ helped as much
as placebo [126]. These patients probably have something other than
persistent B burgdorferi infection causing signs.

Treatment of PLN associated with Lyme disease is the same as that for

other forms of PLN, with the addition of doxycycline. The author often
gives an ACE inhibitor to decrease proteinuria by decreasing glomerular
ﬁltration pressure, and a low dose of aspirin (0.5 mg/kg every 24 hours) as
an antithrombotic. Omega-3 fatty acids also are recommended. If blood

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M.P. Littman / Vet Clin Small Anim 33 (2003) 827–862

pressure measurements are still high despite ACE inhibitor therapy,
amlodipine may be added. Asymptomatic dogs with proteinuria (early
PLN) are monitored monthly with urine protein/creatinine ratio (Up/c),
blood pressure measurements, and chemical screening. If they are stable
after 3 months, visits are less frequent. The author used to give doxycycline
long-term (no one knows how long is long enough), but the new C6 peptide
test will allow clinicians to make wiser decisions about whether antibiotics
are still necessary. Sick dogs (vomiting, dehydrated, anorexic) with PLN
need more intensive therapy, including dietary phosphorus and protein
modiﬁcations, phosphate binders, rehydration, colloid support, and other
treatments.

Prevention

For dogs living in endemic areas, whether seronegative or seropositive, it

is important to decrease the risk of exposure to B burgdorferi by avoiding
wooded areas and vegetation (tick territory) when possible, and using good
tick control. The author advises mowing lawns back and getting rid of brush
where possible. Near the author’s practice, there are multiple tick-borne
diseases besides B burgdorferi to consider (E equi, E canis, Rocky Mountain
spotted fever, and babesiosis). An excellent method of tick control is the
amitraz collar (Preventic collar, Virbac, Inc., Fort Worth, TX). When put
on tightly enough to have contact with the skin (not just the fur), after
a day’s wearing, the amitraz chemical ‘‘coats’’ the skin and is able to keep
ticks from being able to attach to the dog, by paralyzing their mouth parts.
Control dogs all seroconverted, but all dogs wearing the amitraz collar
stayed seronegative, despite exposure to 100 ticks in one study [127]. The
amitraz chemical washes away, however, so this method may not be good
for dogs that swim, and the collar is toxic if eaten (antidote: yohimbine).
Amitraz should not be used on dogs given certain behavioral medications.
Owners with multiple pets should be advised to have the dogs wear a wider
cloth or nylon collar atop the amitraz collar so that other dogs cannot get to
the amitraz collar during rough play. For dogs that swim or are bathed
often, a topical product which does not wash oﬀ (ﬁpronil, Frontline Top
Spot, Merial Ltd., Iselin, NJ) should be used. This product works by killing
ticks after they have attached. The ticks generally die within 24 to 36 hours
of exposure to this chemical, which is before transmission of B burgdorferi,
but it may not be a fast enough kill to prevent exposure to other tick-
borne diseases (eg, E equi is transmitted within 24 hours) [34]. The author
has used amitraz with ﬁpronil together on dogs without adverse reactions.
Permethrin containing products such as Advantix (Bayer Animal Health,
Shawnee Mission, KS) or ProTIcall (Schering-Plough Animal Health,
Kenilworth, NJ) repel and kill ticks, but should not be used around cats.

The author sometimes receives questions when owners pull ticks from

their dogs. The owners ask for a 2- to 4-week course of antibiotics to prevent

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M.P. Littman / Vet Clin Small Anim 33 (2003) 827–862

infection. There is little indication for this. A recent human study showed
that one dose of doxycycline, given within 72 hours of removal of an
engorged tick, prevented people from showing Lyme disease and from
seroconversion [128]. The dose is unnecessary if the removed tick is ﬂat (not
engorged). Although no similar study was done on dogs, the author would
rather give a panicked owner one dose of doxycycline for their dogs rather
than a month’s worth, and monitor the dog before and 5 to 6 weeks later
with a C6 peptide test, if concerns remain. If the C6 peptide test is ever
positive in an asymptomatic dog, the author recommends checking for
proteinuria.

Vaccines available but controversial

Although vaccines are available for prevention of Lyme disease in dogs,

administering any Lyme vaccine is still controversial (see Table 1). Not all
dogs respond, but the preventive fraction for seroconversion in ﬁeld trials
using bacterin was high at 92.2% [129]. Although the recombinant,
nonadjuvanted OspA subunit vaccine protected 100% of the dogs in one
study [130], it did not protect one of two dogs [89] in another study. Because
vaccinal antibody is not long lasting, booster vaccines are recommended
annually after an initial series of two vaccines. Adverse events are considered
moderate [131], with anaphylaxis sometimes reported. More study is needed,
especially concerning possible reactions to proinﬂammatory surface antigens
or long-term immune-mediated sequelae. All vaccines (bacterin and
recombinant subunit OspA vaccines) expose the dog to OspA. Anti-OspA
antibodies taken in by the tick during feeding kill the agent before
transmission. Bacterins also expose the dog to other antigens, which may
induce other bactericidal antibodies but also could introduce other antigens
with proinﬂammatory, anaphylactoid, or immune-mediated sequelae.

The author’s concern about vaccinating against Lyme disease is long-

standing [132,133]. Brieﬂy, most dogs (95%) do not get ill from natural
exposure to B burgdorferi, so they do not need vaccination. They
seroconvert and remain asymptomatic. Although preventive fractions for
seroconversion may look good, there are no data to show preventive
fraction for clinical signs, which would indicate beneﬁt in the beneﬁt/risk
calculation. Veterinarians report that since using Lyme vaccination
programs, they are not seeing as many dogs with acute Lyme disease signs.
Veterinarians who do not vaccinate their patients also are reporting this,
however. Epidemiologically, this makes sense, as B burgdorferi prevalence in
ticks rose rapidly in some areas, exposing dogs of all ages and causing signs
in genetically predisposed dogs. Now there are fewer acute cases per year,
because there are fewer naı¨ve dogs to infect, and even practices not using
vaccine are seeing fewer of these cases. The next round of illness might be
caused by A phagocytophila. Near the author’s practice good tick control is

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M.P. Littman / Vet Clin Small Anim 33 (2003) 827–862

critical anyway, to protect against Ehrlichia, Rocky Mountain spotted fever,
and other tick-borne diseases.

Another reason the author does not use Lyme vaccines is because dogs

that show arthritis attributed to Lyme disease are generally responsive to
a short course of inexpensive oral antibiotics (or they may have several self-
limiting bouts that do not need any treatment, as in the tick challenge dog
model of Lyme arthritis). If Lyme nephropathy is caused by B burgdorferi,
then Lyme disease might involve signiﬁcant morbidity and mortality. The
author still would not recommend vaccination, however, because that
syndrome is associated with immune-mediated glomerulonephritis, and
while the antigens that are involved are unknown, they may be antigens
injected with vaccination. Dogs predisposed genetically to respond to Lyme
antigens by having Lyme nephropathy also might be the ones at risk to
develop problems from vaccine. Anti-OspA antibodies have been associated
with immune-mediated chronic arthritis in people because of molecular
mimicry of OspA with human LFA-1 [134]. OspA induces inﬂammatory
cytokines, arthritis, and sensitization with increased signs in rat and hamster
models [133]. Sensitization causes more intense inﬂammation when the
antigens are seen again.

Sensitization is a problem in vaccine development for diseases with an

immunopathogenic basis (eg, feline infectious peritonitis [FIP]) [132,133].
Monoclonal anti-OspA antibody staining revealed OspA expression in the
kidney from a dog with ‘‘Lyme nephropathy’’ [74]. A young dog that
became critically ill 4 days after an OspA booster vaccine had typical ‘‘Lyme
nephropathy’’ signs and renal lesions at VHUP; it had a high anti-OspA
titer but no evidence of natural exposure to B burgdorferi, based on the
SNAP-3 IDEXX test and Western blot (S.C. Fincham, unpublished data).
Although other tick-borne diseases were not ruled out, a postvaccinal Lyme-
like syndrome was described in dogs without evidence of natural exposure to
B burgdorferi

on Western blot [71]. Because ticks usually carry B burgdorferi

at higher rates than other agents causing fever/arthritis, and the dogs were
seronegative for Lyme antibody, it is possible the dogs reacted to
inﬂammatory antigens from the vaccine. Vaccine development is diﬃcult
for diseases involving immunopathogenetic mechanisms, because the animal
may become sensitized [132]. Because molecular mimicry of Lyme antigens
to tissue auto-antigens is a concern, the author does not recommend
vaccination for Lyme disease until these mechanisms and the real risks of
natural Lyme disease are understood better.

In February 2002, the one vaccine available for people (LYMErix, Table

1) was taken oﬀ the market, citing poor sales, which resulted from concerns
about possible postvaccinal illness in genetically predisposed people who
might get auto-immune disease from the vaccine or its boosters. The basis
for this concern is that 10% of people have treatment-resistant chronic
Lyme disease, even after clearance of the organism, based on PCR and
culture. These people have very high anti-OspA antibodies (even though

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M.P. Littman / Vet Clin Small Anim 33 (2003) 827–862

never vaccinated), OspA-sensitized T cells, and usually have HLA-DR4
haplotypes. This haplotype occurs in 30% of the population. OspA was
shown to have molecular mimicry with the auto-antigen LFA-1, causing
autoimmune arthritis [134,135]. Other antigens of B burgdorferi also have
stimulated antibodies that cross-react with auto-antigens of nerve axonal
protein, myelin basic protein, heat shock protein, muscle or myosin,
cardiolipin, and gangliosides [133,134,136].

Summary

A guild of organisms carried by the same vector (Ixodes ticks) in Lyme-

endemic areas may be confounding the understanding of Lyme disease in
dogs. A new diagnostic method, the C6 peptide test for Lyme, and serology
and PCR testing for Ehrlichia, Babesia, and Bartonella species will help to
sort out seroprevalence and symptomatology caused by exposure to these
agents or by coinfections. In addition, Rickettsia, Leptospira, Mycoplasma
species, and more could be involved in dogs diagnosed with a ‘‘doxycycline-
responsive’’ disease. The author does not recommend treating asymptomatic
Borrelia

carrier dogs, but does recommend screening them for proteinuria

and for exposure to other agents. A positive Lyme titer is a marker of
exposure to Ixodes ticks and the agents they carry. The risk/beneﬁt of
vaccination will be understood better as the symptomatology and im-
munopathogenesis of Lyme disease are deﬁned. Meanwhile, tick control is
highly recommended for all dogs in Lyme-endemic areas.

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Ehrlichiosis and related infections

Leah A. Cohn, DVM, PhD

College of Veterinary Medicine, University of Missouri–Columbia, 379 East Campus,

Clydesdale Hall, Columbia, MO 65211, USA

Ehrlichieae are gram-negative pleomorphic cocci capable of causing

disease in people and in several species of domestic and wild animals. They
have more in common with their close cousins the Rickettsia than they do
with most other gram-negative organisms in that they do not cause
endotoxemia, and they require a vector for transmission. These organisms
are found in membrane-lined vacuoles within the cytoplasm of infected
eukaryotic host cells, most often leukocytes. The obligate intracellular
location of these organisms makes an eﬀective host immunologic response
diﬃcult, and this complicates antimicrobial therapy.

Although ehrlichiosis ﬁrst was recognized as a disease of dogs in Algeria

1935, the prevalence and scope of the disease was not recognized until much
later. During the Vietnam War, US military veterinarians observed an illness
in military dogs that came to be called tropical pancytopenia. This illness was
caused by infection with Ehrlichia canis, which also was documented as a cause
of disease in dogs that had never left the United States. During the ensuing
decades, infections attributed to species of Ehrlichia other than E canis were
documented as important causes of disease in dogs and horses [1,2]. Although
naturally acquired feline infection was documented well over a decade ago,
researchers only now are beginning to elucidate the relevance of ehrlichiosis in
cats [3]. The recognition of ehrlichiosis as a potentially fatal disease of people
in the late 1980s prompted an intensiﬁcation of research centering on these
organisms [4]. The recognition of several species related to but distinct from E
canis,

combined with the availability of sophisticated molecular techniques,

has lead to a number of recent discoveries regarding these organisms, cul-
minating in an extensive reclassiﬁcation with changes in nomenclature. This
article describes the recent proposals for change and refers to each organism
by its new moniker. For purposes of this article, disease descriptions focus on
ehrlichial disease of dogs, but the diseases in other species are mentioned.

Vet Clin Small Anim

33 (2003) 863–884

E-mail address:

cohnl@missouri.edu

0195-5616/03/$ - see front matter

doi:10.1016/S0195-5616(03)00031-7

Taxonomic relationships of ehrlichial species

Traditionally, the species within the genus Ehrlichia were grouped by

cellular tropism. In this system, Ehrlichia were divided into monocytic forms
(eg, E canis and E risticii), granulocytic forms (eg, E ewingii and E equi), and
thrombocytic forms (ie, E platys). This system had a number of limitations,
including the fact that cell tropism is not absolute; a single infection can occur
in more than one cell type. A more objective classiﬁcation scheme uses
sequence homology of ribosomal RNA (rRNA) genes to determine the genetic
relatedness of the various organisms. Most Ehrlichia species have been
sequenced in this manner, leading to a recent proposed major revision in
taxonomic classiﬁcation of these bacterial organisms [5].

Several organisms previously included in the genus Ehrlichia have been

reclassiﬁed and moved into other genus groupings, and all of these
organisms have been moved from the family Rickettsiaceae and into the
family Anaplasmataceae (Fig. 1) [5]. Just before reclassiﬁcation, Ehrlichia
species had been divided into three distinct ‘‘genogroups’’ based on rRNA
relatedness; now, members of the second and third genogroup have moved
out of the genus Ehrlichia altogether. These diﬀerences in genogroup and
genus are more than academic technicalities. For instance, genetic relation-
ships explain why Potomac horse fever (Neorickettsia risticii) shares far
more in common with salmon poisoning disease (N helminthoeca) than it
does with E canis or Anaplasma phagocytophila infections. Additionally,
serologic cross-reactivity is marked among members of the same genus
(previously genogroup), while cross-reactivity between each genus is min-
imal. This means that to diagnose ehrlichiosis, titers should be performed
to a member of the same genus responsible for infection.

The family Anaplasmataceae contains four genra: Ehrlichia, Anaplasma,

Neorickettsia

, and Wolbachia. All but the later contain animal pathogens

formerly described as members of the genus Ehrlichia. Using the
reclassiﬁcation scheme, genogroup 1 Ehrlichia retain the genus name, while
members of genogroup 2 change from Ehrlichia to Anaplasma, and members
of genogroup 3 become Neorickettsia (Fig. 1) [5]. Because the organisms
previously known as E equi, E phagocytophila, and the human granulocytic
ehrlichial agent (HGE) diﬀer in only three nucleotides at most, these
organisms have become one species, A phagocytophila [5]. For purposes of
completeness, several organisms formerly described as Ehrlichia but now
classiﬁed as members of other genra will be included in this article. Correct
names derived from the reclassiﬁcation will be used when referring to
a speciﬁc species (Table 1).

Disease transmission

Ehrlichiosis, anaplasmosis, and neorickettsiosis are transmitted to

mammals by way of vectors. Infection with Ehrlichia and Anaplasma are

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L.A. Cohn / Vet Clin Small Anim 33 (2003) 863–884

Fig

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L.A. Cohn / Vet Clin Small Anim 33 (2003) 863–884

ble

Char

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ase

866

L.A. Cohn / Vet Clin Small Anim 33 (2003) 863–884

transmitted through the salivary secretions of an attached tick, while trem-
atodes serve as vectors for transmission of Neorickettsia species (Table 1).
The life cycle of the various trematode vectors is complicated and includes
multiple other animal species (eg, ﬁsh, snails, and aquatic insects) that may
pass on infection indirectly to pets. Several diﬀerent tick species are capable
of horizontal transmission of infection from vector to eukaryotic host. E
canis

usually is spread by the bite of the brown dog tick (Rhipicephalus

sanguineus

), which also can transmit infection with E ewingii and probably

A platys

. E ewingii is transmitted predominantly (but not solely) by the lone

star tick (Amblyomma americanum). Ixodes species ticks are responsible for
transmission of A phagocytophila infection. Geographic distribution of tick
vectors has a direct impact on disease prevalence in a given region [6]. For
instance, in the United States, A phagocytophila infection is reported most
often in the northwestern, upper midwestern, and northeastern states, the
same regions in which Ixodes ticks are most abundant. E ewingii infection
also follows the distribution of its tick vector, being found most often in the
southeastern and south-central United States. Vectors for several ehrlichial
species remain incompletely deﬁned.

Concurrent infection

Concurrent infection with multiple arthropod-borne pathogens has been

well document in both case reports and in epidemiologic studies [7–13]. A
single tick species can serve as a transmission vector for more than one type
of infectious agent. For instance, R sanguineus is capable of transmitting not
only E canis and E ewingii, but Babesia canis and perhaps B gibsoni as well,
and Ixodes ticks are competent vectors for transmission of A phagocytophila
as well as Borrelia burgdorferi and Babesia microti [1]. Either a single tick
infected with multiple pathogens or multiple ticks on a single animal could
be responsible for simultaneous infection with diﬀerent pathogens in one
host. Co-infection with multiple pathogens no doubt complicates disease
diagnosis and treatment, and has likely confounded results from retrospec-
tive studies that purport to describe the clinical course of ehrlichial infection.
Clinical signs or outcomes ascribed retrospectively to one type of infection
might have in fact been due to a diﬀerent infectious agent altogether, or to
the synergistic eﬀects of multiple infecting organisms.

Ehrlichial pathogens of pet animals

Ehrlichia canis

was the ﬁrst reported ehrlichial veterinary pathogen and

is perhaps the best studied. This organism is found worldwide, although

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L.A. Cohn / Vet Clin Small Anim 33 (2003) 863–884

infection is most prevalent in warmer climates. Persistent infection allows
canid hosts to serve as reservoir for infection when ticks feed ﬁrst on an
infected, then on an uninfected host [14]. Because there is no transovarial
transmission in the tick, the tick itself cannot serve as disease reservoir. Unlike
acute arthropod-borne infection, the chronic nature of the disease caused by
E canis

infection means that there is not a strong seasonal incidence.

Variable clinical ﬁndings associated with E canis infection likely are

inﬂuenced by strain of organism, host immune status, and even dog breed
[14,15]. German shepherds are said to have a more fulminant course of
infection than other breeds [15–17]. Clinical infection with E canis is described
as including three stages; acute, subclinical, and chronic infection. The acute
illness occurs 1 to 3 weeks after a dog is bitten by an infected tick and is
typically mild [18]. The organisms invade and replicate within host mono-
nuclear cells where they form morulae, or bacterial colonies bound by
a vacuolar membrane. The most consistent hematological change during
acute infection is a thrombocytopenia that results from vascular endothelial
inﬂammation with resulting platelet consumption, immunologically mediated
destruction of platelets, and splenic sequestration of platelets [18–20]. Clinical
signs during this phase are vague but often include lethargy, fever, anorexia,
weight loss, splenomegaly, and generalized lymphadenopathy [14,21]. Few
dogs succumb to the acute disease; most clear the organism (likely by way of
cell-mediated immunity) or enter the subclinical stage of infection [14,18].

