Applied and Environmental Microbiology, August 1998, p. 2888-2893, Vol. 64, No. 8
0099-2240/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
Detection of Ehrlichia risticii, the Agent of Potomac
Horse Fever, in Freshwater Stream Snails (Pleuroceridae:
Juga spp.) from Northern California
Jeffrey E.
Barlough,1
Gerhard H.
Reubel,1
John E.
Madigan,1,*
Larisa K.
Vredevoe,2
Paul E.
Miller,3 and
Yasuko
Rikihisa4
Department of Medicine and Epidemiology,
School of Veterinary Medicine,1 and
Department of Entomology,2 University of
California, Davis, California 95616;
19305 Ordway Road, Weed,
California 960943; and
Department of
Veterinary Biosciences, College of Veterinary Medicine, The Ohio
State University, Columbus, Ohio 432104
Received 5 November 1997/Accepted 6 May 1998
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ABSTRACT |
Ehrlichia DNA was identified by nested PCR in operculate snails
(Pleuroceridae: Juga spp.) collected from stream water in a
northern California pasture in which Potomac horse fever (PHF) is
enzootic. Sequencing of PCR-amplified DNA from a suite of genes (the
16S rRNA, groESL heat shock operon, 51-kDa major antigen genes) indicated that the source organism closely resembled
Ehrlichia risticii, the causative agent of PHF. The minimum
percentage of Juga spp. harboring the organism in the
population studied was 3.5% (2 of 57 snails). No ehrlichia DNA was
found in tissues of 123 lymnaeid, physid, and planorbid snails
collected at the same site. These data suggest that pleurocerid stream
snails may play a role in the life cycle of E. risticii in
northern California.
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INTRODUCTION |
Potomac horse fever (PHF), also
called equine monocytic ehrlichiosis, is an important disease of
horses caused by a monocytotropic rickettsia, Ehrlichia
risticii (19-23, 27, 34). PHF was first observed
in 1979 in pastures along the Potomac River in Maryland (20). Soon thereafter it was reported in Virginia and
Pennsylvania, and it has since been identified in a number of other
states and in Europe (27). Clinical signs include anorexia,
lethargy, variable fever, and diarrhea. Laminitis is a complication in
a significant percentage of cases. Fatalities may result if severely
affected horses are not treated promptly with fluids, electrolytes, and antibiotics.
The means by which horses become exposed to E. risticii has
remained mysterious. The disease is infectious but not contagious; this, combined with its seasonality, geographical distribution, and
ease of transmission by intradermal inoculation, has been thought to
reflect spread by an arthropod vector, possibly a tick, but direct
evidence of tick transmission is lacking (3, 18, 27, 29, 30,
37). The disease can be transmitted by oral inoculation, however,
and the causative agent has been identified in feces (25, 28,
35).
Since the earliest reports of PHF, an association of cases with
riverine and other aquatic habitats has been noted. Research has
revealed a close phylogenetic relationship between E. risticii and the following three rickettsiae associated with
aquatic environments: Neorickettsia helminthoeca, the agent
of "salmon poisoning," a frequently fatal systemic,
metacercaria-borne disease of canids; the SF agent, isolated in
Japan from trematode metacercariae (Stellantchasmus falcatus) parasitic on gray mullet fish; and Ehrlichia
sennetsu, the agent of human sennetsu ehrlichiosis in Japan and
Malaysia (15, 16, 31, 33, 43). Based on 16S rRNA gene
sequences, antigenic cross-reactivity, and the proven or suspected
mechanism of transmission, these four agents form a distinct cluster or genogroup distinct from other rickettsiae (11, 33, 36, 43).
