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Protozoa
Chapter 21 Protozoa Katherine Wasson
I.
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
II.
Flagellates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
520
A.
Giardia muris . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
520
1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
520
2. Life C y c l e and M o r p h o l o g y . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
520
3. Cell Biology
520
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4. Disease and D i a g n o s i s
B.
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521
6. R e s e a r c h Implications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
521
Spironucleus muris .........................................
523
1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
523
2. Life Cycle and M o r p h o l o g y . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. Cell Biology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
523
4. D i s e a s e and Diagnosis
524
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524 524
1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
524
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5. Treatment, Prevention, Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
III.
524
Tritrichomonas muris .......................................
4. Disease and D i a g n o s i s
E.
524
6. R e s e a r c h Implications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
2. Life C y c l e and M o r p h o l o g y . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. Cell Biology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
D.
521
5. Prevention, Treatment, Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
5. Prevention, Treatment, Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C.
518
524 525 525 525
6. R e s e a r c h Implications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
525
O t h e r Intestinal Flagellates
525
Trypanosoma musculi
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525
1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
525
2. Life Cycle and M o r p h o l o g y . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. Cell Biology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
526
4. Disease and D i a g n o s i s
526
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526
5. Treatment, Control, R e s e a r c h Implications . . . . . . . . . . . . . . . . . . . Amoebae ....................................................
527
A.
527
Entamoeba muris ..........................................
527
1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
527
2. Life Cycle and M o r p h o l o g y . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
527
3. Cell Biology
527
THE MOUSE IN BIOMEDICAL RESEARCH, 2ND EDITION
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Copyright 9 2007, 1980, Elsevier Inc. All rights reserved.
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KATHERINE 4. D i s e a s e and D i a g n o s i s
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5. Treatment, Prevention, Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6. R e s e a r c h Implications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV.
528
A.
General Introduction
528
B.
E i m e r i a spp . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
529
1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
529
C.
2. Life C y c l e . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
529
3. Cell Biology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
531
4. Disease and D i a g n o s i s
531
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D.
531
Sarcocystis m u r i s . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
532
1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
532
2. Life Cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
532
3. Cell Biology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
532
4. Disease and D i a g n o s i s
532
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533
Klossiella m u r i s . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
533
1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
533
2. Life C y c l e . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
533
3. Cell Biology
533
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533 534
6. R e s e a r c h Implications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
534
Toxoplasma gondii .. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
534
1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
534
2. Life Cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
535 535
3. Cell Biology
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536
5. Treatment, Prevention, Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
536
6. R e s e a r c h Implications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
536
Cryptosporidium muris
538
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1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
538
2. Life C y c l e . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
538
3. Cell Biology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
539
4. Disease and D i a g n o s i s
539
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5. Prevention, Treatment, Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
539
6. R e s e a r c h Implications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
540
Microsporidia A.
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5. Prevention, Treatment, Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
4. Disease and D i a g n o s i s
F.
533
6. R e s e a r c h Implications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
4. D i s e a s e and D i a g n o s i s
E.
531
6. R e s e a r c h Implications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
5. Prevention, Treatment, Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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E n c e p h a l i t o z o o n cuniculi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
540 540
2. Life C y c l e . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
540 541
3. Cell B i o l o g y
541
1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ...........................................
4. Disease and D i a g n o s i s
I.
528 528
5. Treatment, Prevention, Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
V.
527
Apicomplexans ............................................... .......................................
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541
5. Prevention, Treatment, Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
542
6. R e s e a r c h Implications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
543
Acknowledgments ..................................................
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References
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INTRODUCTION
The Kingdom Protozoa contains a diverse collection of single-celled eukaryotic organisms that in many ways bridge the gap between the plant and animal worlds. Protozoa may be free living or parasitic, may be capable of photosynthesis, or may be thriving in a microaerophilic environment. Others can be considered extremophiles, living in ice, hydrothermal vents, or sulfur-emitting fumaroles. A relatively small number
WASSON
of these organisms are parasitic for mammals. Although there is no universal agreement on the correct terminology for these organisms, the parasitic protozoa of mammals fall into one of four phyla: the amoebae (Sarcodina), the flagellates (Mastigophora), the apicomplexans (Sporozoa, or the coccidia), and the ciliates (Ciliophora). Of these, mice can be naturally infected with members from the first three phyla; there are no known reports of murine-specific ciliates. A fifth phylum, the Microspora (which includes Encephalitozoon cuniculi), is now considered a member of the Kingdom Fungi (see Table 21-1 for
21.
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PROTOZOA TABLE 21-1
COMPARISON OF MURINE PROTOZOANS a Phylum
Genus and species
Diagnostic stage(s)
Diagnostic test(s)
Size ( ~ t m )
Description
Amoeba
Entamoeba muris
Cyst
Fecal float, smear
9-19 diameter
Trophozoite
Fecal smear
12-30 diameter
Cryptosporidium spp.