During the subclinical phase of infection, the dog remains apparently

healthy, while organisms are retained at low numbers in splenic mono-
nuclear cells [22]. The length of the subclinical phase may range from weeks
to years. In fact, it is not known what factors inﬂuence progression from the
subclinical to chronic disease state, nor is it known what percentage of
subclinically infected dogs will develop illness. Hematologic changes
suggestive of ehrlichiosis, such as thrombocytopenia and hyperglobu-
linemia, may be detected during the subclinical phase of infection [23].

Ehrlichiosis caused by E canis often is diagnosed during the chronic

phase of infection. Nonspeciﬁc clinical signs include lethargy, anorexia, and
weight loss [15,24]. Signs attributable to bleeding tendencies, including
epistaxis, melena, petechial or ecchymotic hemorrhages, hyphema, retinal
hemorrhage, or hematuria, occur in 25% to 60% of cases [15,24–26].
Lymphadenopathy, fever, pale mucous membranes, and splenomegaly
sometimes are found on physical examination [15,24–26]. Occasionally
anterior uveitis, retinal changes, or neurologic abnormalities are noted
[24,27]. Ataxia, paraparesis, conscious proprioceptive deﬁcits, head tilt,
nystagmus, and seizures have been reported as neurological manifestations
of infection [24]. Although lameness often is listed as a clinical sign of
infection, many cases displaying lameness had granulocytic morulae,
suggesting the infection may have in fact been caused by E ewingii rather
than E canis [28–30]. Other clinical signs of chronic E canis infection relate
to complications of infection, including glomerulonephritis with nephrotic

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L.A. Cohn / Vet Clin Small Anim 33 (2003) 863–884

syndrome or pancytopenia resulting in secondary infections, severe anemia,
or bleeding. Persistent E canis infection results in persistent antibody
formation but fails to clear the organism [14,18,31–33]. Some of the most
severe disease manifestations associated with chronic ehrlichiosis, such as
glomerulonephritis and hyperviscosity syndrome, result from a nonprotec-
tive yet exuberant humoral immune response [14,18].

Like historical and physical examination ﬁndings, clinicopathologic

abnormalities are nonspeciﬁc. Most cases display some degree of
thrombocytopenia, and mild-to-moderate nonregenerative anemia also is
identiﬁed commonly [15,24–26]. White blood cell counts may be normal,
increased, or decreased [24–26]. Hyperglobulinemia is observed in most
cases [15,24,25]. Albeit usually because of polyclonal gammopathy, hyper-
globulinemia may present as a monoclonal gammopathy that can be
mistaken for multiple myeloma [34,35]. Other laboratory abnormalities
noted with some regularity include hypoalbuminemia, elevated alkaline
phosphatase (ALP), and elevated alanine transaminase (ALT) concentra-
tions [24–26]. Proteinuria may occur independently or concurrently with
glomerulonephritis [24–26,36,37]. Although early descriptions of E canis
infection were of a ‘‘tropical pancytopenia,’’ the pancytopenic manifes-
tations appear to account for a small minority of cases in the United States.
Cytologic evaluation of bone marrow aspirates often displays increased
numbers of plasma cells, with hypoplasia (suggests chronic) or hyperplasia
(acute or chronic infection) of the other marrow elements [21,24,25,38].

Ehrlichia chaﬀeensis

was the ﬁrst documented ehrlichial pathogen of people

in North America. In people, the disease manifests with fever, headache, and
myalgias and may be fatal if left untreated [39,40]. E chaﬀeensis is related
closely to E canis, with which it shares a tropism for mononuclear cells.
Although primarily notable as a human pathogen, dogs are also susceptible to
infection with this organism. Experimentally infected dogs demonstrated only
mild-to-inapparent disease [41]. More serious signs, however, including
vomiting, epistaxis, lymphadenopathy, and anterior uveitis were documented
in three dogs naturally infected with E chaﬀeensis [8]. Only specialized testing
(ie, species-speciﬁc polymerase chain reaction [PCR]) will diﬀerentiate
infection with E canis from E chaﬀeensis since routine E canis titers would
be positive in dogs with either disease [8]. Because of this, it is unlikely that the
true incidence of E chaﬀeensis infection in dogs will be known without
epidemiologic studies designed to evaluate this question directly.

Ehrlichia ewingii

is one of two ‘‘ehrlichial’’ agents known to result in

granulocytic infection of dogs, with the other agent being A phagocytophila
(formerly E equi.) [10]. Unfortunately, identiﬁcation of granulocytic

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L.A. Cohn / Vet Clin Small Anim 33 (2003) 863–884

morulae does not diﬀerentiate A phagocytophila infection from E ewingii
infections, although the geographic distribution of the two disease diﬀers
somewhat; E ewingii predominates in the southeastern and south-central
United States. Because E ewingii is related closely to E canis, E canis titers
may be positive during infection [10]. Because E ewingii and A phagocytophila
infection can only be diﬀerentiated by special testing, many times a diagnosis
is left as ‘‘granulocytic ehrlichiosis.’’ Veterinary literature often describes
canine granulocytic ehrlichial infection as presenting with acute polyar-
thropathy, and this is particularly true in geographic regions more likely to
harbor E ewingii infection [28–30,42,43]. Joint pain and occasionally joint
swelling caused by eﬀusion are noted, and dogs are often febrile.
Splenomegaly and hepatomegaly have been reported. Most dogs display
mild-to-moderate thrombocytopenia, and bleeding tendencies may be seen
[10]. Meningitis has been documented, and dual infection with E canis and
E ewingii

has been reported in association with profound ataxia and

epistaxis [44,45]. Fatal infections seem to be extremely rare, and clinical
signs resolve quickly with appropriate therapy. In a review done at the
University of Missouri of dogs diagnosed with E ewingii infection, all
presented with lameness, and all responded rapidly and completely to anti-
microbial therapy (Leah Cohn, unpublished data, 2002). Unlike E canis,
there is little evidence of chronic infection with E ewingii, and thus infected
animals most are often presented in the spring and summer when the tick
vector is most active.

Anaplasma phagocytophila (formerly Ehrlichia equi)

The true incidence of A phagocytophila infection in dogs is unknown, but

A phagocytophila

may account for a signiﬁcant proportion of granulocytic

canine ‘‘ehrlichiosis’’ in the northeastern and upper midwestern states and
California, where equine infections are endemic [10,46,47]. Experimental
infection with A phagocytophila in dogs produced only mild clinical illness,
but naturally infected dogs have presented with nonspeciﬁc illness including
fever, lethargy, and thrombocytopenia [47,48]. There are no unique clinical
ﬁndings attributed to infection with A phagocytophila, but polyarthritis is
described far less frequently than for E ewingii [47]. Most cases presented in
autumn, and females were over-represented, ﬁndings in direct contrast to
the author’s own study of dogs from Missouri with granulocytic ehrlichiosis
attributable to E ewingii, where dogs were most often male and presented in
the spring and summer months (Leah Cohn, unpublished data, 2002) [47].
Routine titers for E canis are often negative during infection with A
phagocytophila

, and will certainly be lower than titers speciﬁcally designed

to detect A phagocytophila. Without an index of suspicion, the veterinarian
may not request the speciﬁc diagnostic testing required to identify infection
with this organism [10,47]. Infection with A phagocytophila is less host-speciﬁc
than are many other ehrlichial infections. The organism is responsible for

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equine granulocytic ‘‘ehrlichiosis,’’ tick fever in small ruminants, human
granulocytic ‘‘ehrlichiosis,’’ and it causes infection in cats also [2,49–51].

Anaplasma platys (formerly Ehrlichia platys)

The platelet tropism of A platys is unique among these ehrlichial-related

organisms, even though all of these infections may result in thrombocyto-
penia. Infection with A platys results in moderate to severe cyclic
thrombocytopenia in dogs, but bleeding is seldom problematic [52,53].
A platys

has a worldwide distribution, and more severe disease manifesta-

tions have been described outside of the United States [54–56]. There is some
evidence that platelets of dogs infected with A platys may assume a more
activated state than do healthy platelets [57]. Coinfection with A platys and
E canis

, which might use the same tick vector, have been documented [11–

13]. As expected, A platys does not share serologic cross-reactivity with
E canis

[11,13]. Veterinarians should consider speciﬁc testing for A platys

infection in dogs with repeatable evidence of thrombocytopenia for which
another cause (including the more common ehrlichial infections) cannot be
documented.

Neorickettsia risticii (formerly Ehrlichia risticii)

Neorickettsia risticii

, the causative agent of Potomac horse fever, can

infect dogs and cats as well as horses. Like the other Neorickettsia, this agent
is not transmitted by the bite of a tick. Only very recently has the complex
life cycle of this organism begun to be understood [58]. Similar to other
Neorickettsia

, trematodes that use snails as intermediate hosts seem to be

key. Infection could be passed orally during the ingestion of snails, free
trematode life-stages, or through ingestion of aquatic insects with encysted
metacercaria [58]. Grazing and drinking from standing water seems may be
an ideal way to facilitate infection and may explain why infection of dogs or
cats is less common than infection of horses. N risticii displays tropism for
mononuclear cells and entrocytes, explaining the common presentation of
acute colitis in horses [2]. When dogs are infected, lethargy, vomiting,
bleeding disorders, and arthralgia have been reported [59]. Because N risticii
is actually a member of the genus Neorickettsia rather than Ehrlichia,
E canis

antibody cross-reactivity is lacking, and dogs with this infection

likely will be E canis titer negative [59].

Neorickettsia helminthoeca

The vector for infection is the trematode Nanophyetus salminocola, but

the trematode itself requires three separate hosts (snail, ﬁsh, and bird or
mammal) for completion of its life cycle [60]. The snail intermediate host,
Oxytrema silicula

, is restricted to the northwestern United States. For this

reason, N helminthoeca causes disease only in dogs in this locale. The

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moniker salmon poisoning disease derives from the method of exposure.
Dogs develop infection after consuming raw ﬁsh (often salmon) or
salamanders that are infected with the trematode, which in turn carries
the Neorickettsia organism. N helminthoeca replicates in intestinal epithelial
cells and in mononuclear cells. The clinical disease picture is characterized
by an acute onset of fever and gastrointestinal [GI] illness similar in
presentation to canine parvovirus. Typical laboratory ﬁndings include
thrombocytopenia, lymphopenia and eosinophilia, hypoalbuminemia, and
elevated ALP [61]. Although fecal exam for trematode eggs becomes
positive at about the same time the dog develops illness, trematodiasis can
occur in the absence of neorickettsial infection. Although the disease is
rapidly progressive and fatal if left untreated, it responds readily to the same
antimicrobial therapy as other types of ‘‘ehrlichial’’ infection. Antimicro-
bials are combined with supportive care and therapy directed at removing
the trematode parasite [60].

Feline

‘‘ehrlichiosis’’

Less is known about the signiﬁcance of ehrlichiosis in cats than in dogs,

but monocytic and granulocytic ‘‘ehrlichial’’ infections have been docu-
mented [3]. Experimentally, cats can be infected with N risticii. Neorickettsia
have a tropism for intestinal epithelial cells and mononuclear cells and often
cause diarrhea in susceptible species. Some cats infected with N risticii
developed a diarrheal disease, but most infected cats displayed only mild or
inapparent illness [62]. Cats also have been infected with A phagocytophila
experimentally. Although granulocytic morulae were documented, illness
was again quite mild [48].

Naturally occurring feline ‘‘ehrlichiosis’’ has been documented in only

a small number of cats presented with a variety of clinical signs including
fever, anorexia, arthropathy, GI signs, and general malaise [3,50,63–65].
These infections were conﬁrmed in a variety of ways including identiﬁcation
of morulae, serologic evidence of exposure, PCR identiﬁcation of
organisms, and transfer of infection. The organisms responsible for infection
have been described as resembling E canis [65], N risticii [63], or A
phagocytophila

[50], or infections simply have been described by cell tropism

with no attempt to speciate infection [64]. Thus far, infection of cats with
E chaﬀeensis

, E ewingii, E platys, or N helminthoeca has not been described.

Recently, three cats presented for polyarthropathy or hematologic dys-
crasias accompanied by a positive antinuclear antibody titer were proven by
PCR assay to harbor an organism very similar to E canis, but no serologic
evidence of infection was documented [66]. This leaves in question how
useful serologic methods may be for the diagnosis of ehrlichial infection in
cats. For now, diagnosis relies on ruling out other causes for the described
clinical illness in combination with either identiﬁcation of morulae within
the peripheral blood cells, serologic evidence of exposure to these agents or

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PCR demonstration of DNA, and resolution of clinical signs after
appropriate antimicrobial therapy [3]. Because of the similarities in clinical
presentation between immunologically mediated disease and ehrlichial
infection, cats with such presentations should be evaluated for ehrlichiosis
before initiating immunosuppressive therapy.

Diagnosis

There is no single method of diagnosis for these diseases. Instead, the

diagnosis is achieved to varying degrees of certainty through a combination
of clinical and hematological indicators, serologic evidence, and molecular
conﬁrmation.

Clinical, hematologic, and culture evidence

Ehrlichial infections can mimic several other infectious and noninfectious

conditions, and clinical presentation varies tremendously. The most
common clinical ﬁndings are nonspeciﬁc (lethargy, anorexia, malaise, or
fever) [1]. Ehrlichial disease is included in the diﬀerential diagnosis for
ﬁndings such as acute onset polyarthropathy or bleeding diathesis, but these
ﬁndings are neither speciﬁc nor consistent. Other ﬁndings compatible with
clinical disease caused by each agent have been described previously. The
most common ﬁndings on routine laboratory evaluation are likewise
nonspeciﬁc but combined with clinical presentation may be suggestive.
These ﬁndings include thrombocytopenia, hyperproteinemia and hyper-
globulinemia, mild nonregenerative anemia, and mild hypoalbuminemia
[67]. Culture of intracellular organisms is diﬃcult and expensive and is used
primarily in a research setting rather than for clinical disease diagnosis.
Occasionally, microscopic inspection of blood smears reveals intracyto-
plasmic inclusions (ie, morulae), thus providing a diagnosis of ehrlichial
disease [16]. Cellular tropism of the organism governs which type of cell
(mononuclear, granulocytic, or thrombocytic) will contain morulae (Table 1).
Morulae have been identiﬁed in leukocytes from peripheral blood,
cerebrospinal ﬂuid (CSF), and joint ﬂuid, and the author has identiﬁed
granulocytic morulae in ﬂuid obtained from prostatic wash (Fig. 2)
[28,29,44,45]. The microscopic search for morulae tends to be more pro-
ductive in acute infection. Perhaps this explains why morulae are more
readily apparent in association with granulocytic infections that tend to be
acute in presentation. [16,24,43,47]. The use of concentration techniques,
such as buﬀy coat exam with a Romanovsky-type stain, maximize the
chance of identifying morulae [38].

Salmon poisoning disease is the exception to the rule for these diseases,

with a classic presentation in a restricted geographic locale such that
a presumptive diagnosis is based on typical clinical and routine laboratory
ﬁndings. Antimicrobial therapy should begin immediately in dogs with an

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appropriate presentation and historical exposure, but disease with a similar
presentation (ie, parvovirus) should be ruled out. Cytologic examination of
lymph node aspirates can reveal morulae within mononuclear cells, but
because of the rapid disease course and associated high morbidity and
mortality, therapy should proceed even when inclusions are not evident.
Because of the inherent time delay, serologic tests are useful only to conﬁrm
infection rather than to initiate treatment decisions.

Serologic evidence

Serologic techniques are the most commonly employed conﬁrmatory

diagnostic tests for suspected ehrlichial infection. These tests do not detect
organisms but rather reactive antibody. Veterinarians employing these tests
must understand that a positive titer in a dog from an endemic area does not
conﬁrm that the illness under investigation is caused by ehrlichial infection.
Rather, a positive titer conﬁrms exposure to the organisms, but may be
observed after exposure and clearance of the organism, during the
subclinical stage of infection, or after successful treatment of infection, as
well as during active infection. Likewise, a negative titer does not rule out
infection because of the delay inherent in mounting an antibody response
to acute infection, and because moribund animals may cease to produce
antibody [68,69]. Essentially, positive serologic test results can be viewed as
circumstantial evidence of infection.

Although a number of serologic tests and techniques are available, the

most commonly employed are the indirect immunoﬂuorescence (IFA) and
enzyme-linked immunosorbent assays (ELISA) tests. Traditionally, the IFA
has been the serologic test of choice for ehrlichiosis. Unfortunately, there is
no standardization between laboratories as to the antigen used in a given

Fig. 2. A round, basophilic morulae of Ehrlichia ewingii is seen in the cytoplasm of the neutrophil.
(Adapted from Preziosi DE, Cohn LA. The increasingly complicated story of Ehrlichia.
Compendium on Continuing Education for the Practicing Veterinarian 2002;24(4):277–288; with
permission.)

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test, standard use of antibody (poly or monoclonal), or even a uniform
reporting system. In fact, diﬀerent results can be documented from the same
serum sample sent to diﬀerent laboratories [67]. Antibody generated to one
species of Ehrlichia may or may not cross react with other species. Species
that fall within the same genus possess the strongest serologic cross-
reactivity [68]. Thus it is likely that E canis titers would be positive in a dog
infected with E chaﬀeensis, but might be positive or negative in a dog
infected with A phagocytophila. It seems that although the organisms are
related, only about half the dogs with E ewingii infection are found to be
serologically positive by E canis titer [70]. Perhaps this discrepancy is related
to the acute nature of E ewingii infection in that dogs may not have had
suﬃcient time to mount a measurable cross-reactive antibody response when
clinical illness is observed. It is expected that the strongest serologic reaction
would occur in response to the agent actually causing infection, but the
veterinarian must request that the laboratory run titers other than those to
E canis

(eg, A platys, A phagocytophila, or N risticii) [68]. Of additional

concern, antigenic diversity can occur among organisms of the same species.
Thus, an IFA test may not detect antibody to a strain of the same species
found in another part of the world [68].

Recently, in-house screening tests using ELISA technology have been

marketed [71,72]. The one most frequently used in the United States is
a combination test for E canis and B burgdorferi antibody and heartworm
antigen (Snap 3Dx assay, IDEXX Laboratories, Inc., Westbrook, ME).
This test kit contains horse radish peroxidase-conjugated peptides (two
speciﬁc E canis proteins are used as antigen source), a membrane ﬁlter, and
substrate. Sera from the test animal are added, and antigen-antibody
complex formation in the positive test is indicated by a color change. The kit
is designed to indicate positive results corresponding to an IFA titer of
greater than 1:100, with a manufacturer reported sensitivity and speciﬁcity
of 95% and greater than 99%, respectively. Independent evaluation of the
test using serum with IFA titers above and below the 1:500 level found
a good correlation between the ELISA and IFA, with sensitivity of 71% and
speciﬁcity of 100%. In most cases in which there was disagreement between
tests IFA titers were 1:320 or less [72]. As with any test, predictive value is
more important than sensitivity or speciﬁcity, but it depends on both, and
on disease prevalence. Thus, a positive screening test in an endemic area is
more likely to be a true positive than the same result in an area with a lesser
disease prevalence. Like the IFA test, a positive result must be interpreted
with caution, as it may represent current infection, resolved infection, or
merely exposure.