These data suggest that the vector of E. risticii may not be
an arthropod but instead an organism closely associated in some way with river and/or irrigation water. Accordingly, we
have directed our efforts recently toward identifying a
potential aquatic reservoir for E. risticii. In
this report we describe the detection of ehrlichia DNA in freshwater
operculate snails (Pleuroceridae: Juga spp.) collected from stream water in a pasture in northern California in
which PHF is enzootic. Analysis of PCR-amplified DNA
indicated that the ehrlichia detected is clearly related to the type
strain of E. risticii and virtually identical to a new
strain (the strain responsible for Shasta River crud [SRC])
identified in the blood buffy-coat cells of an affected horse a few
miles from the pasture which we studied. We hypothesize that operculate
snails of the genus Juga may play a role in the life cycle
of E. risticii in northern California.
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MATERIALS AND METHODS |
Snail collection.
Freshwater snails were collected in August
1996 from a pasture in Weed, Calif. (Siskiyou County), with a history
of PHF in some horses allowed to graze in it. No case had been observed for at least several months prior to sampling. Snails were collected by
hand or with a net from the clear, shallow margins of a stream that
flows through the pasture and is accessible to horses for drinking. The
stream is fed by the nearby Shasta River. A total of 180 snails were
collected, including 55 lymnaeids, 24 physids, 44 planorbids, and 57 pleurocerids (Table 1). The pleurocerid snails (Fig. 1) were identified as
members of the genus Juga (5). Based on current
nomenclature derived from classical shell characteristics, the majority
of the individuals resembled the subspecies Juga hemphilli
hemphilli, in which ribs (costae) are for the most part limited to
the apical whorl of the shell (5). However, because the
systematics of the Pleuroceridae is in need of thorough revision (5, 10), identification of the snails to the species level remains presumptive.
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TABLE 1.
E. risticii nested PCR results (16S rRNA
gene fragment) for freshwater stream snails collected at a site in
northern California in which PHF is enzootic
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FIG. 1.
Pleurocerid snails collected from stream water fed by
the Shasta River in Weed, Calif. (Siskiyou County). Bar = 1 cm.
(Photograph courtesy of Jerry Fields.)
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Processing of snails.
The snails collected were frozen
(
70°C), thawed, segregated by family into pools of 2 to 11 snails
depending on size, and processed for PCR by a modification of previous
methods (2, 3). Briefly, snails were washed in sterile
distilled water, dissected from their shells with sterile scissors, and
placed into 2-ml microtubes. The tissues were mechanically disrupted for 1 min with a Mini-BeadBeater (Biospec Products, Bartlesville, Okla.) by using the high setting. The tubes were centrifuged at the
maximum speed in a microcentrifuge for 1 min. Excess crude extract was
removed until approximately 1 ml remained in each microtube. The tubes
were filled nearly to the top with DNA extraction buffer (10 mM Tris
[pH 8.0], 2 mM EDTA, 0.1% sodium dodecyl sulfate, 500 µg of
proteinase K per ml), vortexed, placed in a heating block (57°C) for
3 h, vortexed again, and then heated at 97°C for 15 min to
inactivate the proteinase K.
After thorough vortexing, 500-µl portions of each extract were
transferred to sterile 1.7-µl Eppendorf tubes, and DNA extraction was
performed by using a phenol-chloroform-isoamyl alcohol (25:24:1) mixture. Each sample was extracted twice. Portions (500 µl) of the
final aqueous layer were transferred to fresh microtubes. The DNA was
precipitated by adding 3 to 4 volumes of absolute ethanol and 100 µl
of 3 M sodium acetate (pH 5.2) to each tube and incubating the tubes at
20°C for 24 to 48 h. The DNA was pelleted by centrifugation at
the maximum speed in a microcentrifuge for 15 min at 4°C and washed
twice with 70% ethanol. After the DNA was dried, it was resuspended in
100 µl of Tris-EDTA buffer (pH 8.0). The DNA content was checked with
a UV spectrophotometer, and the DNA concentration of each sample was
adjusted to approximately 300 ng per µl. One microliter was used in
each PCR mixture.
Nested PCR assays.