Oocyst
Fecal float
5• 7
Eimeria falciformis
Oocyst
Fecal float
14-27 • 11-24
E. ferrisi
Oocyst
Fecal float
12-22 • 11-18
E. papillata
Oocyst
Fecal float
18-26 • 16-24
E. vermiformis
Oocyst
Fecal float
18-26 • 15-21
Klossiella muris
Sporocyst
Urinary centrifugation
16 x 13
Sarcocystis muris Toxoplasma gondii
Bradyzoite-filled cyst Cyst (intermediate host) Oocyst (Cat)
Histology Histology
4-6 • 14-16 Variable
Fecal float, smear
11-14 • 9-11
Tachyzoite
Histology
2-3 • 6-7
Bradyzoite
Histology
Giardia muris
Cyst Trophozoite
Fecal float, smear Fecal smear
15 • 17 7-13 • 5-10
SpironucIeus muris
Cyst
Fecal float
7• 4
Trophozoite
Fecal smear
10-15 x 3-4
Tritrichomonas muris
Trophozoite
Fecal smear
16-19 • 7-9
Trypanosoma musculi Encephalitozoon cuniculi
Trypomastigote Spore
Blood smear Histology
2-3 • 16-34 1.5 • 2.5
Round; 8 nuclei; lumen of cecum and ascending colon Pleomorphic, 1 nucleus; lumen of cecum and ascending colon Round to ellipsoid; gastrointestinal mucosa; intracellular but extracytoplasmic Round to ellipsoid; smooth; clear to light brown; crypt epithelium of cecum and colon Round; smooth; clear to light brown; villus epithelium of cecum and colon Round to ellipsoid; papillated; yellowish brown; villus epithelium of distal small intestine Ellipsoidal; pitted; yellowish brown; crypt epithelium of distal small intestine Round; glomerular endothelium and renal tubule epithelium Banana-shaped; myocytes Bradyzoites within cysts 5-8 • 1-2; prominent cyst wall; CNS, muscle, other tissues Round to subspherical; small intestinal epithelium Crescent shaped; within variety of nucleated cells Fusiform shaped; found within tissue cysts in variety of tissues Ellipsoid; 4 nuclei; lumen of small intestine Bilaterally symmetrical, pear-shaped; two nuclei; 4 pairs of flagella; closely associated with small intestinal villus brush border; "falling leaf' motility Ellipsoid; 2 nuclei; "Easter egg"; lumen of small intestinal Ellipsoid, tapered; found in mucus layer of small intestinal villus and crypts; "zig-zag" motility Pear-shaped; lumen of cecum and colon; "rolling" motility Elongated, vermiform shaped Rod-shaped; free or within parasitophorous vacuoles in brain, renal tubules
Apicomplexans
Flagellates
Microsporidia
aSee text for references.
a summary). Because of E. cuniculi's historical classification as a protozoan, and because of its importance as a pathogen in laboratory mice, this organism will be included in this discussion of murine protozoa. Not all of these organisms are pathogenic for mice. Indeed, the flagellate Tritrichomonas muris and the amoeba Entamoeba muris can be considered commensals of the murine large intestine. Others, for example, the flagellate Giardia muris and the apicomplexan Cryptosporidium muris, tend to cause disease in immunologically na'fve, neonatal mice. Susceptibility to these organisms tends to have a mouse
strain and gender predisposition, and it is expected that genetically engineered mice will vary in their susceptibility to disease as well. Lastly, several of the parasitic protozoa of mammals were first recognized and described in mice, and mice continue to be instrumental in understanding the pathology, immune response to, clearance, and treatment of these infections. Although parasitic infections are rare in well-managed facilities utilizing rederived and barrier-maintained mice, surveillance for and familiarization with these organisms in mouse colonies are still warranted.
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KATHERINE
II.
A.
FLAGELLATES
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G. lamblia, or unspeciated Giardia (Franjola et al. 1995; Sogayar and Yoshida 1995).
Giard& muris 2.
1. Introduction
Life Cycle and Morphology
Genus
Species
Primary host
Giardia
muris agilis ardeae lamblia (duodenalis, intestinalis) microti psittaci
Rodents Amphibians Herons Mammals including humans
G. muris, as with all Giardia species, has a simple and direct life cycle. Infection is acquired through the ingestion of environmentally resistant cysts. Experimental infections indicate that the minimal infectious dose of G. muris in outbred Swiss or athymic nude mice is 10 spores (range, 5 to 20), and the prepatent period prior to fecal shedding is 8 days (range, 5-14; Roberts-Thomas et al. 1976; Stachan and Kunstyr 1983). Cysts undergo excystation upon exposure to gastric pH and secretions from the upper gastrointestinal tract. Upon excystation, two trophozoites are released into the small intestine. Trophozoites are bilaterally symmetrical and pear-shaped, possess four pairs of flagella, and measure 7-13 x 5-10 ~tm. Two nuclei and the median body often give trophozoites a "ghost" face appearance under light microscopy. Trophozoites are found in the upper small intestine where they attach to intestinal epithelial cells by means of a ventral "adhesive" disk. Trophozoites replicate by binary fission to produce two identical daughter trophozoites. As trophozoites are sloughed into the intestinal lumen toward the large bowel, a certain percentage will encyst and be excreted as cysts into the environment. The length of time that cysts are shed in the feces is dependent on the dose of the original inoculum, host immune status, mouse strain, and possibly gender (Belosevic and Faubert 1983; Belosevic et al. 1984; Daniels and Belosevic 1995; Heyworth 1988). Cysts are ellipsoid in shape, possess four nuclei (two trophozoites), and measure 15 • 17 ~tm. To provide organisms for in vitro studies, G. muris trophozoites can be collected from the duodenum and jejunum of infected mice (or other infected hosts) and isolated in axenic culture media (Tillotson et al. 1991). Trophozoites can also be generated by harvesting cysts from fecal pellets and inducing them to excyst in the presence of trypsin and low pH (Schaefer et al. 1984).