Molecular diagnosis

Polymerase chain reaction (PCR) uses primers (small segments of

nucleotides that match conserved areas of the organism’s DNA) to amplify

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a portion of the organism’s DNA. Thus, in a laboratory with excellent
quality control, a positive PCR proves that the organism is present in the
test subject. Primers used to detect ehrlichial organisms may be generic or
species-speciﬁc. Generic primers detect many of the organisms in related
genus groups, while speciﬁc primers designed to amplify highly variable
portions of the genome can be chosen to identify only a particular species of
organism [8]. Commercial laboratories usually use generic primers. A
positive test may prompt the use of more speciﬁc primers. The choice of test
sample will inﬂuence the results of PCR. Generally, whole blood samples in
EDTA are submitted for testing but these organisms, and particularly
E canis

, may be sequestered in low numbers in tissue reticuloendothelial

cells [73]. In such a case, PCR could be negative despite the presence of
sequestered organism. Because a positive result depends on organism being
found in peripheral blood, samples should be collected before treatment.
When used to evaluate eﬃcacy of treatment, sample collection should be
postponed at least 2 to 3 weeks after completion of antimicrobial therapy.
The consensus statement of the American College of Veterinary Internal
Medicine (ACVIM) Infectious Disease Study Group recommends that PCR
should be used in conjunction with serology [74].

Treatment

Tetracycline-related antibiotics have been the treatment of choice for

ehrlichial infections for years [38]. Excellent absorption and an infrequent
dosing interval make doxycycline or minocycline the preferred drugs
[33,74,75]. Although there is no clear answer as to how long is long enough
for treatment, the recent consensus of the ACVIM Infectious Disease Study
group suggests using doxycycline at a dose of 10 mg/kg by mouth every
24 hours for 28 days in infected dogs and cats [74]. Although most studies
have been directed at E canis infections, doxycycline has proven eﬀective
for other ehrlichial species and canine infections with Anaplasma and
Neorickettsia

An alternative treatment for ehrlichiosis that has been used for decades

outside of the United States, but only within the last several years in the
United States, is imidocarb dipropionate, an aromatic diamidine also used
to treat babesiosis. Although a single 1980 study found that imidocarb
produced more eﬀective clearance of E canis than did a 2-week course of
tetracycline, recent studies using doxycycline have documented no diﬀerence
in clinical response of dogs treated with either drug alone, or with both
drugs simultaneously [76,77]. Imidocarb is administered as an intramuscular
injection of 5 mg/kg, with a second injection 2 weeks after the ﬁrst [14,77].
Pretreatment with atropine may lessen the occurrence of unpleasant
anticholinergic adverse eﬀects, including salivation, serous nasal discharge,
diarrhea, and dyspnea.

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Other antimicrobial agents have been evaluated for the treatment of

ehrlichiosis. Chloramphenicol has been used in puppies to avoid dental
discoloration associated with tetracycline, but doxycycline is less likely to
discolor erupting teeth than tetracycline [14]. Although enroﬂoxacin has
been used successfully to treat experimental Rickettsia rickettsii infection in
dogs, it does not seem to be eﬀective for E canis [33]. Similarly, ciproﬂoxacin
was unsuccessful in eliminating infection with E chaﬀeensis in people [78].
Thus, quinolones cannot be recommended as treatment for ehrlichial
infection.

Supportive therapies may be required for treating ehrlichial infection.

Glucocorticosteroids can attenuate the immune-mediated destruction of
platelets associated with infection, and thus a short course (2 to 7 days) of
prednisone may be indicated if thrombocytopenia is severe or pending
diagnostic testing to diﬀerentiate ehrlichiosis from immune-mediated throm-
bocytopenia [14,24]. Steroids also may be indicated for the treatment
of polyarthritis, vasculitis, or meningitis associated with certain types of
ehrlichial infections [14,44,45]. Other supportive treatments might include
the administration of parenteral crystalloid or colloidal ﬂuids, or blood
transfusion, as indicated [14,38]. Complications of chronic E canis infection
such as glomerulonephritis or pancytopenia likewise may require speciﬁc
supportive therapy.

Monitoring treatment eﬃcacy

Evaluation of treatment eﬃcacy is problematic. Resolution of clinical

signs and normalization of platelet counts usually are noted within days
after initiating proper treatment for acute cases, and often for mild chronic
cases also [14,24,77]. Even for chronic cases, if clinical signs and
hematologic parameters fail to improve within 1 or 2 weeks from initiation
of therapy, the diagnosis should be re-evaluated. Such a scenario could
result simply from severe, chronic infection, but it also might be associated
with coinfection with another organism (ie, babesiosis) or concurrent
noninfectious illness. Even after rapid clinical improvement, studies have
found that platelet counts may decrease after completion of doxycycline
therapy, titers remain elevated, organisms can be cultured, and PCR results
remain positive [73,77].

In most dogs, serum antibody titers decline and become negative within 6

to 9 months after therapy. Despite clinical and hematological normalization,
however, serum antibody titers remain elevated in other dogs for months to
years after appropriate treatment [31,32,73,77]. Persistently positive titers
can indicate continued infection, reinfection, or simply may be indicative of
a past infection [24,31]. Because ELISA Snap tests provide only a positive or
negative result, the utility of this test to monitor for serologic evidence of
cure is limited. The PCR test may oﬀer the best option for documenting

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clearance of the organisms after therapy, but even PCR evaluation can be
problematic [77]. For instance, PCR cannot distinguish between living and
dead organisms. Because it is unlikely that killed organisms would persist in
the body for more than several weeks, a positive PCR result weeks after
completing therapy would strongly suggest persistent infection. The greater
problem is false negative tests resulting from sampling tissues (ie, peripheral
blood) containing low levels of organisms [73,79]. Practically speaking,
treated dogs in which the clinical and hematologic evidence of disease
resolves need not be further evaluated for the presence of organisms. In
animals treated appropriately for infection but in which evidence of disease
remains, a positive PCR would warrant continuation or alteration of
therapy.

Prevention and control

Unfortunately, exposure to E canis does not confer protective immunity

against either that organism or any related organism [33,48]. Infection with
N helminthoeca,

however, does result in protective immunity, at least from

very similar strains [60]. Although a vaccine for horses is available for N
risticii

[2], there is no vaccine available for N helminthoeca in dogs or for any

other ehrlichial pathogen. Prevention of these diseases revolves around
minimizing exposure. For N helminthoeca, that means preventing dogs from
consuming raw ﬁsh. To minimize risk of developing N risticii infection, pets
should not be allowed to drink from standing bodies of water. For
prevention of infection with Ehrlichia and Anaplasma species, eﬀective tick
control is paramount. Several highly eﬃcacious products are available for
direct application to the dog, and premises sprays are available to decrease
tick populations in the dog’s local environment.

Other strategies for disease prevention have been considered. The

prophylactic use of tetracycline antibiotics (3 mg/kg doxycycline by mouth
every 24 hours) in endemic regions during tick season has been advocated
for preventing infection. This option is not free of cost or risk associated
with antibiotic use, and theoretically could lead to antimicrobial resistance
[14]. It also has been suggested that dogs be monitored serologically and
positive animals treated regardless of the presence or absence of clinical
signs [14]. It is possible that many untreated animals would remain infected
subclinically and never go on to develop illness. Also, even when using very
sensitive and speciﬁc tests, false-positive results will occur, particularly in
areas of low disease incidence. This could lead to treatment of uninfected
animals with the attendant costs, inconveniences, and potential for adverse
reactions. It seems likely that veterinarians performing screening serology
would test for E canis antibody, and might therefore miss infections with
organisms from the related genus (ie, Anaplasma, or Neorickettsia). If
a veterinarian opts to perform routine screening, positive results should

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elicit further evaluation of the animal. At a minimum, such evaluation
should include a thorough physical examination, quantitative platelet count,
and measurement of serum globulin. Positive screening tests also might
prompt conﬁrmatory tests, including IFA or PCR, before starting antibiotic
therapy.

Zoonotic potential

Because people can be infected with ehrlichial and related organisms, the

zoonotic potential of these infections must be considered. By virtue of the
need for a vector host, there is no evidence that any of these infections are
passed directly from animals to people. Because pets are susceptible to
infection with some of the same organisms that people are, pets might serve
as disease sentinels, or perhaps even as reservoirs of infection [74]. Human
monocytic ehrlichiosis (HME), ﬁrst reported in the United States in 1986
[80], originally was thought to be caused by E canis. Ultimately, the agent of
HME was proven to be the closely related species E chaﬀeensis. Like E canis,
E chaﬀeensis

has a monocytic cell tropism, and as expected by their close

genetic relationship, strongly cross-reactive antibodies are produced [40].
Although primarily a human pathogen, persistent E chaﬀeensis infection of
dogs has been documented in experimental and natural settings [8,41]. These
long-lasting infections raise the concern that dogs could serve as a reservoir
of infection for human disease, although such a phenomenon never has been
proven [40]. Several years after the recognition of HME, a granulocytic form
of human ‘‘ehrlichiosis’’ (HGE) was recognized [49]. The causative agent of
HGE was shown to be nearly identical to E phagocytophila (known to infect
small ruminants) and E equi (known to infect horses and dogs). In fact, the
organisms are classiﬁed as one and the same and have been removed from
the genus Ehrlichia and are known as A phagocytophila. As for E chaﬀeensis,
the fact that dogs harbor these organisms must raise the possibility that dogs
could serve as disease sentinel or as reservoir host [39,47,81,82]. Thus far,
there is little evidence that A phagocytophila causes persistent infections in
dogs, making their role as a reservoir unlikely.

Summary

Ehrlichiosis is a term that has been used to describe infection with any of

a number of related intracellular, vector-borne pathogens. A recent
reclassiﬁcation has resulted in the transfer of several species previously
known as Ehrlichia to the genus Anaplasma or Neorickettsia. Ehrlichia and
Anaplasma

are transmitted largely through the bite of infected ticks, while

vectors for Neorickettsia include trematodes and the intermediate hosts
(ie, ﬁsh, snails, and insects) involved in the trematode life cycle. Dogs (and
cats) are susceptible to infection with several of these pathogens, and

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veterinarians should be aware of the similarities and diﬀerences between
E canis

and related infections. Pets with suggestive clinical signs and

laboratory abnormalities may be started on doxycycline pending speciﬁc
diagnostic testing. The veterinarian practicing in endemic areas must
understand the implications and limitations of serologic and molecular
testing to conﬁrm a diagnosis. For animals in endemic areas, prevention of
exposure to vectors can lessen the risk of disease for pets and might lessen
the potential for animals to become carriers of disease for their human
companions.

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Canine babesiosis

A. Lindsay Boozer, DVM*,

Douglass K. Macintire, DVM, MS

Department of Clinical Sciences, Auburn University College of Veterinary Medicine,

Wire Road, Auburn, AL 36849,

Canine babesiosis is a protozoal, tickborne, hemolytic disease with

worldwide distribution and global signiﬁcance. Over 100 Babesia species
have been identiﬁed, but only Babesia canis and B gibsoni have been shown
to infect dogs [1]. Until recently, Babesia organisms infecting dogs were
identiﬁed based on morphologic appearance. All large Babesia were
designated B canis, whereas all small Babesia were thought to be B gibsoni.
Molecular analysis and serologic surveys have shown these organisms to be
much more prevalent and genotypically diverse than previously recognized.
Genetic sequencing has revealed that there are at least three distinct
subtypes of small Babesia aﬀecting dogs [2–4]. In the United States, there are
two clinically distinct strains of small Babesia. In California, a small Babesia
closely related to Theileria infects a variety of dogs and causes relapsing
parasitemia [2,5]. In the remainder of the United States, the classic Asian B
gibsoni

infects almost exclusively pit bull-type dogs and is often subclinical

[6–8]. A third strain of small piroplasm has been identiﬁed recently in
Europe and is most similar to a human and rodent pathogen, B microti [9].

Babesiosis is considered an emerging disease, as numerous cases are being

reported in new areas throughout the United States and Europe. Trans-
mission of canine babesiosis is facilitated by the international and interstate
transportation of dogs and the availability of tick vectors. The presence of
chronic subclinical carrier states, the inability to completely eradicate all
infections, and transovarial transmission in the tick encourage the es-
tablishment of infected tick populations. Babesiosis also has gained atten-
tion as a persistent endemic disease in greyhound kennels [10,11]; a
newly recognized problem in pit bulls [6–8], as a potential model for falci-
parum malaria in people [12,13], and as an emerging zoonosis [14].

Vet Clin Small Anim

33 (2003) 885–904

* Corresponding author.
E-mail address:

boozeal@vetmed.auburn.edu (A.L. Boozer).

0195-5616/03/$ - see front matter

doi:10.1016/S0195-5616(03)00039-1

The organism

Babesia

species (Fig. 1) (Fig. 2) generally are divided into large and small

organisms. B canis is a larger piroplasm (3 to 5 lm) that is approximately
twice the size of B gibsoni (0.5 to 2.5 lm) [2]. B canis organisms are pyriform
and often occur in pairs. B gibsoni organisms appear in ﬁve to six diﬀerent
shapes and are frequently oval, often single, and may display a ring form
[15]. Small Babesia are found occasionally as a maltase cross-form but are
not reported pairs [5,7,16,17].

Babesia canis

Three distinct subspecies of B canis exist: canis, rossi, and vogeli. The

subspecies demonstrate tremendous variation in clinical signs, geographic
distribution, and infective tick vectors. Immune responses are fairly speciﬁc,
and little cross-protection occurs among the diﬀerent subspecies. Genetic
analysis of rRNA sequences has revealed that B canis canis and B canis
vogeli

are most similar to each other, with 82% identity in nucleotide

positions. B canis rossi is approximately 70% homologous to the other two
subspecies and tends to cause the most severe disease [18]. In North
Carolina, a fourth large canine piroplasm was identiﬁed in a Labrador re-
triever undergoing chemotherapy for lymphosarcoma [19]. This piroplasm
was morphologically identical to B canis; however, polymerase chain re-
action (PCR) and antibody tests for all known Babesia species were
negative. The 18S rRNA gene fragments showed considerable diﬀerences
to B canis and B gibsoni, and this organism probably represents a fourth
species/subspecies of Babesia [19].

Fig. 1. Babesia gibsoni organisms are seen in the red blood cells of a dog.

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A.L. Boozer, D.K. Macintire / Vet Clin Small Anim 33 (2003) 885–904

South African babesiosis is caused by B canis rossi and is transmitted by

the Haemophysalis tick. This strain is very widespread and notoriously
the most virulent. At the Onderstepoort Veterinary Academic Hospital
(OVAH) in South Africa, 12% of sick patients were diagnosed with
babesiosis. Approximately a third of patients were hospitalized, and fa-
talities were common, despite treatment [20].

In Europe and Asia, babesiosis is caused by B c canis, and Dermacentor

reticularis

is the tick vector. In endemic areas of France, up to 85% of over

500 dogs tested had antibodies to Babesia. A maximum of 14% displayed
any clinical signs, however, and mortality was only 1.5% [21].When deaths
occurred, they were attributed to hepatic and renal damage [21].

Babesia canis vogeli

is transmitted by the brown dog tick, R sanguineus,

and causes relatively mild disease in the United States and in tropical and
subtropical areas. B canis ﬁrst was reported in the United States in 1934 [22].
It is recognized throughout the United States, and the disease has been
endemic in southeastern greyhound kennels for over 50 years. Babesiosis in
greyhounds primarily causes anemic pups, which may be parasitemic but
seronegative [10]. Vertical transmission is suspected, and B canis has been
documented in pups under 2 days old [23]. The incubation period of natural
B canis

infection is approximately 10 to 21 days [24].

In a Mississippi kennel, 59% of greyhounds screened in the early 1980s

had positive indirect ﬂuorescent antibody (IFA) titers. Organisms were not
evident on blood smears. The kennel routinely admitted numerous dogs
without quarantine, and additional screening of dogs from source kennels in
Mississippi, West Virginia, Oklahoma, Texas, and Florida revealed enzootic
disease with seroprevalence rates of well over 50% [10]. More recently, 46%
of almost 400 greyhounds of various ages screened in Florida were
seropositive (greater than 1:80) for B canis antibodies [11]. In kennels with
a history of anemic pups, 78.5% of the dogs were seropositive. In kennels

Fig. 2. Babesia canis usually appears as paired piriform organisms in canine red blood cells.

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without a history of anemic pups, 23% were seropositive. None of the dogs
currently racing were seropositive, suggesting reduced performance in
infected dogs [11]. Babesiosis in greyhounds often is attributed to genetic
susceptibility and environmental exposure to numerous ticks. Although
most common in tick-infested kennels, B canis occasionally aﬀects the
general canine population also. In North Carolina, 3.8% of random source
dogs were seropositive for B canis antibodies [25], and in California shelters,
13% of dogs were seropositive for B canis [26].

Babesia canis

has the potential to become a more prevalent disease. The

growing eﬀort to rescue and adopt greyhounds may inadvertently be con-
tributing to the perpetuation of a B canis reservoir in North America. In
1995, over 16,000 greyhounds were rescued and adopted [23]. It has been
estimated that 20% to 60% of these dogs may be seropositive [23]. In reality,
the probability of clinical disease in an adult greyhound dog is low, and
signiﬁcant spread is unlikely [23]. Age-related immunity is an important
protective factor. B canis organisms isolated from anemic pups did not cause
clinical disease, even in splenectomized adults [10]. Disease transmission is
most likely to occur if infected greyhounds are used as blood donors or are
kenneled with breeding animals and young puppies.

Babesia gibsoni

Unlike B canis, documented B gibsoni infection is relatively new in the

United States. B gibsoni ﬁrst was recognized in India in 1910 [27] and since
has been reported in Asia, Northern and Eastern Africa, Brazil, and rarely
in Europe [9,28]. In the United States and Europe, emergence of B gibsoni
infection has been dramatic. In 1968, B gibsoni ﬁrst was reported in the
United States in a chronically infected dog from Malaysia [29]. In Malaysia,
B gibsoni

is endemic, and 18% of dogs demonstrated organisms on blood

smear evaluation [30]. Dog ﬁghting is legal in Malaysia, and the suspected
international transport of dogs to and from this country may be an im-
portant source of small Babesia in the United States.

In 1979, small babesial infection was reported in a dog from Connecticut

with no travel history [31]. Since that time, infections have been documented
in dogs from California [5], North Carolina [7], Indiana [32], Oklahoma [33],
and Alabama [6]. At the Vector-Borne Disease Testing Laboratory at
the North Carolina State College of Veterinary Medicine, a PCR-based
test has identiﬁed infected dogs in Arizona, California, Florida, Georgia,
Kentucky, Maryland, Michigan, Minnesota, Mississippi, Missouri, New
Jersey, New York, Ohio, Pennsylvania, South Carolina, Tennessee, Texas,
Washington, Wisconsin, and Virginia (Birkenheuer et al, unpublished data)
B gibsoni

also has been reported occasionally in areas near military bases,

where international working dogs are housed [23]. In Japan, B gibsoni is the
most common species, except in Okinawa, where B canis is also prevalent
[34]. Approximately 500 dogs are shipped annually from Okinawa to the

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United States, and many of these dogs may be infected with B canis or
B gibsoni

Molecular analysis has led to a much broader understanding of the

phylogenetic relationships and classiﬁcation of the canine piroplasms.
Historically, only one small Babesia was thought to infect dogs. Al-
though morphologically identical, analysis of the 18S rRNA has shown
that there are actually three distinct isolates of small Babesia [2]. Small
Babesia

infections now are attributed to the Asian isolate (B gibsoni), the

Californian isolate, and a recently identiﬁed B microti-like organism in
Europe. The Asian isolate is considered Babesia species sensu stricto and
is identical to the strain identiﬁed in most of the United States, excluding
California [2,4].