A nested PCR that amplifies a 5' segment
(529 bp) of the 16S rRNA gene of E. risticii was used as an
initial screening procedure for infected snail pools. The components
and conditions of this PCR procedure have been described in detail
elsewhere (3). The cycling parameters used in this study
were as follows: preheating at 94°C for 5 min and then 35 cycles
consisting of denaturation at 94°C for 1 min, annealing at 60°C for
2 min, and extension at 72°C for 1.5 min, followed by a final
extension step at 72°C for 7 min. The PCR products were visualized in
ethidium bromide-stained 1.5% agarose minigels.
Segments of two additional ehrlichia genes were amplified by PCR. Sets
of nested primers were designed to detect portions of the E. risticii homolog of the Escherichia coli groESL heat shock operon (39) and the E. risticii 51-kDa major antigen gene (12, 13, 41). The
following primer sequences were used to amplify the heat shock
operon: 5'-ACCAGGCTACCTCACAGGC-3' and
5'-TTGACCCTCGCATCAATG-3' (outer primers); and
5'-CACAAGTTGGTTCAATTTCTGC-3' and
5'-CCGAGATCTTCAACAGTAAGGC-3' (inner primers). The amplified
sequence was entirely contained within the groEL portion of
the operon. The following primer sequences were used to
amplify the 51-kDa major antigen gene: 5'-GGATCGATAACTGCGATGCT-3' and 5'-ACCGGCCTGACCACTAAAG-3' (outer primers); and
5'-TCCTATAATGGCACCACTAGCG-3' and
5'-CCATCCGCAGTAGAGTTTGAG-3' (inner primers).
The predicted molecular sizes for the groESL fragment were
823 bp (first-round product) and 526 bp (nested product). The nested product comprised nucleotides 500 through 1025 of the operon
(numbering based on the sequence deposited under GenBank accession no.
U96732). The predicted sizes for the 51-kDa major antigen gene fragment were 818 bp (first-round product) and 569 bp (nested product). The
nested product comprised nucleotides 1303 through 1871 of the gene
(numbering based on the sequence deposited under GenBank accession no.
U85784). The components and conditions of the PCR assays used for
groESL and the 51-kDa major antigen gene were similar to the
components and conditions used for standard 16S rRNA gene
amplification, except that the annealing temperatures were 45°C (for
the 51-kDa major antigen gene) and 55°C (for groESL).
Cloning and sequencing of amplified PCR products.
A nearly
complete sequence for the 16S rRNA gene was obtained by amplifying and
cloning the gene in three overlapping fragments. The majority of the
gene (ca. 1,440 bases) was amplified in the first round by using
primers ER-3 (5'-ATTTGAGAGTTTGATCCTGG-3') (3, 8)
and PC5 (5'-TACCTTGTTACGACTT-3') (44). In
the nested round the 5' segment of the gene was amplified with
primers ER-3 and ER-2 (5'-GTTTTAAATGCAGTTCTTGG-3')
(3); the middle segment was amplified with primers
ER-2a(R) (5'-CCCGTAAGTTAGGTGTG-3') and ER-X
(5'-CATCTCACGACACGAGC-3'); and the 3' segment was amplified with primers ER-Y (5'-CCAACACAGGTGTTGC-3') and ER-Z2
(5'-ACCCCAGTCACCCACCCC-3'). The cycling conditions used were
the same as those described above for the standard nested PCR except
that annealing was performed at 52°C and each 72°C extension step
was 2 min long.
Nested PCR products of the 16S rRNA, groESL, and 51-kDa
major antigen genes were purified by spin chromatography
(PCR SELECT-II spin columns; 5 Prime 
3 Prime, Inc.,
Boulder, Colo.) and were cloned by using the pNoTA/T7 shuttle vector
and competent E. coli (Prime PCR Cloner cloning system;
5 Prime 3 Prime). Double-stranded maxiprep DNA was isolated with a
PERFECTprep plasmid DNA preparation kit (5 Prime 3 Prime). One microgram of maxiprep DNA was used for restriction
endonuclease digestion with BamHI (New England Biolabs,
Beverly, Mass.) to verify the insert sizes. The inserts were sequenced
by using the universal M13 sequencing primers present in the vector.