Voles, muskrats Parrots
3.
muris andersoni baileyi canis felis galli hominis meleagridis molnari parvum serpentis saurophilium wrairi
Rodents Cattle Chickens, turkeys Dogs Cats Finches, chickens Primates including humans Turkeys Fish Cattle, sheep, goats Snakes, lizards Lizards Guinea pigs
Mice are natural hosts to Giardia muris and can be experimentally infected with G. lamblia. G. lamblia (also referred to as G. intestinalis, or G. duodenalis; Thompson et al. 2000) is a waterborne pathogen and an important cause of human diarrhea on a global basis (Adam 2001). Although the classification and nomenclature of Giardia is still evolving, it is generally agreed that six distinct species exist and that G. lamblia can be further subdivided into two major genotypes (Table 21-2; Adam 2001). The virulence and zoonotic potential between species and genotypes varies. Among the flagellates found in the intestines of mice, only G. muris is considered a primary pathogen of mice (Sebesteny 1969). Giardiasis in laboratory-reared mice should be nonexistent under proper rederivation, husbandry, and management standards. Colonies of G. muris-infected mice still exist, however. G. muris was diagnosed in 6.9% of "white" laboratory mice, and co-infection with other murine intestinal protists was noted (Jalili et al. 1995). Giardia infection is also common in wild rodent populations, which can serve as potential sources of infection for laboratory mice. Recent field studies of wild rodents (including Mus musculus and Rattus rattus) indicated that 14.3 to 100% of the various populations harbored G. muris,
TABLE 21-2
VALIDATED GIARDIA AND CRYPTOSPORIDIUM SPECIESb
Cryptosporidium
bAdapted from Adam (2001) and Xiao et al (2004).
Cell Biology
Giardia spp. possess two membrane-bound nuclei, a complex cytoskeleton, and a Golgi-like membrane system. However, they lack several features found in other eukaryotes, including nucleoli, mitochondria, enzymes involved in oxidative phosphorylation, and most enzymes involved in amino acid and nucleoside synthesis. As a result, Giardia are anaerobic and must forage for amino acids, purines, and pyrimidines from the intestinal milieu. The nuclei of Giardia are unique in that both are transcriptionally active and replicate at the same time. A second unique feature of Giardia is the ventral disk. This cytoskeletal structure allows the trophozoite to attach to intestinal epithelial cells of the duodenum and jejunum, preventing premature exit from the host. It is composed of
21.
several cytoskeletal proteins, including [3-tubulins (Parsons 1995). The median body, located on the midline, is another unique component of the trophozoite's cytoskeleton system. This structure is thought to be the assembly site for microtubule bundles to be incorporated into the ventral disk (Meng et al. 1996). The morphology of the median body can be used to distinguish between some of the Giardia species: G. lamblia has one or two transverse median bodies that are shaped like hammer claws; G. muris has one small, rounded median body (Adam 2001). Giardia lack mitochondria and the ability to generate energy through aerobic metabolism. However, identification of mitochondria-like heat-shock protein genes and mitochondrial remnant organelles ("mitosomes") suggests that Giardia may have lost this organelle during the course of adapting to a parasitic lifestyle (Roger et al. 1998; Tovar et al. 2003). Lastly, Giardia trophozoites are capable of antigenic variation by expressing a family of cysteine-rich, immunodominant proteins across their surfaces known as variable surface proteins (Aggarwal et al. 1988). The ability to undergo antigenic variation is likely a means of evading the host immune response and may give the organisms a survival advantage in differing intestinal microenvironments (Adam 2001). 4.
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PROTOZOA
Disease and Diagnosis
G. muris infections generally do not cause clinical signs in immunocompetent mice. Immunocompromised mice may exhibit weight loss and failure to thrive. These clinical signs are also observed in mice infected as weanlings, presumably due to a less mature immune system at this age (Buret et al. 1990). Unlike giardial infection in other mammals, diarrhea is not a clinical feature of disease in mice (Eckmann and Gillin 2001). Giardia clearance, as well as acquired immunity to the parasite, is primarily dependent on IgA production (Langford et al. 2002). Cellular immune responses and innate immunity factors such as defensins and host microflora also contribute to disease resistance and clearance (Singer and Nash 1999, 2000). G. muris can be diagnosed by identifying the characteristic trophozoites or cysts in feces by light microscopy. A "falling leaf' motility may be observed in fresh, unstained wet-mount samples taken from the duodenum or jejunum. Wright or Giemsa stains can be used to enhance the "ghost" face appearance of trophozoites. Cyst stages can be identified on fecal flotation or smears, and differentiated from other cysts or oocysts by their size (Table 21-1). Four nuclei may be seen on trichrome- or iodine-stained preparations of cyst forms. Owing to irregular shedding of cysts, several fecal examinations may need to be performed prior to ruling out an antemortem diagnosis of giardiasis. Histologically, trophozoites are found attached to the brush border of epithelial cells of the duodenum and jejunum. Trophozoites are lightly eosinophilic, crescentshaped on longitudinal view, and noninvasive. Careful examination under oil immersion may reveal the presence of
the ventral disk (Fig. 21-1A). There may be mild reduction in the villus:crypt ratio, and mild to moderate lymphocyte infiltration in the underlying mucosa and lamina propria (MacDonald and Ferguson 1978). Organisms need to be differentiated from Spironucleus muris, which tend to localize in the distal small intestine and are often found packed in intestinal crypts (Brett and Cox 1982, Owen et al. 1979). Enzyme-linked immunosorbent assays, flourescence antibody assays, and molecular-based methods are also available for diagnosis of giardiasis (Garcia et al. 1992; Nash et al. 1987; Sedinova et al. 2003). These tests are used for the detection of human giardial infection, and several are available in kit format. Use of these or comparable diagnostic kits for screening rodent colonies has not been reported. 5.