Babesia gibsoni

organisms isolated from a dog in Oklahoma were

sequenced and compared with the Californian small Babesia [33]. The
Oklahoma isolate demonstrated 232 nucleotide diﬀerences in the 18S rRNA
gene when compared with the Californian organism [33]. The Californian
small Babesia is related closely to Theileria species and to Babesia isolates
that infect wildlife and people in the Western United States [2]. T equi
was most similar to the Californian strain with 92.7% identity [4]. The
Californian organism showed only 88.2% sequence identity to B gibsoni [4].
There is also a suggestion, based on maximum-likelihood analysis, that the
Californian piroplasm and Cytauxzoon felis are ancestral to the re-
maining Theileria and to the classic B gibsoni [35].

Babesia gibsoni

appears to be less virulent than the Californian strain of

small Babesia. Parasitemia, relapse rates, and overall mortality appear to be
lower with B gibsoni infection in the southeast and Midwest. Breed dif-
ferences also exist. B gibsoni aﬀects almost exclusively pit bulls and
American Staﬀordshire terriers in the United States. The disease has been
reported only sporadically in other breeds. In North Carolina, a mixed
breed dog with concurrent illness was naturally infected with B gibsoni [7],
and in Indiana, a mixed-breed dog was infected with B gibsoni after being
attacked by pit bulls and traveling to Florida [32]. The Californian strain
has been reported in many breeds of dogs.

In 1994, a new small piroplasm was discovered in a German dog that

presumably acquired the infection in Spain [9]. This organism appears to be
related only distantly to B gibsoni. The organism shows the most similarity to
B microti, B rodhainii

, and Theileria equi [9]. Previous reports of small Babesia

in Europe have been relatively scarce. The investigators who discovered the
parasite have suggested the name T annae for this organism [9].

Over 150 dogs in northwest Spain have been identiﬁed with a small

piroplasm (generally less than 2 lm) on blood smear analysis since the
discovery of this organism. Ribosomal DNA analysis conﬁrmed that the
organism is 100% identical to T annae [36]. Parasitemia was subjectively
rated as low in 149 dogs and moderate or severe in eight dogs. Ninety
percent of dogs had a hematocrit less than 31%, and regeneration was

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prominent. Leukocytosis was not common, and thrombocytopenia was
present in about half of animals [36]. Marginal azotemia was very common.
Approximately 10% of dogs had evidence of renal failure on serum
chemistry analysis. Hepatic involvement appears minimal with T annae, and
75% of dogs had normal hepatic serum chemistry values. The Californian
organism is reported to cause signiﬁcant hepatic changes [37]. Elevations in
creatine kinase (CK) were fairly common and may be a result of muscle
damage seen with the sequestration of parasites.

Some Babesia species may be reclassiﬁed as Theileria. Theileria organisms

diﬀer from Babesia in that they have a pre-erythrocytic life stage in a
lymphocyte, and they lack transovarial transmission in the tick [38]. The
identiﬁcation of a lymphocytic stage in B equi resulted in its reclassiﬁcation
as T equi. A lymphocyte stage also has been discovered recently in B microti,
and it likely also will be reclassiﬁed as a Theileria.

The apparent incidence of B gibsoni has been rapidly increasing, since its

ﬁrst in the United States. The additional awareness among practitioners, pit
bull and greyhound owners, and the contribution of sensitive PCR testing
have led to an increase in the diagnosis of this new disease. More stringent
regulations on the importation of dogs from endemic areas are needed to
prevent further spread of this disease.

Pathogenesis

Dogs become infected with Babesia after the bite of an infective tick

releases sporozoites into the circulation. Inside the host, the organisms
attach to the red cell membrane and are engulfed by endocytosis [39].
The membrane disintegrates, and the parasites remain directly inside the
cytoplasm. Babesia species undergo binary ﬁssion, resulting in merozoites.
Ticks become infected with merozoites during feeding and may remain
infective for several generations. Schizogony produces merozoites in the
salivary glands, gastrointestinal (GI) cells, and oocytes of the tick. With B
canis

, ticks must feed for 2 to 3 days for transmission to be complete [40].

The incubation period of B gibsoni is reportedly 7 to 21 days, and for B
canis

, it is 10 to 21 days. Vertical transmission, blood-borne transmission,

and possible transmission through bite wounds is suspected for B gibsoni
and B canis. B canis has been documented in pups that were 36 hours old
[23], and B gibsoni has been found in 3-day-old puppies and a dam [41].

The pathogenesis of canine babesiosis varies with the infective species and

is proving more complex than initially recognized. Many of the pathogenic
mechanisms are actually a result of host immune response to the organism
rather than direct destruction of the erythrocyte by the parasite. Review of
662 cases of South African canine babesiosis suggested two basic disease
mechanisms: hemolytic anemia and a hypotensive shock syndrome induced
by inﬂammatory mediators. Disease in the severely anemic group was

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attributed primarily to immune-mediated hemolytic anemia, and nonanemic
babesiosis was attributed to an overwhelming immune response to the
parasite.

Hemolytic anemia is a prominent feature of large and small babesiosis in

dogs. Direct red blood cell (RBC) damage, intravascular hemolysis, and
extravascular hemolysis are thought to occur [42,43]. Antierythrocyte
antibodies, IgG-bound erythrocytes, erythrocyte oxidation, phagocytosis,
osmotic fragility, and a hemolytic factor in serum have been implicated in
B gibsoni

-related RBC destruction [43–45].

In B gibsoni infections, parasitemia rarely exceeds 10%, yet severe anemia

frequently develops [44,46]. In experimental B gibsoni infections, dogs ex-
perienced concurrent increases in parasitemia and decreases in hematocrit
approximately 2 weeks postinfection. This may be caused by destruction by
escaping parasites [44]. Spherocytes and autoagglutination were observed
approximately 3 to 4 weeks postinfection, suggesting immune-mediated
disease is also a signiﬁcant factor. Anti-RBC IgG has been identiﬁed in
B gibsoni

-infected dogs with ELISA testing. The antibody levels are inversely

related to the degree of anemia [47].

Complement dependent immune-mediated lysis and removal by the

mononuclear phagocytic system are also important factors. Hemoglobin-
uria and diﬀuse erythrophagocytosis and hemosiderosis suggest intravas-
cular and extravascular hemolysis [37]. It also has been suggested that
a ‘‘hemolytic factor’’ is present in the serum of dogs infected with B gibsoni
[46]. Sera from B gibsoni-infected dogs caused in vitro hemolysis of normal
beagle RBCs. The relative activity of the hemolytic factor correlates well
with the degree of parasitemia and anemia [48]. Soluble antigens also are
thought to be involved in B canis anemia. Over 40% of complicated
B canis rossi

infections demonstrated autoagglutination [49]. Immune-

mediated disease often causes continued hemolysis after babesiacidal
treatment.

In many cases, the degree of tissue hypoxia is relatively severe for a given

level of anemia [50]. A quantitative and qualitative deﬁcit of hemoglobin can
occur with B canis infections. The hemoglobin that remains in intact cells
may function abnormally at the tissue level, especially under acidic and
hypercapnic conditions [50]. Other Babesia and Plasmodium species are
known to contain enzymes that cleave hemoglobin. Preliminary electro-
phoretic studies with B canis suggest similar enzymes may be present in
canine infections [51]. It is not known whether these mechanisms exist for
B gibsoni

Canine babesiosis has several similarities to malarial infections and may

prove to be a valuable model for studying human disease. In both diseases,
a small subset of victims experience hemoconcentration, shock, neurologic
signs, and multiple organ failure [13,52,53]. Nonanemic babesiosis is
associated with an acute, overwhelming inﬂammatory response mediated
by cytokines, nitric oxide, platelet activating factor, and eicosanoids. The

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syndrome (‘‘red babesia’’ ‘‘red biliary’’) is often peracute and associated
with simultaneous massive hemolysis and increase in vascular permeability
resulting in a hemoconcentrated state. Plasma, rather than a ﬁltrate of
plasma, is lost through the leaky endothelium [49]. Dogs with nonanemic
babesiosis often have severe azotemia, electrolyte and acid–base disturban-
ces, and minimal leukocyte responses or even leukopenia [13]. Leukopenia
may be caused part by sequestration in the pulmonary vasculature. The se-
verity of the inﬂammatory response may be so rapidly fatal that a signiﬁcant
leukocyte bone marrow response and the hemolytic state may not have time
to develop.

Cerebral and cerebellar signs are encountered occasionally in complicated

malarial and babesial infections [54]. Signs may include posterior paresis,
muscle tremors, nystagmus, anisocoria, aggression, paddling, crying, and
altered states of consciousness. In people, parasite-derived surface proteins
serve as endothelial cell receptors. Parasites become sequestered in the
microvasculature of the muscle and central nervous system (CNS) and
concentrate the mediators of the inﬂammatory response in the areas where
they lodge. Parasites also have been visualized in canine cerebral capillaries.
Cerebellar signs may present acutely or may be delayed by several days to
weeks in malaria and babesiosis [54].

Multiple organ dysfunctions may occur in severe cases of hematozoan

infection [55]. Secondary lung injury is one of the most common ﬁndings.
Pulmonary edema may result from increased endothelial permeability as-
sociated with systemic cytokine release, nitric oxide, free oxygen radicals,
eicosanoids and platelet activating factor [49,56,57]. In malarial infections,
approximately 3% to 10% of patients develop acute onset pulmonary
edema relatively late in infection [58]. In dogs, acute respiratory distress
syndrome (ARDS) is suspected when there is hypoxemia, acute dyspnea,
poor response to diuretics, and diﬀuse inﬁltrates in the caudal lung ﬁelds.
Cardiogenic overload and cardiac depression associated with acidemia
should be ruled out. Evidence for cytokine-based disease has been doc-
umented in malarial infections. High levels of tumor necrosis factor
(TNF), interleukin-1 (IL-1), IL-6, IL-10, lymphokines, and neutrophil
products have been found in malarial infections [49,59]. Nitric oxide also
may play an important role in cytokine-mediated disease and TNF-me-
diated hypotension. Attempts to correlate levels of reactive nitric oxide
metabolites with the severity of disease in canine babesiosis have been
discouraging [60].

Acute renal failure is an uncommon but severe complication of virulent

babesiosis and complicated malaria. In some malarial infections, intra-
vascular hemolysis and gross hemoglobinuria (‘‘blackwater fever’’) are
found in association with renal failure. Historically, obstruction of renal
tubules with hemoglobin was thought to cause anuria in malaria [61]. The
role of hemoglobin in development of acute renal failure is debatable.
Hypoxia and cytokine-mediated renal damage have been proposed more

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recently as likely causes of renal damage [62,63]. In particular, TNF has
been correlated with the development of renal failure [64].

In general, hemolytic anemia is the primary manifestation of B gibsoni

infection. B gibsoni infection usually causes acute parasitemia followed
by gradual decline in parasite numbers and development of a chronic
asymptomatic carrier state. Carriers may have spontaneous relapses of
parasitemia. Relapses also have been reported with corticosteroid ad-
ministration, splenectomy, or stress [37,65]. Experimental infections with
B gibsoni

have helped deﬁne the histopathologic changes and mechanisms

of disease of the Oklahoma/Asian strain [66] and the Californian small
Babesia

[37].

Dogs experimentally infected with the Californian isolate showed

signiﬁcant hepatic lesions. Findings included diﬀuse nonsuppurative peri-
portal and centrilobular hepatitis and extramedullary hematopoiesis.
Perivenular ﬁbrosis, Kupﬀer cell hypertrophy, and erythrophagocytosis
also were observed. An increased population of CD3 + lymphocytes were
observed in the liver. Cells also showed an increased expression of ICAM-1,
2, 3 and VCAM 1, and LFA-3 ligands. Activated macrophages were seen
within the central hepatic veins. Similar cells were seen in the closely related
WA1 infection in hamsters and people. Renal lesions included IgM antibody
deposition in inﬂamed arteries and glomeruli [37]. Infected dogs often
maintain extremely high IFA titers, and chronic antigenic stimulation may
lead to sequelae such as the membranoproliferative glomerulonephritis with
IgM deposits that has been demonstrated in experimental infection with the
Californian isolates [37].

Multifocal, segmental, necrotizing arteritis was seen in ﬁve of six dogs in

the GI tract, mesentery, and skeletal muscle. Lymphadenopathy was seen in
all dogs, mainly in the hepatic and peripancreatic nodes. Plasma cells were
prominent in lymph nodes. Splenomegaly was noted in four of four dogs.
Experimental infections in spleen intact animals resulted in less para-
sitemia but more severe anemia than in splenectomized dogs. This most
likely is related to rapid removal by the mononuclear phagocytic system
(MPS). All six dogs had antibody titers and eventually became Coomb’s
positive [37].

Experimental infection with the Oklahoma isolate produced milder signs

of clinical disease. Five dogs (one splenectomized) were inoculated with
organisms from one of two naturally infected dogs in Oklahoma.
Parasitemia was relatively mild and ﬁrst appeared 1 to 5 weeks after
inoculation. At 4 to 6 weeks, parasitemia peaked at 1.9% to 6.0%.
Parasitemia is reportedly higher in naturally occurring Californian in-
fections, which reached parasitemias of 5% to 40% [5]. Clinical signs in-
cluded lethargy, fever, and pale gums. All dogs were anemic and
thrombocytopenic. The thrombocytopenia developed before and persisted
longer than the parasitemia. [66]. Transient severe neutropenia also was
noted in some dogs.

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Clinical signs

The clinical signs of babesiosis can range from subclinical infections to

a hyperacute fulminant fatal disease similar to complicated malarial in-
fections. The clinical course of babesiosis is determined by the particular
strain or subspecies of Babesia, the immune response of the host, the age of
the host, the presence of concurrent infections, and previous exposure to the
organism. Hyperacute, acute, chronic, and subclinical forms of the disease
are known to exist.

Babesia canis

is capable of causing a wide range of clinical signs that may

involve subclinical infection, anemia, thrombocytopenia, lethargy, anorexia,
splenomegaly, hemoglobinuria, bilirubinuria, fever, and jaundice. Severe
cases may be accompanied by acute renal failure, acute respiratory distress
syndrome (ARDS), disseminated intravascular coagulation (DIC), hemo-
concentration (‘‘red biliary’’), icterus, hepatopathy, and a neurologic
syndrome referred to as cerebral babesiosis. Cerebral babesiosis and hemo-
concentration are associated with high mortality rates [12]. Babesial shock
can result from severe anemia or as a result of inﬂammatory mediators
associated with multiple organ dysfunction syndromes and resembling septic
shock [49]. Severe cases also may be complicated by lactic acidosis and
acidemia. Rhabdomyolysis has been described recently as a complication of
B canis rossi

infection in two dogs [67]. One dog exhibited caramel colored

urine, increased serum myoglobin, increased creatine kinase, and acute renal
failure. A second dog exhibited muscle pain, tremors, cerebral babesiosis,
pulmonary edema, and death. Muscle necrosis and hemorrhage were seen
on histopathology [67]. Masseter muscle pain also has been reported. A
disproportionate elevation in serum urea nitrogen (BUN) often is seen in
complicated babesiosis and may be caused by increased muscle catabolism
[67]. In human malaria, rhabdomyolysis is unusual, but asymptomatic
muscle damage is very common. Biochemical evidence of muscle damage is
also common in bovine babesiosis. Rarely reported complications of canine
babesiosis include respiratory disease, diarrhea, vomiting, ascites, edema,
periorbital edema, and hemorrhages.

Babesia gibsoni

Clinical signs of B gibsoni infection are similar to B canis and include

fever, thrombocytopenia, regenerative anemia, splenomegaly, lymphade-
nopathy, and lethargy [66]. Thrombocytopenia is a prominent feature of
small babesial infection and may occur before and after parasitemia [66].
Clinical icterus is uncommon. Mortality was one out of ﬁve in dogs
experimentally infected with an Oklahoma strain of B gibsoni [66]. Two
of nine naturally infected dogs in North Carolina died [7]. Six of 15

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dogs naturally infected with the Californian strain died or were euthanized
because of severe illness [5]. Subclinical B gibsoni was identiﬁed in over half
of 33 pit bulls screened in Alabama [6]. Only 1 of 18 PCR-positive dogs
displayed clinical signs [6]. The mean hematocrit was 31%, and the
mean platelet count also was decreased at 154,000/lL with an increased
mean platelet volume (MPV) [6].

Coinfection

In many cases, babesiosis is complicated with concurrent tickborne

diseases or hemoparasitism. The role of Ehrlichia may be especially im-
portant as a source of immunosuppression. Experimental infection with
Ehrlichia canis

can precipitate clinical babesiosis in B. canis carrier dogs

[68,69] Five of 15 dogs in Greece with babesiosis had another hemoparasitic
infection (Ehrlichia, Hemobartonella, or Leishmania) [70]. In North Carolina
Walker hounds, coinfection was prevalent. E canis, B canis, some Rickettsia,
and possibly Bartonella are transmitted by Rhipicephalus sanguineus [71].
Coinfection with babesiosis and borreliosis (Lyme disease) also has been
reported in people. The diseases share a common Ixodes tick vector, and
coinfection may explain poor response to single-agent therapy.

Immune suppression may be induced by B gibsoni infection [72], resulting

in an increased likelihood of concurrent infection and clinical relapse. Dogs
with subclinical B gibsoni infection demonstrated suppressed lymphocyte
blastogenesis. Dogs suﬀering from clinical relapses also had depressed
lymphocyte blastogenesis and decreased production of antiparasite anti-
body [72].

Diagnosis/clinical pathology

Diagnosis of B canis and B gibsoni often is made by identifying organisms

on blood smears. Organisms may be visualized better with Giemsa or Field
stains than with quick stains [5,73]. Blood smears should be made im-
mediately, since storage, even with refrigeration, may make organisms
impossible to identify [73]. Diagnosis may be complicated, because clinically
aﬀected animals do not always have organisms visible on blood smears, and
low levels of parasitemia are common even with patent infections. Ex-
amining buﬀy coat smears or blood smears made from capillary blood may
increase parasite detection. In chronic or subclinical infections, organisms
may be in such low numbers that it is diﬃcult to ﬁnd the organisms.

Percoll gradients also have been used in an attempt to improve diagnostic

sensitivity [74]. Erythrocytes from Italian dogs with B canis were con-
centrated with discontinuous gradients of Percoll. Upper layers had higher
parasitemia, because cells were larger and had lower speciﬁc weight [74].
Percoll gradient smears had greater than 134 times the concentration of

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parasites than the center of a normal blood smear and greater than 30 times
the number of parasites seen at the periphery of a blood smear.