Sequencing was performed with a fluorescence-based automated sequencing
system (Applied Biosystems, Foster City, Calif.). The sequences in both
directions were obtained for all inserts. Three cycles of cloning and
sequencing were performed to verify the sequence and the repeatability
of the procedure.
Nucleotide sequence accession numbers.
Most of the 16S rRNA
gene sequences used were obtained from the GenBank database, and the
accession numbers were as follows: E. risticii Illinois
(type strain), M21290; SF agent, U34280; E. sennetsu,
M73225; N. helminthoeca, U12457; Ehrlichia canis,
M73221; Ehrlichia chaffeensis, U23503; Ehrlichia equi, M73223; Ehrlichia ewingii, M73227; HGE agent,
U02521; Anaplasma marginale, M60313; Cowdria
ruminantium, U03776; Rickettsia rickettsii, U11021; and
Rickettsia prowazekii, M21789. The sequences of the
Kentucky, Ohio 081, and SRC strains of E. risticii were
obtained from previously published sources (21, 42). The
GenBank accession numbers for the groESL and 51-kDa major
antigen gene sequences of E. risticii were U96732
and U85784, respectively. Other relevant groESL
sequences used were the groESL sequences of E. sennetsu (GenBank accession no. U88092), E. chaffeensis (L10917), E. canis U96731 and C. ruminantium (U13638). The groESL and
51-kDa major antigen gene sequences of the SRC agent were
determined in our laboratory at the University of California, Davis.
Phylogenetic analysis of sequence data.
The sequences were
subjected to a BLAST analysis (1) performed with GenBank
nucleic acid sequences to determine similarity ranks, levels of
identity, and deduced amino acid sequences (the latter for the
groESL protein and the 51-kDa major antigen only). Multiple-sequence alignments of the 16S rRNA genes were constructed with CLUSTAL W (40). A phylogenetic analysis was performed
by using the DNAML maximum-likelihood method of PHYLIP (14).
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RESULTS |
Initial PCR screening.
During the initial screening by nested
PCR (5' segment of the 16S rRNA gene), two positive pools, which
consisted of 8 and 10 pleurocerid snails of the genus Juga,
were identified (Fig. 2). The positive
pools were designated Shasta snail-1 and Shasta snail-2. The minimum
percentage of Juga spp. harboring the microorganism in the
population studied was 3.5% (2 of 57 snails). Pools derived from
lymnaeid, physid, and planorbid snails produced negative PCR results.
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FIG. 2.
E. risticii nested PCRs performed for
the 16S rRNA, groEL (of groESL), and 51-kDa
antigen genes by using DNA from the Shasta snail-1 (lanes 1) and Shasta
snail-2 (lanes 2) pools and positive (lanes +) and negative (lanes )
E. risticii DNA controls. Lane  , X174
replicative-form DNA HaeIII digest (molecular size
marker).
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16S rRNA gene.
Nearly complete 16S rRNA genes (length, ca.
1,440 bp) from the two positive snail pools were cloned, sequenced, and
compared to previously described sequences of E. risticii strains (Table 2). Both
Shasta snail-1 and Shasta snail-2 sequences were closely related to the
sequence of the SRC agent, a strain of E. risticii identified in May 1996 in blood buffy-coat cells of an affected horse
at Mt. Shasta City, Calif., a few miles from the pasture where the
snails were collected (21). The Shasta snail-2 sequence differed from the E. risticii sequence by a single
nucleotide (position 828), while the Shasta snail-1 sequence differed
by two nucleotides (positions 36 and 309). The next most closely related sequence was that of E. risticii
Kentucky. As expected, E. sennetsu and the SF
agent, both of which are closely related to E. risticii, exhibited similarity to the snail ehrlichiae (Table 3). N. helminthoeca, the agent
of salmon poisoning, has been found in snails of the genus
Juga but was clearly different from the snail ehrlichiae (94 to 95% identity). The most closely related arthropod-borne agent was
A. marginale (level of identity, ca. 85%).
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TABLE 2.