Prevention, Treatment, Control
G. muris is readily transmitted between mice and other rodent species (Belosevic et al. 1986a; Kunstyr et al. 1992; Saxe 1954). Therefore, rederivation is the treatment of choice for eliminating Giardia spp. from rodent colonies. Metronidazole and albendazole are the drugs of choice for treating humans infected with G. lamblia (Gardner and Hill 2001). The efficacy of these compounds may not be complete in mice (Bemrick 1963; Oxberry et al. 1994). Treatment with oral metronidazole resulted in only a 58.3% cure rate in mice naturally infected with G. muris, based on histologic examination of the small intestine for the presence of trophozoites (Cruz et al. 1997). Cyst forms shed into the environment are resistant to chlorine and ozone exposure, but can be inactivated by autoclaving of bedding and caging material (Lane and Lloyd 2002).
6.
Research Implications
Subtle alterations in intestinal epithelial cell kinetics, brush border enzyme composition, and cytokine production have been documented in immunocompetent mice with G. muris infection. Although villus height remained the same, a higher turnover rate of intestinal epithelial cells was observed in mice naturally co-infected with G. muris and Spironucleus muris when compared to control mice (MacDonald and Ferguson 1978). Significant decreases in the brush border enzymes lactase, sucrase, trehalase, and maltase were documented in C57BL/6 mice by day 10 of infection with G. muris (Daniels and Belosevic 1992). Reduced production of the epithelial cytokine IL-6 was reported in CD-1 mice (Scott et al. 2000). In addition, an increased epithelial lymphocyte infiltration of the small intestine has been observed with resolving Giardia infection (Brett and Cox 1982). All of these factors make endemically infected mice unsuitable for research involving intestinal physiology. A decreased T cell response has also been reported in mice experimentally infected with G. muris (Brett 1983). This was documented by a decreased ability of infected mice to mount
522
KATHERINE
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Fig. 21-1 Intestinal flagellates of mice. (A) Two Giardia spp. trophozoites in the upper small intestine of a mouse. Note the pear-shaped trophozoite with two nuclei and posterior flagella, and its close association to the intestinal brush border; the second trophozoite is oriented longitudinally, demonstrating the cup shape of the ventral adhesive disk. (B) Spironucleus muris trophozoites in the distal small intestine of a mouse; trophozoites are small and tapered, and associated with the mucus layer. (C) Tritrichomonas muris and (D) T. minuta in the ceca of mice; note the foamy cytoplasm and larger size of T. muris compared with T. minuta. H&E; A through D, bar = 50 ~tm.
2 1.
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PROTOZOA
an immune response to sheep red blood cells. T cell responses returned to normal with resolution of infection (Brett 1983). Not surprisingly, exogenous administration of the immunosuppressive drugs cortisone and cyclosporin A resulted in increased cyst shedding and prolonged time to clearance of infection (Nair et al. 1981; Belosevic et al. 1986b). In the case of cortisone, recrudescence of subclinical infection could also be induced (Nair et al. 1981). These results suggest that administration of immunosuppressive agents to mice on experiment may exacerbate occult G. muris infection. A murine model of giardiasis was first described in 1976, and although it does not replicate all the features of human giardiasis, mice have been valuable in understanding the pathophysiology of this disease (Roberts-Thomas et al. 1976). Choice of mouse to use for infection studies should take into account strain, major histocompatibility complex haplotype, and perhaps gender. Early work demonstrated that C57BL/6, C57BL/10, and DBA/2 strains have a longer prepatent period, lower fecal cyst output, and faster resolution of experimental G. muris infection than A/J, BALB/c, or C3H/He strains of mice (Belosevic et al. 1984; Daniels and Belosevic 1992). The differences in the time course of infection were hypothesized to be due to differences in production of IgG2a and gamma interferon (7-INF) inherent in the different mouse strains. A more virulent giardial infection could be induced in C57BL/10 mice by the administration of anti-y-INF antibodies (Venkatesan et al. 1996). No gender differences were observed in the number of cysts shed in feces in BALB/c mice during acute infection with G. muris (Heyworth 1988); however, female C57BL/6 mice cleared G. muris infection by 18 days post-inoculation, while male mice of the same strain continued to shed cysts in the feces beyond 60 days (Daniels and Belosevic 1995). Murine haplotype also plays a role in clearing infection. BALB mice of the H-2b haplotype shed G. muris cysts for a longer period of time than BALB mice of the H-2d or H-2k haplotypes (Venkatesan et al. 1993). The commercial source of mice should also be considered. Isogenic mice from two different vendors were found to have different susceptibilities to experimental G. lamblia infection (Singer and Nash 1999). The inherent resistance in mice from one vendor could be overcome when the intestinal flora was altered with the antibiotic neomycin. Resistance in these mice could also be overcome when housed with susceptible mice from the second vendor (Singer and Nash 1999). These results suggest that differences in the endogenous microflora and fauna can affect both experimental and natural Giardia infection.