Serologic testing of IFA titers often is used, but this may have limitations

in endemic areas, suﬀer from cross reactivity, and be negative in very young
animals or early in infection [7,8,75]. Antibodies usually take 8 to10 days to
develop. ELISA and complement ﬁxation tests are also available but are
used less commonly. In a large survey of stray dogs and kenneled pit bulls in
North Carolina, IFA titers for B gibsoni, the Californian isolate, and B canis
were performed. Microscopic examination and PCR analysis also were per-
formed in some of the dogs. Several dogs that were PCR positive did not have
positive IFA titers, and in some cases the PCR result was not in agreement
with the highest titer for a given species. The clinical signiﬁcance of dogs
that are seropositive but PCR and blood smear negative is problematic. It
is possible that animals are infected persistently with levels of parasitemia
that are below the detection of microscopic and PCR analysis [8].

A nested PCR to amplify Babesia small subunit ribosomal RNA is

extremely sensitive and able to detect a parasitemia of 0.0001% [34]. A
seminested PCR test oﬀered through North Carolina State is also extremely
sensitive, capable of detecting 50 organisms/mL, and it can diﬀerentiate B
gibsoni, B canis vogeli, B c rossi

, and B c canis [76]. Seminested PCR test-

ing can detect parasitemia levels that are over 1000 times lower than the
approximate 0.001% parasitemia detectable by light microscopy [76,77].
Diagnosis of canine babesiosis should be as speciﬁc as possible, since
virulence, prognosis, and response to treatment are variable.

Clinical pathology

The severity of anemia is highly variable with babesial infections.

Researchers found that 50% of dogs infected with B canis rossi were severely
anemic (packed cell volume [PCV] less than 10) at presentation; 32% had
moderate anemia, and 18% were nonanemic or polycythemic [13]. Disease
in the severely anemic group was attributed primarily to immune-mediated
hemolytic anemia, and approximately 90% of severely anemic dogs were
Coombs positive [13].

Regeneration is usually proportional to the degree of anemia. Autoag-

glutination can be quite common and may occur in over 20% of dogs [49].
The leukocyte count is also extremely variable with babesiosis and can range
from leukopenic to leukamoid [66]. Leukocytosis usually is associated with
regenerative immune-mediated anemias.

Thrombocytopenia also is encountered frequently with babesial infec-

tions. The mechanism for the decrease in platelets is understood poorly. The
MPV generally is increased, suggesting that there is normal bone marrow
response to the use, sequestration, or destruction of platelets [6]. Local and
systemic DIC, immune-mediated destruction, and sequestration of platelets
in the spleen are possible mechanisms. Babesia infection triggers the host

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coagulation system; ﬁbrinogen degradation products (FDP’s) are typically
low, and a consumptive coagulopathy occurs at areas of local parasitism
[78].

Biochemical evidence of hypoxic hepatic damage occurred in severely

anemic B canis rossi infected dogs. Hepatic changes also were found in dogs
infected with the Californian small piroplasm, but changes were rare with
T annae

[36]. Electrolytes tended to be normal, and leukocyte counts

frequently were elevated in dogs with severe anemia [13,36]. Acute renal
failure is reported to occur in less than 3% of B canis infections. Clin-
icopathologic data from small Babesia infections in Spain indicated that
approximately 10% of parasitemic dogs had biochemical evidence of renal
failure [36].

Therapy and immunity

Species of Babesia vary in their susceptibility to babesiacidal treatments.

In general, small Babesia are considered more resistant to treatment. The
development of a chronic carrier state is suspected for small Babesia, while
most large Babesia infections are considered cleared with treatment. Some
drug resistance also is seen with pathogenic South African strains. Several
drugs have been used to treat babesiosis including: diminazene aceturate
(Berenil), imidocarb dipropionate (Forray [65]; Imizol), trypan blue, phe-
namidine isethionate (Oxopirvedine) [68], and quinuronium sulfate [70].
Diminazene is not available in the United States, but it is probably the most
commonly used drug worldwide. Diminazene is a diamidine derivative
closely related to phenamidine isethionate and pentamidine isethionate(Lo-
midine) [23]. The drug is thought to interfere with aerobic glycolysis
and inhibit DNA synthesis in the parasite [79]. A survey of practitioners
in South Africa [68] indicated that drug resistance and relapses were
suspected with diminazene treatment. Diminazene also may cause pain
at the injection site, vomiting, neurologic signs, hypotension and para-
sympathomimetic eﬀects [63]. Diminazene toxicity can cause depression,
stupor, vocalization, ataxia, increased extensor tone, nystagmus, seizures,
and possible death [63]. One dose of diminazene (3.5 mg/kg intramuscularly
[IM]) is recommended. B gibsoni does not respond as well as B canis to
treatment with diminazene.

Phenamidine isethionate is available in many countries for the treatment

of canine Babesia. Pentamidine isethionate, a closely related drug, is ap-
proved in the United States as an orphan drug for the treatment of
Pneumocystis

pneumonia in people. The drug has shown eﬃcacy against B

canis

and B gibsoni [80] but does not totally eliminate small babesial

infections [38].

Imidocarb, also a diamidine, is the only drug approved for the

treatment of Babesia in the United States. Imidocarb has direct eﬀects on
the nucleus and cytoplasm of the parasite. For treatment of B canis a single

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dose of 6 mg/kg IM or subcutaneously may be eﬀective for treating acute
disease and eliminating the carrier state. B gibsoni infections typically
are treated with a second injection repeated in 2 weeks. Possible adverse ef-
fects include pain at the injection site and parasympathomimetic signs.
Salivation, depression, lacrimation, muscle tremor, restlessness, tachycar-
dia, dyspnea and diarrhea may occur within 10 minutes of receiving the
injection [63]. Shivering, periorbital edema, and fever have been reported
10 to 12 hours after injection [63]. Premedication with atropine often is
given to reduce these adverse eﬀects. Imidocarb toxicity can cause severe
hepatic and renal damage, tachycardia, arrhythmias, lacrimation, diarrhea,
and tremors [63].

Trypan blue is used frequently to treat dogs with severe shock caused

by B canis rossi infection [68]. It may be able to block the entrance of the
parasite into the RBC. Infections usually are not eliminated with trypan
blue treatment, but parasitemia and clinical signs are reduced. Parasitemia
often returns within 2 weeks. Adverse eﬀects are minimal, and trypan blue
has a much broader therapeutic index than other babesiacidal drugs. Trypan
blue can be given as a 1% solution at a dose of 10 mg/kg intravenously (IV).
Tissue irritation and bluish discoloration of the mucosa and urine may
occur.

Ten of 15 dogs in Greece with B canis were treated with a 0.5% solution

of quinuronium sulfate (two injections given subcutaneously 48 hours apart
at a dose of 0.05 mL/kg) Clinical signs resolved within 48 hours of the ﬁrst
treatment [70]. The PCV may drop initially with treatment. This drop in
hematocrit also is seen with treatment for malaria and may be related to
parasite death or a lag phase before RBC destruction stops. More persistent
declines in PCV after babesiacidal treatment may be caused by the devel-
opment of immune-mediated destruction. Short-term therapy with pre-
dnisone may be indicated in some cases.

Chemotherapeutics are reportedly unable to completely eliminate B

gibsoni

infection. Theileria infections in cattle often are treated with

hydroxynaphthoquinone derivatives like atovaquone (Mepron). Based on
phylogenetic relationships, this also may be useful in Californian B gibsoni,
human B microti infections, and other small Babesia species. Pravaquone
has been suggested as a treatment for B gibsoni [15]. In hamster models,
atovaquone used alone caused drug resistance but was eﬀective when
combined with azithromycin. Azithromycin/quinine, azithromycin/atova-
quone and azithromycin/clindamycin/doxycycline were eﬀective in human B
microti

infections that failed to respond to standard clindamycin/quinine

therapy [81,82]. Preliminary studies using atovaquone and azithromycin in
canine B gibsoni infections appear promising, and several treated dogs be-
came PCR negative approximately 2 months after treatment [83].

Iron chelators are of potential beneﬁt with cerebral babesiosis. Therapy

with desferroxamine (DFO), an iron chelator, has proven beneﬁcial in
children with cerebral malaria [84]. Parasite clearance and recovery from

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coma were faster in pediatric patients treated with an infusion of DFO than
controls. Beneﬁts are attributed to radical scavenging properties, possible
enhancement of the TH1 immune response, and to reduced iron availability
for parasites [84]. Oral iron chelators eventually may play a role in therapy
of drug-resistant or relapsing strains of malaria. These agents have not been
examined in dogs.

Other less frequently used drugs include amicarbalide, euﬂavine, and

chloroquine. These drugs are less eﬀective and have more severe adverse
eﬀects [63].

Immunity

Young dogs are more susceptible to babesiosis and frequently have more

severe infections. [70]. At OVAH, 77% of dogs aﬀected with B canis were
less than 3 years old [85]. This is diﬀerent from the situation with bovine
babesiosis, where calves are more resistant than older animals [21]. A report
of 70 naturally occurring cases of babesiosis in Nigeria found that greater
than 70% were less than 1 year old [86]. Acute and hyperacute forms were
seen in dogs as young as 4 weeks old [86]. In suckling pups, B canis vogeli is
capable of causing severe disease that is manifested by anemia, marked
thrombocytopenia, jaundice, renal failure, and death [10]. Identical strains
subinoculated into splenectomized and nonsplenectomized adult dogs did
not cause obvious disease. Breed predispositions are not deﬁned strictly, but
anecdotal information revealed that in South Africa, rottweilers, German
shepherds, pit bulls, and border collies were over-represented [68]. Pit bulls
also were over-represented among dogs that died of nonanemic B canis rossi
infection [13]. The apparent susceptibility of pit bulls to large and small
babesiosis is not understood. Genetic factors, environmental conditions,
and vertical transmission likely are involved.

The development of immunity to babesial infection is characterized

poorly and often questioned. Veterinarians in South Africa report that dogs
treated with sterilizing drugs often relapse in the same season. Imidocarb
and diminazene often are thought to clear B canis infection totally and may
prevent the dog from developing protective immunity. Premunition is
thought to be important in South Africa, where babesiosis is extremely
virulent and widespread. Experimental studies have demonstrated that
heterologous strains did not provide cross-protection. Antibody levels are
thought to decline between 3 and 5 months after infection. In France, a ﬁrst-
generation vaccine has been available since 1984 [87]. The vaccine (Pirodog)
is made from B canis cell culture supernatants with a saponin adjuvant.
Eﬃcacy is around 89% for B c canis. The vaccine is not cross-protective for
the South African or other strains [87]. Vaccination reduces parasitemia but
does not eliminate infection. The lack of cross-protection is probably
partially responsible for vaccine failures in the ﬁeld.

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Human babesiosis

The ﬁrst case of human babesiosis was described in 1968. Since then,

hundreds of cases have been reported. Cases primarily occur along the East
Coast and in the Great Lakes area. Virtually all of the zoonotic cases in the
United States are caused by the rodent parasite B microti. In Europe, the
bovine pathogen B divergens is responsible for zoonotic infections primarily
in asplenic individuals with exposure to cattle [81]. Mortality rates for B
divergens

infections were approximately 38%, whereas mortality rates were

only 5% for B microti infections in the United States [81]. B microti infections
are typically treated with a combination of clindamycin and quinine [38]. In
severe cases, erythrocyte exchange transfusion may be life-saving [81]. B
divergens

infections are more serious and are treated with IV clindamycin

and oral quinine. In vitro studies showed eﬃcacy with imidocarb and
a combination of cotrimoxazole and pentamidine. Imidocarb is not approved
for use in people but has been used successfully in two Irish patients [81].

A new species of small Babesia has been recognized on the West Coast

[14]. In 1991, the pathogen, WA1, was isolated from a 41-year-old im-
munocompetent man in Washington with acute malarial-like symptoms.
WA1 caused more serious and prolonged illness than previously reported
human cases of B microti. Neighbors of the WA1 patient also showed
serologic evidence of exposure. Since this discovery, seroreactivity has been
found in many northern Californians. A similar situation exists with B
microti

, which often is accompanied by up to 17 silent infections for every

positive case found [88]. Each year, more cases of WA1 are being diagnosed
in Washington [14]. Two new strains, CA1 and MO1, also have been
reported in primarily in asplenic individuals in California [14] and Missouri
[89], respectively. Phylogenetic analysis revealed that the Californian small
Babesia

is the species most closely related to WA1.

Prevention

The best method of prevention in endemic areas is aggressive control of

the tick vector. An eﬀective topical acaricide combined with a ﬂea/tick collar
is usually very eﬃcacious in preventing tick exposure. Owners should inspect
their dogs daily for ticks. Prompt removal of ticks within 24 hours should
prevent disease transmission, because the tick must be attached for 2 to 3
days to transmit the organism [40]. In kennels where puppies are being lost
to disease, aggressive tick control measures should be instituted, including
spraying the environment and treating animals. Because babesiosis can
be transmitted vertically from dam to oﬀspring, serologic testing should be
performed to remove infected dogs from the breeding pool.

To avoid transmission through blood contamination, poor kennel

practices such as sharing needles for vaccination or reusing surgical
instruments for tail docking and ear cropping should be avoided. Dogs

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with positive Babesia titers should never be used as blood donors. More
stringent regulations concerning serologic testing, quarantine, and treatment
of dogs entering the United States from endemic areas may prevent
continued spread of the disease into this country. Dog ﬁghting should be
avoided, as it is also a potential method for spreading disease.

Treatment of asymptomatic dogs with subclinical infection is controver-

sial, and it is not known how eﬀective treatment is in eliminating the carrier
state. Treatment with imidocarb followed by a negative PCR test in
2 months may help eliminate the reservoir of disease, however. Development
of new vaccines, such as the French vaccination for B canis, is needed,
because there is no cross-protection for the diﬀerent Babesia organisms. As
evidence mounts concerning the spread of this emerging tickborne disease to
new areas, continued research is needed to determine better methods of
prevention and treatment.

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American canine hepatozoonosis

Nancy A. Vincent-Johnson, DVM, MS

United States Army Veterinary Corps, 10015 Theote Road,

Building 610, Fort Belvoir, VA 22060-5400, USA

The ﬁrst cases of canine hepatozoonosis in the United States were

recognized in 1978 near the Gulf Coast region of Texas [1]. Since that time,
the disease has been diagnosed in dogs throughout much of the southeastern
United States. Although originally believed to be caused by Hepatozoon
canis

, the course of the disease was much more severe than canine hepato-

zoonosis seen elsewhere in the world. Not only did the clinical signs and
laboratory abnormalities diﬀer but so did pathological characteristics, tissue
tropism, parasite morphology, and tick vectors. In 1997, the organism was
described as a separate species, Hepatozoon americanum [2]. Genetic and
antigenic comparative studies further substantiate that it is related to but
a distinct species from H canis [3–5].

Hepatozoon americanum

and H canis are apicomplexan protozoa from

the family Haemogregarina. H canis was reported ﬁrst in dogs in India in
1905 [6] and has since been identiﬁed in Africa [7,8], Southeast Asia [9–11],
the Middle East [12,13], and Southern Europe [14–17]. H canis appears to be
adapted to dogs, causing subclinical disease or only mild clinical signs in
most cases. Immunosuppression caused by concurrent disease or other fac-
tors appears to play an important role in manifestation of clinical signs. This
is in contrast to H americanum, where immunosuppression or concurrent
disease is not necessary to induce the more severe clinical signs typically
seen. H americanum appears to be poorly adapted to dogs and is likely
a natural parasite of another species, with the dog becoming an accidental
host after ingestion of infected ticks. The vector for H americanum is the
Gulf Coast tick, Amblyomma maculatum [18], while the vector for H canis is
the Brown Dog tick, Rhipicephalus sanguineus [19]. Both species of Hepato-
zoon

are transmitted trans-stadially in their tick vectors. A comparison of

the major diﬀerences between H americanum and H canis is shown in
Table 1.

Vet Clin Small Anim

33 (2003) 905–920

E-mail address:

nvincent@LN.amedd.army.mil

0195-5616/03/$ - see front matter

doi:10.1016/S0195-5616(03)00028-7

Table

Com

parison

patozoon

ame

ricanum

and

Hepatoz

oon

can

Hepatoz

oon

america

num

Hepatoz

oon

can

Tick

vect

Ambly

omm

maculatum

(Gu

oast

tick)

Rhipicephalus

sanguine

(bro

dog

tick)

Prima

clinical

sign

Feve

pain,

lam

eness,

mucop

urulent

ocula

disc

harge;

may

and

wane

Freque

ntly

asym

ptoma

tic;

can

cause

lethargy

fever,

weight

loss

Com

mon

labora

tory

abnorm

alities

Extr

eme

leukocy

tosis

(20,00

0–200,00

leu

kocytes/

anem

ia,

ele

vated

alka

line

osphat

ase,

low

gluco

Anemia;

extre

leukocy

tosis

rare

may

see

dogs

ith

very

high

parasitemia

Con

current

infe

ctio

immunosu

ppres

sion

Occasio

nal

Very

mmon

Geog

raphic

distribu

tion

United

State

Cent

ral

and

South

Americ

Africa,

Middle

East

Asia,

South

ern

Euro

pe,

South

Americ

Tiss

stage

Muscle

lesio

nsisting

‘‘oni

on-skin’’

cysts

eronts,

pyogra

nulom

as,

myositis

muscle

lesions

Meron

exh

ibit

blast

ophore

formatio

‘‘Wheel-spoke

d’’

ronts

fou

primarily

spleen

bone

mar

row,

lymph

nodes

Rad

iograp

hic

lesions

Periost

eal

pro

liferation

None

(exc

ept

case

repor

ted

Japan

)

Sever

ity

dise

ase

Sever

guarded

progn

osis

Subclin

ical

severe,

usually

mild;

good

progn

osis

Freq

uency

gam

onts

ripheral

blood

Rare;

parasitemia

usually

0.1%

Common

par

asitemia

1–10

(usually

1%)

Gamo

charac

teristics

Size:

8.8

3.9

ultrast

ructure:

tail-like

appe

ndage

;

lacks

ﬁn

ﬁ

ril-like

ructure

see

canis

Size:

11.0

4.3

ultra

struct

ure:

ﬁne

ﬁbril-l

ike

struct

ure

surr

oundin

parasit

ophoro

vac

uole

Antib

odies

america

num

IFA

show

good

correlation

ith

muscle

bio

psy

Low

cro

ss-reactivity

canis

IFA

can

IFA

shows

high

equenc

antibo

dies

gener

dog

populat

ion

(Israe

Treatm

ent

Trime

thoprim

/sulfadiaz

ine,

pyr

imethamine,

clin

damycin

ecoquin

ate

Imidocar

dipro

prionat

xycycline

Abbre

viatio

TFA,

imm

unoﬂu

orescen

antib

ody.