Nucleotide differences in the 16S rRNA genes of the
Shasta snail-1 and Shasta snail-2 ehrlichiae and four strains of
E. risticii (Illinois, SRC agent, Kentucky, Ohio 081)
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TABLE 3.
Comparison of the Shasta snail-1 and Shasta snail-2
16S rRNA gene sequences with the 16S rRNA gene sequences of
other rickettsiae
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groESL heat shock operon.
The gene
fragments amplified from both snail pools (Fig. 2) were most similar to
groESL of the SRC agent and E. risticii
(Table 4). The sequences of Shasta
snail-1 and the SRC agent were, in fact, identical; the sequence of
Shasta snail-2 differed by a single nucleotide. The next most similar
sequences, those of E. sennetsu, E. chaffeensis, E. canis, and C. ruminantium, differed more. The deduced amino acid sequences of
the GroEL fragments from the SRC agent and Shasta snail-1 were ientical
to each other and to the GroEL fragment of E. risticii,
indicating that the few nucleotide changes were silent (Fig.
3). For Shasta snail-2, the single added
nucleotide resulted in a conservative amino acid substitution (Fig. 3).
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TABLE 4.
Comparison of the Shasta snail-1 and Shasta snail-2
partial groESL gene sequences with the partial
groESL gene sequences of other rickettsiae
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FIG. 3.
Deduced amino acid sequences of GroEL segments. Periods
indicate conserved positions relative to the E. risticii sequence.
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51-kDa major antigen gene.
As expected, the sequences
obtained from the snail pools showed greater variation from
E. risticii in the 51-kDa major antigen gene
sequence than in the more conserved 16S rRNA and groESL
sequences. The nucleotide sequences amplified from Shasta snail-1,
Shasta snail-2, and the SRC agent were identical, and all three
exhibited 91.74% identity to the E. risticii gene
sequence deposited in the GenBank database (derived from a Maryland
strain). An analysis of the deduced amino acid sequences for Shasta
snail-1, Shasta snail-2, and the SRC agent revealed 12 amino acid
changes compared to E. risticii, and 6 of the 12 changes were conservative in nature (Fig.
4).
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FIG. 4.
Deduced amino acid sequences of E. risticii 51-kDa major surface antigen segments. Periods indicate
conserved positions relative to the E. risticii
sequence.
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Phylogenetic analysis of sequence data.
A phylogenetic tree
generated by TreeView PPC from the DNAML analysis of the 16S rRNA gene
sequences is shown in Fig. 5. The sequences of the snail ehrlichiae were found to cluster with the sequence of the SRC agent and were most closely associated with the
Kentucky and Illinois strains of E. risticii.
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FIG. 5.
Phylogenetic tree generated by GrowTree from a
DNAML-based alignment of 16S rRNA gene sequences (length, ca. 1,400 bp), showing the relationship of the Shasta snail-1 and Shasta snail-2
ehrlichiae to other rickettsiae. ER, Ehrlichia risticii.
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DISCUSSION |
Here we describe identification of ehrlichia DNA in two pools of
pleurocerid snails of the genus Juga collected from stream water in a pasture in Siskiyou County, Calif., in which PHF is enzootic. Determination of the 16S rRNA gene sequences ruled out the
possibility that the organism detected was N. helminthoeca, the agent of salmon poisoning, which is related to E. risticii and occurs in snails belonging to the family
Pleuroceridae (24, 31). The 16S rRNA gene sequences of both
Shasta snail-1 and Shasta snail-2 were virtually identical to the 16S
rRNA gene sequence of the SRC agent and were clearly similar to the 16S
rRNA gene sequences of other variants of E. risticii
reported previously (42). This result was mirrored by the
results obtained for the DNA sequences and deduced amino acid sequences
of the less conserved groESL heat shock operon and
the E. risticii 51-kDa major antigen gene. These data
provide evidence that pleurocerid stream snails may play a role in the
life cycle of E. risticii and the transmission of PHF
in northern California.