B. 1.
Spironucleus muris
Introduction
Originally called Hexamita muris, Spironucleus muris has been renamed to differentiate the exclusively parasitic ("Spironucleus")
from the usually free-living ("Hexamita") members of this genus (Brugerolle et al. 1980). Recent surveys of laboratory and wild mouse populations suggest that the prevalence of S. muris ranges from 4.1 to 38.6% (Franjola et al. 1995; Jalili et al. 1995). Spironucleus muris is probably represented by subspecies with differing host preferences and infectivity. In several separate transfaunation studies, isolates of S. muris from mice and golden Syrian hamsters were infectious for the reciprocal host; isolates obtained from a rat and European hamster were not infectious for mice; and in a third study, isolates from a mouse and rat were infectious for hamsters and rats (Saxe 1954; Schagemann et al. 1990; Sunstyr et al. 1993). Regardless of host specificity, these organisms are likely commensals in their respective rodent hosts (Baker et al. 1998; Sebesteny 1969). Several clinical reports from the 1970s describe "outbreaks" of spironucleosis in laboratory mice, with mortality rates ranging from 20 to 50%. Clinical signs included chronic wasting in athymic nude and thymectomized mice (Boorman et al. 1973); or depression, distended abdomens, "sticky stools," and nonbloody, catarrhal enteritis in weanling age mice (Flatt et al. 1978; Wagner et al. 1974). Large numbers of Spironucleus muris trophozoites, and occasionally G. muris cysts, were observed at necropsy. Thirty years later, descriptions of these "outbreaks" are reminiscent of disease seen with enteric mouse hepatitis virus (MHV), transmissible murine colonic hyperplasia (Citrobacter rodentium), Tyzzer's disease (Clostridium piliforme), and/or salmonellosis rather than primary infection with S. muris. Today, large numbers of S. muris organisms are frequently observed secondary to underlying disease, often MHV infection (Percy and Barthold 2001). A diagnosis of spironucleosis should prompt the clinician to search mouse colonies for an underlying infectious agent or disease condition. 2.
Life Cycle and Morphology
S. muris has a simple and direct life cycle. Infection is initiated with the ingestion of cysts. A single trophozoite is released from S. muris upon excystation in the upper gastrointestinal tract (Schagemann et al. 1990). Trophozoites replicate by longitudinal binary fission. Trophozoites have a slender, tapered body measuring 10-15 x 3-4 ~tm. They possess four pairs of flagella and two nuclei, but lack the specialized cytoskeletal features seen in Giardia spp. (Brugerolle et al. 1980). Trophozoites encyst as they move down the intestinal tract. Cysts measure 7 • 4 ~tm and resemble "Easter eggs" due to the presence of flagella under the cyst membrane, giving the cysts a banded appearance (Kunstyr !977; Kunstyr et al. 1977). The minimal infective dose is one cyst, and the prepatent period has been reported between 2 and 8 days for mice (Kunstyr 1977; Stachan and Kunstyr 1983). In vitro culture and manipulation of S. muris have not been reported; organisms are passaged in flagellate-free mice for experimental manipulation (Schagemann et al. 1990).
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3. Cell Biology Relatively little is known regarding the cellular and molecular biology of S. muris. Like Giardia, they are early eukaryotic organisms, possessing a membrane-bound nucleus, but lacking mitochondria and, presumably, the enzymes involved in aerobic metabolism (Brugerolle et al. 1980). S. muris lacks the ventral disk and median body observed in Giardia (Januschka et al. 1988). Trophozoites feed on luminal flora, as evidenced by the presence of bacteria in digestive vacuoles seen on electron microscopy (Brugerolle et al. 1980). An inverse ratio of Giardia and Spironucleus trophozoites has been observed in naturally infected mice, suggesting that these organisms compete for similar resources in the host (Sebesteny 1969). 4.
Disease and Diagnosis
Early descriptions of disease due to spironucleosis must be interpreted with caution, as S. muris is likely a "facultative pathogen" in mice infected with other disease agents (Kunstyr et al. 1977). In contemporary mouse colonies, heavy S. muris burdens are associated with enteric mouse hepatitis virus infection (Percy and Barthold 2001). Spironucleosis has also been observed in otherwise pathogen-free, genetically engineered mice with underlying immune alterations. Experimental S. muris infection of athymic nude mice resulted in abdominal distention, failure to thrive, and death after 2 to 3 months of infection (Kunstyr et al. 1977). A more recent experimental report noted a lack of clinical disease and intestinal pathology in a variety of inbred strains of mice orally inoculated with S. muris cysts (Baker et al. 1998). Diagnosis of S. muris depends on observing trophozoites or cysts in fecal smears or histopathology. Trophozoites have a fast, zigzag movement on fresh wet-mount samples taken from the distal small intestine and colon (Sebesteny 1969). The nuclei are best visualized with Giemsa or hematoxylin stain (Sebesteny 1969). Cysts are found in the large intestine and feces, and can be differentiated from Giardia cysts by their smaller size and "Easter egg" appearance (Kunstyr et al. 1977). On histologic sections, organisms are located in the distal small intestine and are usually found in the mucus layer or packed within intestinal crypts (Wagner et al. 1974; Brugerolle et al. 1980). Invasion of the intestinal mucosa and lamina propria by organisms has also been reported (Flatt et al. 1978). S. muris trophozoites appear as smudgy, lightly eosinophilic structures with hematoxylin-eosin stain (Fig. 21-1B). Under 100X oil immersion, they appear as plump banana-shaped organisms, with a single mid-body nucleus visible. Diagnostic immunologic or molecular techniques have not been reported for S. muris. 5.