906

N.A. Vincent-Johnson / Vet Clin Small Anim 33 (2003) 905–920

Geographical distribution and epidemiology of Hepatozoon americanum

Canine hepatozoonosis caused by H americanum is an emerging disease

that has spread north and east from the Gulf Coast of Texas where it was
detected originally. It subsequently has been reported in dogs from
Louisiana, Alabama, Georgia, Oklahoma, Florida, and Tennessee [20–24].
Researchers at Oklahoma State University performed a retrospective study
of muscle from dogs necropsied there from 1989 to 1995 and conﬁrmed that
H americanum

had not been overlooked in dogs before 1995 [25]. The geo-

graphical distribution of the parasite parallels that of the deﬁnitive host, the
Gulf Coast Tick, Amblyomma maculatum. Although the range of this tick
once was believed to be limited to the warm, humid areas near the Gulf
Coast, it appears that the range has expanded to as far north as southern
Kansas and Kentucky [26]. This tick also exists throughout Central America
and the northern part of South America. Eight cases of hepatozoonosis were
reported from Brazil, but it is unclear whether the dogs were infected with
H americanum

or H canis [27].

In addition to dogs, H americanum or a similar organism has been

diagnosed in coyotes, bobcats, and ocelots in the southern United States by
identiﬁcation of cysts and meronts in muscle or gamonts on blood smears
[28–31]. Most of these wild animals were in good physical condition at the
time of their capture. In contrast to adult animals, coyote puppies that were
infected experimentally did develop classic signs of canine hepatozoonosis,
including the typical bone lesions [32].

Life cycle and transmission

Hepatozoon americanum

requires two hosts to complete its life cycle. The

Gulf Coast tick, Amblyomma maculatum, is the deﬁnitive host in which
sexual reproduction occurs. Dogs, other canids, and possibly another spe-
cies of vertebrate serve as the intermediate host where asexual reproduction
takes place.

While feeding on an infected dog or other intermediate host, the nymphal

tick ingests white blood cells containing H americanum gamonts [33]. After
they are freed from the leukocytes, the gamonts form into male and female
gametes, and fertilization occurs. The resulting zygotes develop into oocysts
in the gut of the tick as the tick undergoes a molt. The newly emerged adult
tick contains several mature oocysts within its hemocoele (Fig. 1). Each
oocyst contains hundreds of sporocysts, each of which contains 10 to 26
sporozoites. A dog becomes infected when it ingests an adult tick containing
oocysts [34]. The sporozoites are released from the oocysts within the dog’s
gastrointestinal [GI] tract, pass through the gut wall, and are transported
through the circulation to striated muscle and other target tissues. The
parasite develops between myocytes within a protective host cell [35]. Layers
of mucopolysaccharides are laid down around the host cell, forming an

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N.A. Vincent-Johnson / Vet Clin Small Anim 33 (2003) 905–920

‘‘onion-skin’’ cyst (Fig. 2). The organism undergoes merogony within the
cyst and upon maturation releases merozoites from the ruptured cyst. This
elicits a severe inﬂammatory response, bringing neutrophils and monocytes
into the area to form a pyogranuloma where the meront once existed
(Fig. 3). Many of the inﬂammatory cells become infected with single zoites.
The pyogranuloma undergoes angiogenesis, thus forming a highly vascular
structure through which infected neutrophils and monocytes can return to
the circulation. In this manner, the intracellular parasites may travel to
other sites to continue the asexual cycle or they may become circulating
gamonts that complete the life cycle when ingested by ticks feeding on the
dog. In experimental infections, completion of the life cycle in the dog from

Fig. 1. Five H americanum oocysts, each containing hundreds of sporocysts. Freed sporocysts
also can be seen in this cytological preparation from the hemocoele of an infected Amblyomma
maculatum

tick. (Modiﬁed Wright-Giemsa stain. Original magniﬁcation

40.)

Fig. 2. A typical ‘‘onion-skin’’ cyst in a dog infected with H americanum. (Hematoxylin and
eosin stain. Original magniﬁcation

200.)

908

N.A. Vincent-Johnson / Vet Clin Small Anim 33 (2003) 905–920

ingestion of an infected tick to development of circulating gamonts occurs
in as few as 32 days [35]. In the tick, it takes 42 days for the organism to
develop into mature oocysts that are present in the hemocoele of the newly
molted adults [33].

Recent studies have shown that larval ticks experimentally exposed to

H americanum

through feeding on an infected dog also develop oocysts

and are infectious to dogs as nymphs and adults [36].

Transmission of H americanum occurs by way of ingestion of infected

Amblyomma maculatum

ticks. Unlike most tickborne diseases, Hepatozoon

organisms do not migrate to the salivary glands of the tick and cannot be
transmitted through a tick bite. Researchers at Oklahoma State University
have shown that coyotes also can be experimentally infected with
H americanum

and that they can transmit the organism to susceptible ticks

feeding on them [32]. Because the host range for larval and nymphal
Amblyomma maculatum

ticks is so diverse, there is a large potential variety

of vertebrate candidates that could act as intermediate hosts in maintaining
an endemic cycle for H americanum.

Vertical transmission from dam to puppies has been documented in

H canis

infection [37] and probably also occurs with H americanum. It also has

been hypothesized that dogs may become infected through carnivorism by
way of ingestion of tissue stages of the organism. Although this is known to
occur with other species of Hepatozoon [38], it has not been proven with
H americanum

Pathogenesis

Immunosuppression is not necessary for H americanum to produce disease

in dogs. In one retrospective study, concurrent illness was identiﬁed in only
half of the dogs with H americanum infection, and some households had more

Fig. 3. Pyogranuloma in skeletal muscle. Zoites can be seen displacing the nucleus in several
infected cells. (Hematoxylin and eosin stain. Original magniﬁcation

400.)

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N.A. Vincent-Johnson / Vet Clin Small Anim 33 (2003) 905–920

than one dog diagnosed with hepatozoonosis [22]. Age is not an important
factor either, as clinical hepatozoonosis occurs in dogs of various ages.

Experimentally, the earliest lesions are seen at 3 weeks following infection

when the organism can be observed within a host cell located between
myocytes [35]. As the zoite matures, the host cell produces layers of
mucopolysaccharides that surround the cell, likely protecting the zoite from
the dog’s immune system. Clinical signs are associated with the inﬂam-
matory response that occurs after the cyst ruptures and pyogranuloma
formation occurs. Cysts apparently can lie dormant and may be found in
dogs years after successful treatment. Activation of these cysts results in
continuation of the cycles of asexual reproduction, which account for the
waxing and waning nature of the disease and the occurrence of relapses
following treatment.

Clinical signs

Gait abnormalities, including lameness, weakness, stiﬀness, recumbency,

or diﬃculty rising are the most common reasons for presentation of dogs
infected with H americanum. On physical examination, these dogs may
exhibit pain, poor body condition, muscle atrophy, weakness, depression,
and mucopurulent ocular discharge (Table 2). Fevers of up to 106

F (41

are common, although body temperature may be normal and often
ﬂuctuates with the waxing and waning of clinical signs. Dogs typically lose
weight and exhibit muscle wasting, despite the fact that many of them

Table 2
Frequency of clinical signs in Hepatozoon americanum infection

Clinical sign

Number of dogs

Percent

Fever

19/22

Weight loss

18/22

Mucopurulent ocular discharge

17/22

low tear production

8/22

Muscle atrophy

14/22

Pain (all types)

14/22

joints

2/22

lumbar

4/22

cervical

5/22

generalized

3/22

Stiﬀness

12/22

Generalized weakness

9/22

Rear limb paresis and ataxia

5/22

Inability to rise

5/22

Anorexia

5/22

Data from

Macintire DK, Vincent-Johnson N, Dillon AR, Blagburn B, Lindsay D, Whitley

EM, Banﬁeld C. Hepatozoonosis in dogs: 22 cases (1989–1994). J Am Vet Med Assoc 1997;
210:916–22.

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N.A. Vincent-Johnson / Vet Clin Small Anim 33 (2003) 905–920

maintain a fairly normal appetite. Muscle atrophy may be very marked, and
chronic cachexia or emaciation may be evident (Fig. 4).

Clinical signs may mimic those of meningitis or discospondylitis. Dogs

frequently exhibit hyperesthesia or pain, which can manifest as cervical,
back, joint, or generalized pain. Guarding of the cervical region may result
in the so-called ‘‘Master’s Voice’’ stance. Pain results from inﬂammation
associated with the pyogranulomatous muscle lesions and also from the
periosteal reaction that causes bony proliferation.

An appearance of matted eyes is common because of the occurrence

of a mucopurulent ocular discharge that sometimes is associated with
decreased tear production (Fig. 5). The matted eyes tend to correlate with
the waxing and waning of clinical signs, and owners often report an ocular
discharge as the ﬁrst sign of a relapse.

Dogs may have a history of polyuria and polydipsia, especially if

secondary glomerulonephritis or amyloidosis is present. Transient, bloody

Fig. 4. Severe muscle atrophy and emaciation is evident in this Rottweiler with chronic
American hepatozoonosis.

911

N.A. Vincent-Johnson / Vet Clin Small Anim 33 (2003) 905–920

diarrhea is seen at times. Less common clinical signs include abnormal lung
sounds, cough, pale mucous membranes, and lymphadenopathy.

Laboratory abnormalities

The most consistent laboratory abnormality is an elevated white blood

cell count (Table 3). It commonly ranges from 20,000 to 200,000 cells/mm

and typically consists of a mature neutrophilia, although a left shift may be
present. Neutrophils may appear hypersegmented on microscopic exami-
nation. In two diﬀerent retrospective studies, the mean initial leukocyte
count in infected dogs was 76,807 and 85,700 cells/mm

, respectively [20,22].

A mild-to-moderate normochromic, normocytic, nonregenerative anemia is
also a frequent ﬁnding. The platelet count is typically normal to elevated.
Thrombocytosis, with counts of 422,000 to 916,000 platelets/mm

, has been

reported in a number of dogs in one study [22]. In those cases that have
a decreased platelet count, there is often concurrent disease with ehrlichiosis,
babesiosis, Rocky Mountain spotted fever, or other tickborne disease.

Fig. 5. A mucopurulent ocular discharge is very common in dogs infected with H americanum.
Note the severe muscle atrophy of the temporalis muscles in this dog.

912

N.A. Vincent-Johnson / Vet Clin Small Anim 33 (2003) 905–920

Serum chemistry abnormalities are also common. A mild elevation in

alkaline phosphatase is usually present. The blood glucose level is decreased
in a large number of cases and may occasionally be as low as 5 mg/dL (0.28
mmol/L). This is an in vitro artifact caused by increased metabolism of
glucose by the high number of leukocytes rather than a true hypoglycemia.
When blood is drawn into a sodium ﬂuoride tube to stop glucose metab-
olism, the glucose level is usually normal. Albumin frequently is decreased
and may be caused by decreased protein intake, chronic inﬂammation, or
renal loss. Unless severe renal disease is also present, the blood urea nitrogen
(BUN) often is decreased. The combination of low glucose, albumin, and
BUN, along with an elevation of alkaline phosphatase, gives the impression
of liver failure. Fasting and postprandial bile acids in infected dogs are
usually normal or only slightly elevated, however. Despite the muscle
inﬂammation, creatine phosphokinase (CPK) is almost always normal.
Hyperglobulinemia is uncommon.

Infected dogs may develop glomerulonephritis or amyloidosis. Urinalysis

in these cases will show proteinuria and an elevation in the urine
protein:creatinine ratio.

Radiographic ﬁndings

Dogs infected with H americanum frequently exhibit periosteal pro-

liferation that results in new bone formation along the periosteum,
particularly of the long bones. Radiographically, the lesions range from

Table 3
Frequency of laboratory and radiographic ﬁndings in Hepatozoon americanum infection

Finding

Number of dogs

Percent

Extreme leukocytosis

22/22

100

mature neutrophilia

15/22

mild left shift

7/22

Elevated alkaline phosphatase

22/22

100

Low glucose

20/22

Low albumin

19/22

Periosteal proliferation on x-ray

18/21

Anemia (nonregenerative)

14/22

Low calcium

14/22

Normal platelet count

11/22

Thrombocytosis

9/22

Elevated phosphorous

7/22

Low blood urea nitrogen

7/22

Hyperglobulinemia

4/22

Elevated calcium

2/22

Elevated creatine phosphokinase

1/22

Data from

Macintire DK, Vincent-Johnson N, Dillon AR, Blagburn B, Lindsay D, Whitley

EM, Banﬁeld C. Hepatozoonosis in dogs: 22 cases (1989–1994). J Am Vet Med Assoc 1997;
210:916–22.

913

N.A. Vincent-Johnson / Vet Clin Small Anim 33 (2003) 905–920

subtle bone irregularity to a smooth laminar thickening (Fig. 6). The
pathogenesis of the new bone formation is unknown. Morphologically, the
development of the bone lesions is very similar to that of hypertrophic
osteopathy [39].

Diagnosis

Gamonts of H americanum are observed infrequently on microscopic

examination of blood smears from infected dogs [40]. When present, they
typically are found in less than 0.1% of the neutrophils or monocytes.
The gamonts appear within the cytoplasm of leukocytes stained with
Romanowsky-type stains as a light blue to clear oblong capsule measur-
ing approximately 8.8

3.9 microns with a faintly staining nucleus (Fig. 7).

Special staining techniques [41] have been advocated for making the infected
cells stand out, but diagnosis by examination of blood smears is still
unreliable because of the extremely small numbers of organisms normally

Fig. 6. Proliferative bone lesions range from subtle bone irregularity to a smooth laminar
thickening as seen in this radiograph.

914

N.A. Vincent-Johnson / Vet Clin Small Anim 33 (2003) 905–920

present. Gamonts may exit the cells rapidly after blood is drawn, leaving
behind an empty capsule that is diﬃcult to identify. Examination of buﬀy
coat smears increases the chance of seeing gamont-infected cells.

A more reliable method of diagnosis is by muscle biopsy. With the dog

under general anesthesia, two or three small pieces of muscle (approximately
2 cm

2 cm) are taken from the biceps femoris or semitendinosus muscle,

although epaxial or other muscles also may be sampled [24]. Cysts, meronts,
and pyogranulomas with zoite-containing neutrophils and monocytes are
diagnostic of H americanum infection. Myositis is a less speciﬁc but common
ﬁnding. False negatives occur when a biopsied muscle piece does not contain
any lesions. Taking several pieces of muscle during biopsy increases the
probability of obtaining a diagnosis in an infected dog.

A relatively new method of diagnosis is by serology. An ELISA blood

test that detects antibody to H americanum by using antigen obtained from
sporozoites [42] has been developed by researchers at Oklahoma State
University. The test appears to be highly sensitive (93%) and speciﬁc (96%)
when compared with muscle biopsy.

Bone marrow aspirates typically show an increased myeloid:erythroid

ratio indicative of granulocytic hyperplasia, but they normally fail to dem-
onstrate any H americanum organisms. Likewise, lymph node aspirates usu-
ally demonstrate lymphoid hyperplasia but not organisms.

Pathological ﬁndings

Gross necropsy ﬁndings of dogs chronically infected with H americanum

usually include muscle atrophy and cachexia. Bone surfaces may be
roughened and thickened. Pyogranulomas can appear as 1 to 2 mm
diameter white or tan foci scattered throughout muscle and other tissue [40].

Fig. 7. A gamont of H americanum in a blood smear from an infected dog. Note the
hypersegmentation of the adjacent neutrophils. (Modiﬁed Wright-Giemsa stain. Original
magniﬁcation

1000.)

915

N.A. Vincent-Johnson / Vet Clin Small Anim 33 (2003) 905–920

Histologic exam reveals various stages of the organism in skeletal and

cardiac muscle. The most common stage is the large ‘‘onion-skin’’ cyst: a
round-to-oval structure usually containing a single round basophilic nucleus
surrounded by concentric layers of light blue staining laminar membranes,
averaging almost 200 lm in diameter [2]. Less commonly, a developing meront
is found inside of the cyst. Pyogranulomas are comprised of large accumu-
lations of macrophages and neutrophils, many of which contain round intra-
cellular parasites. Myositis with muscle atrophy, necrosis, and inﬁltration of
inﬂammatory cells between muscle ﬁbers is a common ﬁnding. In addition to
skeletal and cardiac muscle, the ‘‘onion-skin’’ cysts or pyogranulomas have
been found in intestinal smooth muscle, adipose tissue, pancreas, spleen,
lymph node, liver, skin, kidneys, and lungs.

Other histologic ﬁndings include: vascular changes, consisting of ﬁbri-

noid degeneration of vessel walls, mineralization and proliferation of vas-
cular intima, and pyogranulomatous vasculitis; and lung inﬁltrates with
diﬀuse interstitial thickening of alveolar septa. Renal lesions are common
and include focal pyogranulomatous inﬂammation with mild glomerulone-
phritis, lymphoplasmacytic interstitial nephritis, and mesangioproliferative
glomerulonephritis. Amyloid deposits have been found in the spleen, lymph
nodes, small intestine, liver, and kidneys of dogs with chronic H americanum
infection. Schizonts with the characteristic wheel-spoke pattern typical of
H canis

have not been observed [40].

Treatment

Treating dogs with American canine hepatozoonosis has been frustrating

in the past because of frequent relapses that often result in worsening
episodes of disease. Evaluation of treatment also has been diﬃcult because
of the waxing and waning course of the disease. Various antiprotozoal drugs
have been tried, but no drug has been found to eliminate all tissue stages of
the organism. Remission of clinical signs usually can be obtained through
the following combination therapy, which will be referred to as TCP:
trimethoprim-sulfadiazine (Tribrissen, Di-Trim) at 15 mg/kg by mouth
every 12 hours; clindamycin (Antirobe) at 10 mg/kg by mouth every 8 hours;
and pyrimethamine (Daraprim) at 0.25 mg/kg by mouth every 24 hours [22].
This treatment should be continued for 14 days. Dogs generally respond
with resolution of fever and pain within 48 hours of initiating therapy.
Remission is often short-lived as dogs frequently relapse within 2–6 months.
Although response to treatment is generally good with subsequent relapses,
the relapses may recur more frequently over time. With each relapse, the
chances of the dog developing complications of glomerulonephropathy,
amyloidosis, vasculitis, and cachexia increase. A relatively new treatment
to assist in prevention of clinical relapses is decoquinate (Deccox), an
anticoccidial drug available for large animals [43]. This drug does not
appear to be useful in resolving active clinical disease, but it is markedly

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N.A. Vincent-Johnson / Vet Clin Small Anim 33 (2003) 905–920

useful in preventing relapses if given daily after clinical signs have resolved.
Dogs are started on decoquinate at 10 to 20 mg/kg by mouth every 12 hours.
The drug is given continuously for 2 years. If one or more dosages of
decoquinate are missed, relapse frequently follows approximately 1 week
later. When a relapse occurs, the dog should be given another course of the
TCP combination therapy. Decoquinate is available in the United States as
a livestock feed additive at a concentration of 27.2 g of decoquinate per
pound of premix. The powder can be mixed into moist dog food at a rate of
0.5 to 1.0 teaspoons per 10 kg body weight and fed twice a day.

Symptomatic treatment consisting of nonsteroidal anti-inﬂammatory

drug (NSAID) administration is used to relieve fever and pain associated
with the inﬂammation, particularly during the ﬁrst couple of days of therapy
before the antiprotozoal drug takes eﬀect.