Sequences of 16S rRNA genes are known to vary in an orderly fashion
throughout the phylogenetic tree and thus are desirable targets for PCR
amplification and relatedness testing (26, 44, 45). The
variation observed in the 16S rRNA gene sequences obtained from the two
snail pools was equal to or less than the variation observed in the
sequences of other strains of E. risticii associated with disease in horses (8, 21, 42). This information by itself normally would be sufficient to relate the snail ehrlichiae to
E. risticii.
To investigate this relationship further, we sequenced portions of two
additional genes, the groESL heat shock operon and the 51-kDa major antigen gene; the latter represents a DNA sequence more specific for E. risticii. Like the 16S rRNA gene,
groESL is considered to be a useful molecular clock for
phylogenetic studies (17). We found that the sequences of
amplified groESL fragments from the snail ehrlichiae and the
SRC agent differed by either eight or nine nucleotides from the
homologous sequences of E. risticii. The deduced amino
acid sequences, however, were identical or nearly identical to the
amino acid sequence of E. risticii, indicating that the
nucleotide changes in the snail ehrlichiae were for the most part
silent. Thus, the GroEL segments of E. risticii, the
snail ehrlichiae, and the SRC agent were virtually indistinguishable,
providing further evidence of relatedness.
Although 16S rRNA gene and groESL sequences are
valuable in studies of phylogenetic relationships, the variation
inherent in other genes, particularly the genes coding for
antigenic proteins, is more likely to affect such important properties
as host range, virulence, and vector competence. Accordingly, we
investigated the 51-kDa major antigen gene of E. risticii, whose sequence is not significantly related to any other
sequence deposited in GenBank, which makes it, to the best of
knowledge, specific for E. risticii. We were able
to amplify this gene from both positive snail pools and from the
SRC agent; as expected, we found more variation from the E. risticii sequence than we found in either the 16S rRNA gene
sequence or the groESL sequence. The deduced amino acid
sequences of the snail ehrlichiae and the SRC agent were identical
but differed from the E. risticii deduced amino
acid sequence at 12 positions, and 6 of the 12 changes were
conservative (Fig. 4). Variation in antigenic proteins is a widespread
and common characteristic of many bacteria, including E. risticii (8, 12, 13, 41). For example, there are
strains of E. risticii which have been described that
differ in their antigenic components but exhibit identity at the 16S
rRNA gene level (41). Such findings are in accord with the
findings of the present study, particularly in light of the sequences
obtained from the SRC agent and the known association of the SRC agent
with equine diarrheal disease (21).
Whether the pleurocerid snail itself or another agent harbored by the
snail is the actual vector of the snail ehrlichiae remains unclear.
Certainly, a trematode vector is a likely possibility, considering the
heavy trematode burden of snails and the historical precedent in the
region concerning Nanophyetus salmincola, the vector which
transmits the salmon poisoning rickettsia to canids (4, 9, 24,
38). In addition to snails, intermediate hosts of N. salmincola include salmonid fish and the Pacific giant salamander;
definitive hosts are represented by mammals and certain fish-eating birds (24, 38). Areas where fluke infestation and hence salmon poisoning are enzootic are determined by the geographical distribution of the host snail. A similar situation occurs
with the SF agent, which was isolated from a trematode of gray mullet
fish (15, 16, 43). Human disease caused by E. sennetsu has been associated with the consumption of
gray mullet (15, 16). The SF agent causes mild clinical
signs of fever in dogs (16), while E. sennetsu can establish infections in horses but evidently is not
pathogenic in this species (36).