WASSON
dimetronidazole, metronidazole, or tinidazole was ineffective in eliminating cyst shedding in mice (Kunstyr et al. 1977; Sebesteny 1969). Albendazole, a microtubule inhibitor with some efficacy against Giardia infection, had minimal effect on the ultrastructure of S. muris (Oxberry et al. 1994). Cysts are susceptible to several common disinfectants, fixatives (70% ethanol, household bleach, aldehyde-based compounds), and temperatures above 45~ suggesting that standard laboratory animal husbandry and management procedures will control cysts in the environment (Kunstyr and Ammerpohl 1978). 6. ResearchImplications S. muris infection complicates research with experimental intestinal flagellate infection (Owen et al. 1979). In that report, mice colonized with S. muris were less susceptible to experimental G. muris infection. Alterations in intestinal epithelial cell kinetics were observed in mice co-infected with S. muris and G. muris (MacDonald and Ferguson 1978). A decreased villus to crypt ratio, and decreased T cell-dependent immune response to sheep red blood cells, have been observed in S. muris-infected mice (Brett 1983; Brett and Cox 1982).
C.
Tritrichomonas muris
1. Introduction Tritrichomonas muris is a nonpathogenic flagellate of mice, rats, hamsters, and other rodents. Early literature refers to this organism as Trichomonas muris, Trichomonas cricetus, or Tritrichomonas cricetus. However, these were determined to be synonyms for morphologically identical organisms found in a variety of laboratory and wild rodents (Daniel et al. 1971; Honigberg 1963). An exhaustive transfaunation experiment demonstrated the ease with which several trichomonads (including T. muris and Pentatrichomonas hominis) could be established in and transmitted between mice, rats, and hamsters (Saxe 1954). T. muris is related to the more pathogenic Trichomonas vaginalis and Tritrichomonas foetus of humans and cattle, respectively. These latter organisms are sexually transmitted protists that live in the genitourinary tracts of their hosts, causing reproductive disorders (BonDurant 1997; Petrin et al. 1998). T. muris resides in the cecum and colon of rodents, where it is considered a component of the normal fauna (Sebesteny 1969). Recent surveys of laboratory and wild colonies of mice demonstrate a 29.6 to 47.4% prevalence of the organism in the large intestines (Franjola et al. 1995; Stachan and Kunstyr 1983). Although little is known of T. muris, much of its life cycle and cell biology can be inferred from work done on more pathogenic family members.
Prevention, Treatment, Control
S. muris is readily transmitted between rodents (Saxe 1954).
Rederivation and fostering on flagellate-free dams will eliminate S. muris from mouse colonies. Chemotherapy using
2. LifeCycle and Morphology T. muris has a simple and direct life cycle, and exists as motile trophozoites within the host. Pseudocyst forms of
2 1.
T. vaginalis and other trichomonads have been observed and are thought to represent degenerating trophozoites responding to unfavorable environmental conditions (Honigberg 1963; Petrin et al. 1998). The minimal infectious dose of T. muris "pseudocysts" for mice is 5, and the prepatent period is 10 days (Stachan and Kunstyr 1983). Trophozoites are pear- or teardropshaped, measuring 16-19 • 7-9 ~tm (Selukaite 1977; Fig. 21-1C). Trophozoites replicate by binary fission. Trophozoites reside in the cecum and colon but have been reported in the stomach and small intestines, likely as a result of recent ingestion (Koyama et al. 1987; Selukaite 1977). Newborn hamsters, and probably other rodent pups as well, are colonized by T. muris within a week after birth (Mattern and Daniel 1908). Cell-free cultivation of T. muris has not been reported; trophozoites have been harvested from mono-infected rodents for experimental manipulation (Saxe 1954).
3.
Cell Biology
All trichomonads possess a single membrane-bound nucleus, three anterior flagella, and a fourth posterior flagellum that forms the "backbone" of the undulating membrane. The undulating membrane runs partway down the length of T. muris and can be visualized in less motile trophozoites under light microscopy (Osada 1962). The axostyle is the cytoskeletal structure that appears to give the organism a "backbone." It originates at the nucleus and tapers to a tail-like appendage at the distal end of the parasite (Daniel et al. 1971). In T. vaginalis, the axostyle is thought to be an attachment organ on vaginal epithelial cells (Petrin et al. 1998). On electron microscopy, a comb-like structure called the costa can be seen anchoring the base of the undulating membrane (Daniel et al. 1971). The parabasal body is a collection of flattened cisterns similar in structure to the Golgi complex in higher eukaryotes. Trichomonads lack mitosomes but possess double-membrane bound granules that metabolize carbohydrates and produce ATP and hydrogen, and are referred to as hydrogenosomes (Petrin et al. 1998). 4.