Prognosis

In the past, prognosis for dogs with H americanum infection was guarded

to poor. Many dogs died or were euthanized because of the severe clinical
signs of disease during the initial episode or relapses. Many dogs succumbed
to sequelae of the disease, particularly glomerulonephritis or amyloidosis.

With the advent of the TCP combination therapy and daily decoquinate

therapy, the prognosis for dogs with H americanum infection has improved.
Relapses are not as frequent or severe; glomerulonephritis or amyloidosis is
less common; and survival rates have markedly increased. In a comparison
of treatment protocols, the 2-year survival rate for dogs receiving only
toltrazuril (Baycox) or the TCP combination therapy was 12.5% [43]. When
the combination therapy was followed by long-term daily decoquinate
therapy, the 2-year survival rate increased to greater than 84%. The median
survival time for those that received the toltrazuril or TCP combination
therapy alone was approximately 12 months.

Prevention

Prevention of American canine hepatozoonosis is through tick control

using an eﬀective acaricide and examining dogs frequently to remove ticks,
particularly after hunting or roaming outdoors. Although transmission by
ingestion of infected tissue has not been proven to occur with H americanum,
it is plausible, because it happens with other species of Hepatozoon. There-
fore, dogs should not be allowed to eat raw meat or organs from wildlife in
endemic areas.

Summary

Hepatozoon americanum

infection is an emerging tickborne disease in the

southern United States. This organism causes a very diﬀerent and much

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N.A. Vincent-Johnson / Vet Clin Small Anim 33 (2003) 905–920

more severe disease than does Hepatozoon canis, the etiologic agent of
canine hepatozoonosis in the rest of the world. H americanum is transmitted
through ingestion of the deﬁnitive host, Amblyomma maculatum (the Gulf
Coast tick). Clinical signs of American canine hepatozoonosis tend to wax
and wane over time and may include lameness, weakness, pain, muscle
atrophy, fever, and mucopurulent ocular discharge. Radiographs typically
reveal periosteal proliferation of various bones. Extreme leukocytosis is
the most common laboratory ﬁnding, along with a mild elevation of serum
alkaline phosphatase. Diagnosis is made by visualization of gamont-con-
taining neutrophils or monocytes on examination of blood smears; obser-
vation of typical cysts, meronts or pyogranulomas on muscle biopsy; or
detection of serum antibodies against H americanum sporozoites. Common
complications of chronic infection include glomerulopathies, amyloidosis,
and vasculitis. Although the prognosis for this disease in the past was
guarded to poor, recent advances in treatment have increased the long-term
survival rate of infected dogs.

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Canine visceral leishmaniasis and its

emergence in the United States

Alexa C. Rosypal, BS

, Anne M. Zajac, DVM, PhD

David S. Lindsay, PhD

Department of Biomedical Sciences and Pathobiology, Virginia–Maryland Regional College

of Veterinary Medicine, 1410 Prices Fork Road, Blacksburg, VA 24061

Department of Biomedical Sciences and Pathobiology, Virginia–Maryland Regional College

of Veterinary Medicine, Duck Pond Drive, Blacksburg, VA 24061

Department of Biomedical Sciences and Pathobiology, Virginia-Maryland Regional College

of Veterinary Medicine, 1410 Prices Fork Road, Blacksburg, VA 24061

Leishmaniasis is an important insect vectored disease that accounts for

approximately 57,000 human deaths worldwide annually [1]. Leishmaniasis
is a zoonosis. Dogs are important reservoirs of human visceral leishmaniasis
(HVL) [2], and dog ownership is considered a risk factor for HVL in most
endemic areas [3]. Leishmania species can cause visceral, cutaneous, or
mucocutaneous disease in people and animals. Members of the Leishmania
donovani

complex cause human and canine visceral leishmaniasis (CVL) in

parts of Europe, the Middle East, Africa, Asia, China and the Americas [4].
The member of the L donovani complex identiﬁed in dogs in the United
States [5] and most often found in the Mediterranean basin, China, and the
Middle East is L infantum [6]. A similar or identical parasite, L chagasi, is
recognized as a cause of HVL and CVL in Latin America [4].

Until early 2000, most public health oﬃcials and veterinarians thought

CVL was an unimportant disease in the United States. Most reported cases
were in dogs that had originated or traveled from areas where leishmaniasis
is endemic [7]. The recognition that L infantum was actually endemic in
foxhounds in the United States in 1999 [5] changed this thinking, and now
there is concern that the disease may make its way into the human
population [8].

Vet Clin Small Anim

33 (2003) 921–937

The authors wish to recognize the ﬁnancial support of grant MAF DO1CA-16 from the

Morris Animal Foundation. Alexa C. Rosypal is a Morris Animal Foundation Fellow.

* Corresponding author.
E-mail address:

lindsayd@vt.edu (D.S. Lindsay).

0195-5616/03/$ - see front matter

doi:10.1016/S0195-5616(03)00030-5

Leishmania life cycle

Leishmania

species are ﬂagellated protozoan parasites in the Phylum Sar-

comastigophora

that are transmitted by the bite of infected female sand ﬂies.

Sand ﬂy vector biology

Sand ﬂies are small bloodsucking Dipteran ﬂies in the family Psychodidae,

subfamily Phlebotominae. Six genera of sand ﬂies are recognized, but only
two genera are of medical importance: Phlebotomus in the Old World and
Lutzomyia

in the New World [9]. They are found mainly in the tropics,

subtropics, and temperate regions and are distributed between 50

N and

S in the northern and southern hemispheres (Tables 1 and 2). Sand ﬂies

live in a variety of habitats, including desert, rainforest, and intra-
domiciliary. In addition to leishmaniasis, sand ﬂies also vector the etiologic
agent of bartonellosis and several arboviruses.

Phlebotomine

sand ﬂies are small, delicate ﬂies with long legs and

mouthparts. They are covered with hair and hold their wings above their
bodies in a characteristic ‘‘V’’ shape. Sand ﬂies are poor ﬂiers and exhibit
a short hopping ﬂight when approaching a host. Because of their poor ﬂight
ability, it is assumed that sand ﬂies generally stay close to their breeding
sites, although they can travel longer distances if carried by the wind [10].

Sand ﬂies are crepuscular or nocturnal in activity, but they will bite

during the daylight if disturbed. Resting places are cool, humid micro-
environments and include tree holes, caves, houses, and animal burrows.
Males and females feed on a sugar source, but only females blood feed
and use nutrients in the blood for egg development. Leishmania-infected
sand ﬂies are unable to take a single full blood meal and tend to probe
their host before feeding, which increases the possibility of parasite trans-
mission. Probing may be induced by damage to the cardiac valve by way
of the activity of chitinolytic enzymes produced by promastigotes, which
subsequently obstructs engorgement [11]. Infected sand ﬂies tend to probe
and feed more often to complete engorgement. While blood feeding,
sand ﬂies salivate into the bite wound of the vertebrate host. Sand ﬂy
saliva contains vasodilatory, anti-inﬂammatory, and immunomodulatory
compounds that enhance Leishmania infection in the mammalian host.

Sand ﬂies undergo complete metamorphosis, and their lifecycle consists

of egg, four larval instars, pupa, and adult. Most species are terrestrial for
their entire lives. The life cycle of sand ﬂies is variable and depends on the
species, food availability, and environmental factors. Depending on the
Phlebotomine

species, females mate before, during, or after blood feeding.

Eggs are laid after blood feeding, and the time from taking a blood meal to
oviposition depends on the species and ambient temperature. Because of
their small size and diverse ecological habitats, immature sand ﬂies have
proved diﬃcult to ﬁnd. As a result, most of what is known about the sand ﬂy

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A.C. Rosypal et al / Vet Clin Small Anim 33 (2003) 921–937

Table 2
Phlebotomine sand ﬂies (genus: Lutzomyia) in the United States

Species

Distribution

Vector
competency

Species vectored
(Leishmania species)

L cruciata

Florida

Suspected

L mexicana

L diabolica

Texas

Proven

L mexicana

L xerophila

Southern California

Unknown

L anquilonia

Colorado, Washington

Unknown

L anthophora

Texas

Proven

L mexicana

L shannoni

Southeastern United States

Proven

L infantum

L tanyopsis

Arizona

Unknown

L texana

Texas

Unknown

L apache

Arizona, Texas

Unknown

L oppidana

Texas, Colorado, Washington, Montana

Unknown

L stewarti

California

Unknown

L vexator

Western, southeastern United States

Unknown

L californica

California, Washington

Unknown

L cubensis

Florida Keys

Unknown

Data from

Young DG, Perkins PV. Phlebotomine sand ﬂies of North America

(Diptera:Psychodidae). Mosq News 1984;44(2):264–304.

Table 1
Proven or suspected vectors of canine leishmaniasis

Vector species

Causative organism
(Leishmania species)

Distribution of sand ﬂy

Old world vectors

Phlebotomus

P chinensis

L infantum

North and central China

P longiductus

L infantum

North Africa and central Asia

P perniciosus

L infantum

Mediterranean basin

P ariasi

L infantum

Western Mediterranean

P perﬁliewi

L infantum

Mediterranean basin

P longicuspis

L infantum

North Africa, Spain

P neglectus

L infantum

Eastern Mediterranean

P tobbi

L infantum

Eastern Mediterranean

P kandelakii

L infantum

Lebanon, Turkey, Iran, Afghanistan

P syriacus

L infantum

Israel, Jordan, Syria

P langeroni

L infantum

North Africa, Spain

P smirnovi

L infantum

Central Asia

P transcaucasicus

L infantum

Azerbaijan

New world vectors

Lutzomyia

L longipalpis

L infantum

L. chagasi

)

Central and South America

L evansi

L infantum

Columbia, Costa Rica, Venezuela

L youngi

L infantum

Central and South America

L shannoni

L infantum

Southeastern United States, South America

Proven vector.

Data from

Killick-Kendrick R. The biology and control of Phlebotomine sand ﬂies. Clin

Dermatol 1999;17(3):279–89.

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A.C. Rosypal et al / Vet Clin Small Anim 33 (2003) 921–937

lifecycle comes from observations of common laboratory reared species such
as Phlebotomus papatasi [12].

Fourteen species of sand ﬂies (genus: Lutzomyia) have been recorded in

North America (Table 2) [13]. The most common species in the United
States are Lutzomyia shannoni, L cruciata, L anthophora, and L diabolica.
None of these has been shown to transmit Leishmania infection to dogs;
however, L shannoni has been infected experimentally by feeding on
symptomatic dogs [14]. L anthophora and L diabolica are suspected as
vectors of cutaneous leishmaniasis caused by L mexicana [15].

Developmental cycle in the host

The female sand ﬂies inject ﬂagellated stages (promastigotes) into the skin

of the host while feeding (Fig. 1). Promastigotes are elongate stages that have
a single nucleus, an anterior ﬂagellum, and a kinetoplast (Fig. 2). They divide
by longitudinal binary ﬁssion. The kinetoplast is an area of the parasite’s
mitochondrion that contains large amounts of mitochondrial DNA. The
kinetoplast stains dark like the nucleus with blood stains. The ﬂagellum
originates in the vicinity of the kinetoplast. Promastigotes are motile and pull
themselves along by their anterior ﬂagellum. The parasites are ingested by
macrophages, but they are not killed. Inside macrophages, the promastigotes
withdraw their external ﬂagellum and become amastigotes. Amastigotes are
round to slightly oval stages (Fig. 3). They have a single nucleus,
a kinetoplast, and a rudimentary ﬂagellum. The amastigotes divide by
binary ﬁssion, until they rupture the host cell. The released amastigotes are
ingested by other macrophages, and new cells are infected. Infection spreads
from the skin to internal organs by movement of infected macrophages or
amastigotes in the vascular system. Sand ﬂies become infected when they
ingest amastigotes while feeding on an infected host. Inside the gut of the ﬂy
the amastigotes become promastigotes. The promastigotes migrate to the
hypostome of the sand ﬂy and are inoculated when the ﬂy feeds. The
promastigotes are ingested by macrophages, and the new host is infected.

Evidence indicates that transmission Leishmania species also may occur

through exchange of blood or other bodily secretions [16]. Transmission by
blood transfusion [17] and by packed red blood cell (RBC) transfusion [18]
recently has been demonstrated in US dogs that have received blood from
donor foxhounds. Congenital transmission of visceral leishmaniasis has
been reported several times in people [19]. It is less clear if congenital
transmission occurs in dogs. A case of Leishmania infection observed in
a puppy [20] could have occurred during birth or transplacentally. In a well-
designed and controlled study of congenital transmission of leishmaniasis in
dogs from Brazil, no evidence of congenital transmission was found in 63
puppies from 18 naturally infected dogs [21]. This is strong evidence that
congenital transmission is not an eﬀective means of maintaining CVL in the
Brazilian dog population.

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A.C. Rosypal et al / Vet Clin Small Anim 33 (2003) 921–937

Clinical signs of visceral leishmaniasis in dogs

In endemic areas, there are no diﬀerences in the prevalence of infected

male or female dogs. In general, all breeds appear to be equally susceptible,
although North African Ibizian hounds may be naturally resistant [22].

Fig. 1. Life cycle of Leishmania infantum.

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A.C. Rosypal et al / Vet Clin Small Anim 33 (2003) 921–937

There is some indication that short-haired dogs may have a higher
prevalence than long-haired dogs in the same area [23]. Dogs kept outdoors
or that are frequently outdoors are more likely to be infected because of
more exposure to infected sand ﬂies.

Clinical signs of canine leishmaniasis are variable [24,25]. Many dogs are

naturally resistant to disease and appear clinically normal despite being

Fig. 2. Scanning electron micrograph of promastigotes of Leishmania infantum from culture.
Note the dividing promastigotes (D). Bar

¼ 2 lm.

Fig. 3. Transmission electron micrograph of amastigotes (a) of the LIVT-1 strain of Leishmania
infantum

in the spleen of an experimentally infected mouse. A kinetoplast (arrowhead ) and

rudimentary ﬂagellum (arrow) are labeled in one amastigote. Bar

¼ 1 lm.

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A.C. Rosypal et al / Vet Clin Small Anim 33 (2003) 921–937

infected. These dogs may show only a localized nodular reaction at the site of
the sand ﬂy bite. In endemic areas, it is believed that only 10% of infected
dogs develop overt clinical disease [24]. Clinical signs include local or gen-
eralized lymphadenopathy, alopecia, cutaneous lesions, splenomegaly,
onychogryphosis, epistaxis, emaciation, ocular lesions, renal failure,
lameness, diarrhea, and onychogryphosis. Body temperature is usually
normal or below normal. Canine leishmaniasis is a slowly progressive disease.

Immunity to canine visceral leishmaniasis

Dogs naturally or experimentally infected with L infantum develop

a spectrum of disease ranging from asymptomatic to oligo- or poly-
symptomatic. The outcome of disease in CVL is largely mediated by the
development of the cell-mediated immune (CMI) response. Resistance to
canine leishmaniasis is associated with a strong Th1 type cellular immune
response. Analysis of cytokines from peripheral blood mononuclear cells
has revealed that asymptomatic dogs develop a Th1 type response marked
by increased secretion of interleukin-12, tumor necrosis factor (TNF), and
interferon-c (IFN-c) compared with symptomatic dogs [26,27]. Resistant
dogs typically have low or unapparent speciﬁc antibody production and
positive leishmanin skin test [28]. Conversely, the lack of an appropriate
CMI results in disease progression in symptomatic dogs. Symptomatic
dogs develop a marked humoral response and simultaneous lack of
peripheral blood mononuclear cell proliferation in vitro to Leishmania
antigens [29].

CD4+T-cells play a key role in immunity to leishmaniasis by inﬂuencing

the production of a particular cytokine proﬁle and by interacting with
infected macrophages. Symptomatic dogs have decreased levels of CD4+
T-cells compared with noninfected and asymptomatic dogs. These decreased
levels are responsible for the lack of CMI in susceptible dogs [29–31].
Moreover, infected macrophages can alter the canine immune response,
because they are deﬁcient in costimulatory molecules that are required for
T-cell activation [28]. Thus, infected macrophages have a reduced ability to
interact with T cells, and this reduced ability subsequently interferes with
initiation of IFN-c production and parasite destruction.

Diagnosis and antibody responses

The diagnosis of CVL is often diﬃcult because of the variability of

clinical presentation. The simplest method is to demonstrate amastigotes
(Fig. 4) in stained smears from skin lesions, bone marrow smears, or
aspirates from enlarged lymph nodes. Unfortunately, this method will detect
only about 60% (bone marrow) to 30% (lymph nodes) of cases [24].

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A.C. Rosypal et al / Vet Clin Small Anim 33 (2003) 921–937

Histological examination of biopsies also can be used, but this suﬀers from
low sensitivity. Parasites can be isolated in culture of clinical samples, but
this method usually is limited to research and diagnostic laboratories.

Serological methods can be used to detect antibodies. The indirect

ﬂuorescent antibody test (IFAT) is the ‘‘gold standard’’ by which other
serological tests are measured. Serological examinations by IFAT are done
at initial dilutions of serum at 1:16. An IFAT cutoﬀ titer of 1:64 usually is
considered positive. Leishmania parasites, however, have been isolated from
CVL dogs with IFAT titers of 1:16.

Dogs that initially present with clinical signs of CVL demonstrate high

levels of IgG1 and IgG2 by IFAT or ELISA [32–34]. Levels of these isotypes
are variable in asymptomatic dogs [34]. With treatment, IgG1 levels
decrease, while IgG2 titers remain relatively constant. Regardless of the
health status, however, the levels of IgG1 antibodies are usually lower than
those of IgG2 [32].

Other serological methods include complement ﬁxation, indirect hemag-

glutination, latex agglutination, direct agglutination of amastigotes or
promastigotes, FAST agglutination, counter-immunoelectrophoresis, whole
parasite, antigen-speciﬁc ELISA, colloidal gold immunoassay, Western blot,
and rK39 antigen dipstick assay [24,35]. In areas where Trypanosoma cruzi is
also present, there is a possibility of serological cross-reactivity in many of
these tests. This may be overcome by using an rK39 based immunoassay,
because it is based on a Leishmania-speciﬁc amastigote protein. Immuno-
assays based on the rK39 antigen also are good indicators of active CVL
and HVL [36]. A rK39 antigen assay is commercially available in other
countries (Leishmania RAPYDTEST, Intersep, Workingham, United
Kingdom). The Western blot test may be helpful in monitoring speciﬁc
responses to treatment of CVL cases [37]. There is a decrease in the intensity

Fig. 4. Impression spear of amastigotes of the LIVT-1 strain of Leishmania infantum in the
spleen of an experimentally infected mouse. An arrow points to a group of amastigotes.
Bar

¼ 10 lm.

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A.C. Rosypal et al / Vet Clin Small Anim 33 (2003) 921–937

of reactive promastigote bands, especially those in the region of 12 to 30 kd,
after treatment.

Polymerase chain reaction (PCR) can be used to detect Leishmania in

blood, skin biopsies, and lymph node and bone marrow aspirates [38]. Prev-
alence of infection is usually higher using PCR methods than serolog-
ical methods, and many dogs are PCR-positive and serologically negative
[39]. PCR is powerful, but a false-negative test can result if few parasites
are present. Bone marrow is the tissue of choice, followed by lymph node
and blood. Animal inoculation (usually hamsters) and in vitro culture also
can be conducted, and they are about 80% as sensitive as PCR using bone
marrow [40].