Several trematodes that occur in western pleurocerids and could
potentially serve as vectors for ehrlichiae have been described. The trematode Acanthatrium oregonense, for example, produces
virgulate cercariae that penetrate the gills of caddisfly
larvae (order Trichoptera) (6, 7). The insects are eaten by
bats, the definitive hosts, in which the adult flukes develop in the
intestine. Many caddisfly larvae were observed in close association
with Juga spp. in the stream water described here, and bats
are known to be common in the area. In our laboratory at the University of California, Davis, we have observed many virgulate and rare furcocercous cercariae in secretions of Juga specimens from
the Siskiyou County site that have been maintained in aquarium culture (32). Cercariae characteristic of N. salmincola
were not observed in these secretions. Horses could conceivably be
exposed to E. risticii by consuming infected
cercariae (which are released from the snails after a period of
environmental warming and so are available to horses in drinking water
at certain times of the year), either free in the water or encysted on
vegetation or in a second intermediate host, such as an insect. The
seasonality of cercaria release from Juga spp. could explain
the widespread seasonal occurrence of PHF only in late spring, summer,
and early fall.
The known geographical distribution of Juga spp.
encompasses northern California, northern Nevada, Oregon, and
Washington (5), an area similar to the geographical
distribution of the pleurocerid host of N. salmincola. If
snails play a role in the life cycle of a trematode vector of
E. risticii in other areas of the United
States, different snails certainly must be involved. Pleurocerid genera
in the midwestern and eastern states that might be examined for
E. risticii include the genera Elimia,
Pleurocera, Lithasia, and Gyrotoma
(5). We have recently identified a putative E. risticii strain, resembling Ohio 081 in its 16S rRNA
gene sequence, in a pool of lymnaeid snails (Stagnicola
spp.) collected from an irrigation canal at Klamath Falls, Oreg., 70 miles north of the Siskiyou County site (32). The
distribution of the genus Stagnicola is particularly wide,
so members of this genus could provide a separate line of investigation
in other areas of the United States where PHF occurs. It is possible
that strains of ehrlichiae that cause PHF occur in association with a
variety of snail genera, a host range that may be reflected in the
genetic and antigenic variation commonly observed among
E. risticii isolates. Thus, the "agent" of
PHF may actually be an array of closely related ehrlichia strains that
differ in their vectors and snail hosts and, perhaps, in their
pathogenicities for horses as well.
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ACKNOWLEDGMENTS |
We thank Stacia Hoover, Eric Bowman, and Elfriede DeRock for
laboratory assistance; Norm Anderson for help with snail
identification; Jerry Fields for help with molluscan photography; Jerry
Theis, Steve Dumler, Ron Hedrick, Janet Foley, and Walter Boyce for
useful discussions; and Tom Sampson, Mike Ronne, Jon Goodell, Carie
Owings, Bob Cook, Wayne Merhoff, John Ballestin, Harry Sumner, Inderpal K. W. Singh, and Tim and Christi Saltonstall for valuable
contributions to our studies of SRC and PHF.
This work was supported by grants from the Center for Equine Health,
University of California, Davis, with funds provided by the Oak Tree
Racing Association, by the State of California satellite wagering fund,
and by contributions from private donors; and by discretionary funds
from the Department of Medicine and Epidemiology, School of Veterinary
Medicine, University of California, Davis.
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FOOTNOTES |
*
Corresponding author. Mailing address: School of
Veterinary Medicine, University of California, Davis, CA 95616. Phone
(530) 752-6513. Fax: (530) 752-0414. E-mail: jemadigan{at}ucdavis.edu.
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Applied and Environmental Microbiology, August 1998, p. 2888-2893, Vol. 64, No. 8
0099-2240/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
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Abstract
Introduction