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Disease and Diagnosis
No disease has been attributed to T. muris in rodents. Impressive numbers of trophozoites can be seen in the cecal lumen of some mice in the absence of disease (Fig. 21-1C). Diagnosis can be made by examining fresh wet mounts from the cecum and colon by light microscopy. Organisms can be identified by their "rolling" or "quivering" movement (Petrin et al. 1998). Staining wet mounts with Giemsa or iodine enhances the appearance of the nucleus and undulating membrane.
indicator of breach in barrier maintenance of otherwise pathogen-free mice. Chemotherapeutic elimination of T. muris has not been reported. The lack of T. muris cyst forms implies that standard husbandry and management practices will eliminate organisms from the environment.
6.
Interference with research results due to the presence of T. muris in mice has not been reported. Due to the health significance of T. vaginalis in the human population, there has been interest in using the laboratory mouse as a model for this sexually transmitted disease. Patent, long-term infection requires pretreatment of female mice with estrogen and intravaginal doses of Lactobacillus spp. prior to the introduction of T. vaginalis organisms (McGrory and Garber 1992). Using this model, investigators have demonstrated that IgA antibodies protect against severe infection with T. vaginalis and that T-lymphocytes are important in parasite clearance in infected women (Paintlia et al. 2002).
D.
Rederived and barrier-maintained mice are free of T. muris. The ease with which T. muris can be transmitted between mice and other rodents suggests that trichomonads can be used as an
Other Intestinal Flagellates
In addition to the above-mentioned protists, several other nonpathogenic flagellates have been identified in the intestines of the laboratory mouse. Most of these can be identified according to their morphology and size. Trichomonas minuta (4-9 x 2-5 l.tm) and Trichomonas wenyoni (4-16 x 2.5-6 l.tm) are smaller in size and have a less vacuolated or "foamy" cytoplasm compared to T. muris (Levine, 1973b). T. minuta has a more prominent and eosinophilic axostyle than T. muris (Fig. 21-1D). Chilomastix bettencourti is a cyst-forming flagellate found in the cecum of mice, rats, and hamsters. Trophozoites are asymmetrically piriform and measure 8-20 x 7-8 l.tm; cysts are lemon-shaped and measure 6.5-9 x 5.5-7 l.tm (Nie 1948). They possess three anterior flagella and a short, posteriorly directed flagellum. A distinctive, pouch-like cytostome can be seen in both forms, and is thought to be a feeding organelle (Levine 1973a). Octomitus pulcher is a bilaterally symmetrical flagellate that morphologically resembles Giardia spp. They measure 6-10 x 3-7 ~tm and possess six anterior and two posterior flagella, as well as two anteriorly located nuclei. O. pulcher is found in the cecum of mice, rats, hamsters, and other wild rodents (Gabel 1954; Levine 1973a).
E. 1.
5. Treatment, Prevention, Control
Research Implications
Trypanosoma musculi
Introduction
Trypanosoma musculi is a nonpathogenic hemoflagellate parasite of wild mice living in the Mediterranean basin, West Africa, and Central America. T. musculi is host-specific for Mus musculus (Krampitz 1969). First described in 1909, this
526
KATHERINE
WASSON
organism is also referred to as T. duttoni in the older literature (Kendall 1906). A relatively recent survey identified T. musculi in 3.8% of wild house mice in the Arabian peninsula (Molan and Hussein 1988). There are no published reports of natural infection with this parasite in laboratory-raised mice.
recta of the kidneys, and suggested that mice can be persistently infected for life (Viens et al. 1972). Subsequent work has shown that capillaries of the vasa recta provide nutrients and an "immunologically" privileged site for T. musculi trypomastigotes (Monroy and Dusanic 1997).
2.
3.
Life Cycle and Morphology
The life cycle and morphology of T. musculi is similar to that of other African trypanosomes. T. musculi is transmitted by fleas, including the Oriental rat flea, the Northern rat flea, and the mouse flea (Xenopsylla cheopis, Nosopsyllus spp., and Leptosylla segnis, respectively). Mice become infected by ingesting infected fleas or flea feces containing trypanosomes. Mice become parasitemic, during which time trypomastigotes replicate by multiple fission, and can be identified in peripheral blood smears. Trypomastigotes have an elongated, vermiform shape and measure 2 to 3 ktm in width and 10 to 34 ~m in length (Taliaferro and Pavlinova 1936; Fig. 21-2). Parasitemia lasts 2 to 3 weeks, after which time organisms are difficult to identify in blood smears. The life cycle is perpetuated when fleas consume a blood meal from parasitemic mice. The peak parasite burden varies considerably in immunocompetent strains of mice (Derothe et al. 1999). Mice develop immunity and are resistant to reinfection with T. musculi. However, Viens et al. demonstrated that a small percentage of parasites persisted (and continued replicating) in the vasa
Trypanosomes possess a membrane-bound nucleus and a single flagellum. Trypanosomes also possess a single mitochondrion referred to as a kinetoplast. In addition to performing metabolic functions as in other eukaryotic organisms, this organelle has a unique mitochondrial genome organization and function (McFadden 2003). Indeed, several unique RNA processing mechanisms were first described in the kinetoplast of trypanosomes (Gott and Nilsen 2003). These include transsplicing and RNA editing. The DNA in the kinetoplast is organized in 20 to 50 "maxicircle" DNA segments and 5000 to 10,000 small circular DNA segments. The large content of DNA in this organelle contributes to its intense staining. Instead of the conventional linear method of transferring genomic information from DNA to RNA (transcription), and then to protein (translation), various transcripts of RNA from different areas of the kinetoplast genome can be ligated together prior to being translated (trans-splicing). The ability to perform trans-splicing is the mechanism behind the phenomenon of antigenic variation in African trypanosomes, a major means by which parasites evade the host immune system. In addition to modifying RNA by ligating pieces together, sections of transcribed RNA can be altered by the insertion or deletion of uridine residues in order to create messenger RNA that will code for functional proteins (RNA editing). Once thought to be unique to the kinetoplastid parasites, trans-splicing has since been identified in parasitic and free-living nematodes, and RNA-editing has been described in plant mitochondria and chloroplasts. 4.