Abnormal biochemical features also may be helpful in conﬁrming

a diagnosis of leishmaniasis. Table 3 lists common biochemical alterations
seen in 139 dogs with leishmaniasis [34].

Treatment

Treatment of CVL is challenging, because few dogs are parasitologically

cured [41,42]. This means that many dogs will relapse within a few months
when treatment is stopped, because all the amastigotes have not been killed.
The pentavalent antimonials, meglumine antimonite (Glucantime) or
sodium stibogluconate (Pentostam), are used commonly to treat CVL.
Allopurinol (Zyloric) is used for maintenance to prevent relapse or in
combination with one of the antimonials in initial treatment. There is some
evidence that resistance to antimonials develops in dogs.

In the United States, Pentostam is available from the Centers for Disease

Control and Prevention (CDC). Pentostam is an aqueous solution of 330
mg/mL of agent, which is equivalent to 100 mg/mL pentavalent antimony.
Sodium stibogluconate and meglumine antimonate are administered on the
basis of their antimony content. Antimony compounds are eliminated faster

Table 3
Biochemical features in 139 dogs with leishmaniasis

Feature

Percentage of doses (%)

Ratio of albumin/globulins \0.59

63.2

Hypoalbuminemia

60.1

Hypergammaglobulinemia

57.8

Hypergammaglobulinemia and hypoalbuminemia

52.0

High total serum protein

49.2

Azotemia

24.2

Hypercreatinemia

18.7

Data from

Solano-Gallego L, Riera C, Roura X, Iniesta L, Gallego M, Valladares JE, et al.

Leishmania infantum

-speciﬁc IgG, IgG1 and IgG2 antibody responses in healthy and ill dogs

from endemic areas. Evolution in the course of infection and after treatment. Vet Parasitol
2001;96(4):265–76.

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A.C. Rosypal et al / Vet Clin Small Anim 33 (2003) 921–937

if given intramuscularly (IM) than subcutaneously (SC) or intravenously
(IV) in dogs, and it is important to maintain serum levels of the compound
to treat leishmaniasis. Additionally, IM injection has the potential to cause
severe adverse eﬀects, including muscular ﬁbrosis and abscess formation.
For these reasons, IM injection of antimonials is not recommended.
Pentavalent antimonials are relatively well-tolerated. Adverse reactions
include pain at the injection site, gastrointestinal (GI) symptoms, delayed
muscle pain, and joint stiﬀness. Canine leishmaniasis is treated with 30 to 50
mg/kg body weight pentavalent antimony in the form of sodium stiboglu-
conate by IV or SC injection administered daily for 3 to 4 weeks [42]. Relapses
may occur a few months to 1 year after treatment and should be treated with
another round of pentavalent antimony. The use of pentavalent antimonials is
contraindicated in patients with myocarditis, hepatitis, or nephritis.

Allopurinol is an attractive alternative to antimonial treatment of CVL

[43]. Allopurinol interferes with normal protein synthesis in Leishmania
parasites. Like other Leishmania treatments, it does not remove all of the
parasites [44]. Allopurinol given orally at 10 to 30 mg/kg once or twice daily
is eﬀective in producing clinical cures but relapses are common once therapy
is discontinued. Oral treatment with 20 mg/kg allopurinol for 1 week
a month is highly eﬀective in preventing relapse in dogs [45].

Combination therapy of antimonials plus allopurinol may be superior to

either agent used alone in the initial treatment of CVL [42]. The antimonials
are given SC at the regular dosage levels for 3 to 4 weeks along with
allopurinol twice daily at 15 to 20 mg/kg. Allopurinol then is continued for
maintenance therapy.

Several other chemotherapeutic agents have been used in the treatment of

CVL including pentamidine, paromomycin, and several formulations of
Amphotericin B [41,42]. These may be alternatives to antimonials, especially
if antimonial drug resistance is suspected.

Preventing infection in dogs by insecticide-impregnated collars
and vaccination

Recent studies have indicated that the use of deltamethrin-impregnated

collars will reduce transmission of L infantum infection to dogs [46–49].
Experimental studies indicate that the deltamethrin collars decrease sand ﬂy
bites by 80% to 96% [46,47]. Field studies using the deltamethrin collars in
dogs in an endemic area of southern Italy found a reduction in serocon-
version rates from 50% to 86% [48]. A study conducted in Iran found that
the use of deltamethrin collars on dogs signiﬁcantly decreased the serocon-
version rates in dogs and children living in villages with collared dogs [49].

Vaccination of dogs against leishmaniasis would beneﬁt the canine and

human populations in endemic areas. A recent report reviewed the current
knowledge and status of canine Leishmania vaccines, and it was suggested

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A.C. Rosypal et al / Vet Clin Small Anim 33 (2003) 921–937

that the main requirement for an eﬀective Leishmania vaccine is the in-
duction of a stable and long-lasting TH1 type CMI response [50]. Attenu-
ated or otherwise genetically modiﬁed live vaccines often show promise
in animal models [51] but because of safety and other limitations are more
valuable as tools for deﬁning immune responses than actual vaccines.
Killed promastigote antigens with BCG adjuvant have shown promise
in some studies, but complete protection has not been achieved [50,51].
Deﬁned protein, recombinant protein, and DNA vaccines have been devel-
oped and tested [50,51]. Eﬃcacy of these vaccines depends on the antigen
chosen and often on the type of TH1 and type of CMI-inducing adjuvant
administered. In a recent phase III ﬁeld trial in Brazil, dogs were vaccinated
with a Leishmania fucose mannose ligand (FML) vaccine [52]. The results
of the FML vaccine trials were encouraging in that 92% of vaccinated
dogs were protected, and 8% developed only mild signs of CVL.

Canine culling programs for preventing human infection

In Brazil, culling of Leishmania IFAT-positive dogs has been used as

a means of reducing HVL in people [2]. The eﬀectiveness of culling
programs has been debated for many years [53], but current opinion is that
culling IFAT antibody positive dogs is of little value [54]. The two most
important reasons for failure of culling programs to control CVL in endemic
areas and decrease HVL are lack of sensitivity of the IFAT and other
serological tests and a delay in culling positive dogs, which usually occurs
between 80 and 180 days of sampling [54]. These ﬂaws permit sand ﬂy
transmission to continue from these dogs and hampers the control
programs. It has been suggested that the use of deltamethrin collars could
replace dog culling because of the signiﬁcant impact they have on human
transmission.

History of endemic canine visceral leishmaniasis in the United States

The ﬁrst case of endemic CVL was reported in a 7-year-old female

foxhound from Oklahoma [55]. This dog had generalized alopecia
associated with Demodex folliculorum infestation. The dog had never been
outside the United States and had been conﬁned to a 150 mile radius around
Oklahoma City. This dog came from a kennel of 17 foxhounds, and
additional dogs in the kennel were found to be infected [56]. The second case
of endemic CVL came from an English foxhound in an Ohio research
colony [57]. A male dog developed CVL and eventually died. Both its dam
and sire were born in the United States. Serological examination of 25 dogs
from the colony revealed that eight had antibodies to Leishmania. Other
isolated cases of leishmaniasis in dogs with no history of foreign travel oc-
curred in a pet Basenji in Texas [58] and a pet toy poodle in Maryland [59].

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A.C. Rosypal et al / Vet Clin Small Anim 33 (2003) 921–937

In early 2000, leishmaniasis was diagnosed in foxhounds from a hunt club

in New York [5,60]. Beginning in the late summer of 1999, a number of
foxhounds at the hunt club had developed signs of disease including
bleeding, wasting, seizures, hair loss, skin lesions, kidney failure, and
swollen limbs and joints. Several dogs died. Some of the aﬀected dogs were
sent to North Carolina State University College of Veterinary Medicine for
diagnostic studies, and cytopathologic examination of joint ﬂuid from one
hound revealed amastigote forms of Leishmania species. This ﬁnding was
conﬁrmed at necropsy of several dogs, and promastigotes were isolated and
grown in culture. Diagnostic studies at the New York hunt club kennel
revealed that 39 of 93 (42%) foxhounds were seropositive for antibodies to
Leishmania

. Culture of aspiration or biopsy material from lymph nodes and

other tissues of 15 seropositive dogs resulted in isolation of Leishmania
species promastigotes from 15 dogs. Genetic typing of the parasites at the
Institute of Public Health in Rome found them to belong to the L. infantum
MON1 zymodeme, which is the most common type isolated from HVL
cases in the Mediterranean [5]. The authors have obtained three isolates of
L. infantum

from three naturally infected foxhounds from Virginia by

culture of bone marrow and lymph nodes. They have transmitted one of
these strain of L infantum to mice and beagle dogs experimentally [61] and
deposited type parasite cultures in the American Type Culture Collection,
Manassas, VA (LIVT-2 strain ATTC# 5D918 and LIVT-3 ATTC strain
#5D919). Because the potential for selection of these American isolates for
nonvector transmission, the authors are focusing research eﬀorts on ex-
amining transmission of these parasites in dogs by maternal and other
nonvector means.

In view of the sporadic cases seen in hounds in the past 20 years in the

US, the CDC have undertaken a large scale survey of foxhounds in the
United States. Since early 2000, sera from more than 10,000 foxhounds and
other hunting dogs in 35 states and Canada have been tested [60]. High
IFAT titers of at least 1:64, indicative of active infection, have been found in
approximately 2% of foxhound samples, although some culture-positive
dogs had lower titers. Seropositive dogs have been detected in 60 kennels in
22 states and two Canadian provinces. Serological testing of 600 pet dogs
and 300 wild canids from geographic localities close to the infected
foxhounds have not yielded positive animals [60], and no cases of
autochthonous HVL have been reported in the United States.

The means by which infection is transmitted in dogs in the United States

is not understood. Sand ﬂies are distributed widely in much of the United
States, although they usually are not regarded as an important human or
livestock pests. One species, L shannoni, has been infected with L infantum
by feeding on symptomatic CVL dogs [14]. L shannoni is distributed widely
through the eastern United States as far north as New Jersey, and evidence
indicates that this species may be a competent vector [13] of another
Leishmania

species, L mexicana. Much of the epidemiological data collected

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A.C. Rosypal et al / Vet Clin Small Anim 33 (2003) 921–937

do not support a major role for sand ﬂies in transmission of CVL in the
United States. The management and patterns of movement of foxhounds
have led to the suggestion that direct dog-to-dog contact may be more
important than sand ﬂy transmission in the United States. The risk of CVL
to people in the United States has not been established. Cases of HVL
originating in the United States have not been reported. Sand ﬂies are not
a familiar human pest, and although direct transmission of the organism
from dogs is theoretically possible, there are no cases reported. Even in the
Mediterranean region, where 20% of dogs are infected, there are few clinical
cases in people. People most at risk of developing disease are those with
weakened immune systems.

Human visceral leishmaniasis in the immunosuppressed patient

Of concern is the possible establishment of human leishmaniasis in the

United States and the emerging importance of leishmaniasis as an
opportunistic infection in cases of HIV infection. Recently HVL has
emerged as an important opportunistic infection of individuals coinfected
with HIV in endemic areas. Coinfection is increasingly more common be-
cause of overlapping geographical distribution, as HIV spreads to rural
areas, and leishmaniasis spreads from rural to more urban environments.
Immunocompetent people frequently are bitten by Leishmania-infected sand
ﬂies but do not develop overt disease. In people dually infected with HIV
and Leishmania, however, leishmaniasis rapidly develops into a severe and
life-threatening disease. HIV and Leishmania are able to infect and destroy
macrophages [62]. Coinfection produces a cumulative immune deﬁciency
that hastens the onset and severity of both diseases [63]. Following initial
infection, both pathogens are able to develop latent infection [62], but it is
unclear if the severity of leishmaniasis in coinfected patients is caused by
reactivation of latent infection or primary infection [64].

Diagnosis of HVL in HIV patients is challenging because of overall

immune depression that results from dual infection. The typical clinical
signs (fever, hepatosplenomegaly, and lymphadenopathy) are not always
present in coinfected individuals, and diagnosis may be confused further by
presentation of atypical sites of lesions such as the lungs and GI tract [64].
Impairment of the immune system also decreases antibody production.
Thus, serologic diagnosis of HVL in patients with HIV yields a high
proportion of false negatives and should be conﬁrmed by bone marrow
aspirate or culture [65]. Also, coinfected patients exhibit peripheral
parasitemia caused by uncontrolled multiplication of the parasite, and as
a result they can act as a reservoir host for infection of sand ﬂies or
intravenous drug users [62,63].

Treatment of immunocompromised patients is aimed at clinical and

parasitological cure, although relapses are more common and more frequent

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A.C. Rosypal et al / Vet Clin Small Anim 33 (2003) 921–937

in coinfected patients. Treatment usually consists of pentavalent anti-
monials, alone or in combination with amphotericin B, followed by pro-
phylactic

treatment

with

antimony,

allopurinol,

pentamidine,

amphotericin B [66]. Eﬀective treatment results in clinical cure, but rarely
parasitological cure because of latently infected cells. Relapse of leishman-
iasis occurs as the HIV patient’s immune system weakens, and dormant
infection is reactivated [64]. Since the introduction of highly active
antiretroviral therapy (HAART), the incidence of most AIDS-associated
opportunistic infections has decreased signiﬁcantly and is likely the result of
increased CD4+ T lymphocytes [67]. Leishmaniasis is not considered an
AIDS-deﬁning illness, although HVL does occur commonly in patients with
HIV. HAART appears to reduce the incidence of VL in coinfected
individuals [68,69].

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Index

Note:

Page numbers of article titles are in boldface type.

American canine hepatozoonosis, 905–920.

See also Hepatozoon americanum;
Hepatozoonosis, canine.

Amphotericin B lipid complex, update on,

749–751

Anaplasma

spp

A phagocytophila,

in pet animals,

870–871

A platys,

in pet animals, 871

Antifungal therapy, update on, 749–758

amphotericin B lipid complex, 749–751
azoles, 751–753
chitin synthase inhibitors, 755
echinocandins, 754
ﬂuconazole, 752–753
fungal protein synthesis inhibitors, 755
b-glucan synthase inhibitors, 754
itraconazole, 752
lufenuron, 755
nikkomycins, 755
pneumocandins, 754
terbinaﬁne, 754
triazoles, 753

Azole(s), update on, 751–753

Babesia

spp

B canis,

canine babesiosis due to, 885,

886–888

clinical signs of, 894

B gibsoni,

canine babesiosis due to,

888–890

clinical signs of, 894–895

canine babesiosis due to, 885–890

Babesiosis

canine, 885–904

B canis

and, 885, 886–888

B gibsoni

and, 888–890

clinical pathology of, 896–897
clinical signs of, 894–895
coinfection with, 895
described, 885

diagnosis of, 895–896
pathogenesis of, 890–893
prevention of, 900–901
transmission of, 885
treatment of, 897–899
vaccines for, 899

human, 900

Bartonella

spp, described, 809–810

Bartonellosis, 809–825

causes of, 809–810
diagnosis of, 814–816
epidemiology of, 810–811
in cats

clinical ﬁndings of, 813–814
pathogenesis of, 811–812
prevention of, 817–818
treatment of, 816–817

in dogs

clinical ﬁndings of, 814
pathogenesis of, 812–813
treatment of, 817

pathogenesis of, 811–813
prevention of, 817–818
public health issues related to,

818–819

treatment of, 816–817

Basidiobolomycosis, 707

Blastomycosis

causes of, 721–722
clinical signs of

in cats, 723
in dogs, 723

described, 721–722
diagnosis of, 723–725
prognosis of, 728
treatment of, 725–728
update on, 721–728

Borrelia burgdorferi

borreliosis due to, 827
described, 834
detection of, 842–848
DNA of, 834–835
exposure to, 832–837
genes in, 835–836

Vet Clin Small Anim

33 (2003) 939–943

0195-5616/03/$ - see front matter

doi:10.1016/S0195-5616(03)00074-3

Borreliosis. See also Borreliosis, canine.

Borrelia burgdorferi

and, 827

canine, 827–862

diagnosis of, 849–850
diﬀerential diagnosis of, 849–850
experimental, 839–842
historical background of, 827,

830

prevention of, 852–853
spontaneous, 837–839
treatment of, response to,

850–852

vaccines for, 853–855
vs. human borreliosis, 828–829

historical background of, 827, 830
human

clinical signs of, 837
vs. canine borreliosis, 828–829

Calcivirus, feline, 759–772. See also Feline

calicivirus (FCV).

Canine babesiosis, 885–904. See also

Babesiosis, canine.

Canine borreliosis, 827–862

Canine culling programs, in canine visceral

leishmaniasis prevention, 931

Canine visceral leishmaniasis, 921–937.

See also Leishmaniasis, canine visceral.

Cat(s)

bartonellosis in

clinical ﬁndings of, 813–814
pathogenesis of, 811–812
prevention of, 817–818
treatment of, 816–817

blastomycosis in, clinical signs of, 723
coccidiomycosis in, clinical signs of,

734

cryptococcosis in, clinical signs of,

738–739

fungal diseases in, update on, 721–747.

Document Outline

00 Erratum
01 Preface
02 Update on molecular techniques for diagnostic testing of infectious disease
- Update on molecular techniques for diagnostic testing of infectious disease
03 Pythiosis, lagenidiosis, and zygomycosis in small animals
- Pythiosis, lagenidiosis, and zygomycosis in small animals
04 Update on canine and feline fungal diseases
- Update on canine and feline fungal diseases
  - Blastomycosis
    - Clinical signs
      - Dogs
      - Cats
    - Diagnosis
    - Treatment
    - Prognosis
  - Histoplasmosis
    - Clinical signs
      - Dogs
      - Cats
    - Diagnosis
    - Treatment
    - Prognosis
  - Coccidiomycosis
    - Clinical signs
      - Dogs
      - Cats
    - Diagnosis
    - Treatment
    - Prognosis
  - Cryptococcosis
    - Clinical signs
      - Cats
      - Dogs
    - Diagnosis
    - Treatment
    - Prognosis
  - Drug therapy for systemic mycoses
  - Summary
  - References
05 Update on antifungal therapy
- Update on antifungal therapy
06 Update on feline calicivirus_new trends
- Update on feline calicivirus: new trends
07 Feline hemotropic mycoplasmosis (feline hemobartonellosis)
- Feline hemotropic mycoplasmosis (feline hemobartonellosis)
08 Leptospirosis_A re-emerging zoonotic disease
- Leptospirosis - A re-emerging zoonotic disease
09 Bartonellosis
- Bartonellosis
  - Epidemiology
  - Pathogenesis
    - Cats
    - Dogs
  - Clinical findings
    - Cats
    - Dogs
  - Diagnosis
  - Treatment
  - Prevention
    - Cats
    - Dogs
  - Public health
  - Summary
  - References
10 Canine borreliosis
- Canine borreliosis
11 Ehrlichiosis and related infections
- Ehrlichiosis and related infections
12 Canine babesiosis
- Canine babesiosis
13 American canine hepatozoonosis
- American canine hepatozoonosis
14 Canine visceral leishmaniasis and its emergence in the United States
- Canine visceral leishmaniasis and its emergence in the United States
15 Index

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