Fig. 21-2 Trypanosoma sp. in a peripheral blood smear from a cow. Note the kinetoplast, undulating membrane, and single flagellum. Similar features would be seen in T. musculi from mice. H&E; bar - 50 gm (Image courtesy of H. Gelberg).
Cell Biology
Disease and Diagnosis
T. musculi is relatively nonpathogenic for immunocompetent mice. Mice develop mild anemia, splenomegaly, hepatomegaly, and lymph node hyperplasia that resolve after one month (Hirokawa et al. 1981). Mice experimentally infected between the fourth and fifteenth day of gestation, however, develop fatal parasitemia, with large numbers of replicating trypomastigotes present in the maternal vessels of the placenta (Krampitz 1969). Athymic nude mice develop persistent infection, while splenectomized mice develop fatal parasitemia, when experimentally inoculated with T. musculi trypomastigotes (Rank et al. 1977; Taliaferro and Pavlinova 1936). The trypomastigote forms of T. musculi are morphologically similar to those of other mammalian trypanosomes. When stained with Giemsa, they have a large, red, centrally placed nucleus and a smaller, posteriorly located kinetoplast. The cytoplasm
2 1.
527
PROTOZOA
stains blue. An undulating membrane can be seen by light microscopy running the length of the trypomastigote. The membrane continues past the main body of the parasite as a free, anteriorly located flagellum. Infection is best diagnosed on peripheral blood smears. Trypomastigotes may also be observed in histologic sections of spleen, liver, and kidneys (Hirokawa et al. 1981). 5. Treatment, Control, Research Implications
Because of the self-limiting nature of T. musculi infection, treatment of infected mice has not been reported. Experimental infection in laboratory mice has been proposed as an in vivo model for screening chemotherapeutics against Chagas' disease in humans (Jennings and Gray 1982). However, laboratory mice can be experimentally infected with the etiologic agents of both human American and African trypanosomiasis (T. cruzi and T. brucei spp., respectively). Infection in mice with these organisms mimics many of the pathologic and immunologic features of chronic human infection (Kennedy 1999; Marinho et al. 2004). Consequently, T. musculi infection in mice is rarely reported as a model for human trypanosomiasis. In areas where T. musculi is endemic in wild mouse populations, prevention of infection in the laboratory setting involves appropriate vermin and insect control. There are no published reports of natural T. musculi infection confounding experimental data obtained from mice.
III.
A.
AMOEBAE
Entamoeba muris
1. Introduction The genus Entamoeba is composed of a diverse group of parasitic and free-living, single-celled organisms with an amoeboid mode of locomotion (Silberman et al. 1999). Most Entamoeba are nonpathogenic. Entamoeba muris is the only amoeba identified in laboratory mice and is considered a commensal inhabitant of the cecum. The older literature refers to this organism as Amoeba muris, Entamoeba muris decumani, Councilmania muris, Councilmania decumani, Endamoeba ratti, or Entamoeba coli var ratti. Based on
morphology, these terms are now considered synonyms for E. muris (Neal 1950). Surveys of laboratory and wild mouse populations demonstrate a prevalence of E. muris between 5 and 55% (Franjola et al. 1995; Jalili et al. 1995; Livingston 2004; Pruss 1960). Organisms morphologically identical to E. muris are also identified in laboratory and wild rats, and hamsters (Pruss 1960). E. muris is related to the more pathogenic E. histolytica and E. invadens of humans and reptiles, respectively.
2. LifeCycle and Morphology Entamoeba spp. have a simple and direct life cycle. Trophozoites are found in the cecum and anterior colon, where they feed on bacteria, protozoa, and other luminal material (Lin 1971). They are pleomorphic on wet mounts, round or ovoid in histologic sections, and possess a single nucleus (Neal 1950). The pseudopod, an ectoplasmic extension distinct from the endoplasm (cytoplasm), and a trailing uropod may be observed in wet-mount preparations and account for the amoeboid motion of these organisms. They lack cilia, flagella, or other organized cytoskeletal structures. The cytoplasm contains vacuoles of varying size. Trophozoites replicate by binary fission. Encystation occurs in the cecum, with mature cysts possessing eight nuclei and measuring 9 to 20 ktm in diameter. Immature or "precyst" stages may possess four nuclei. Amoebae that pass from the cecum without encysting do not survive outside the host (Lin 1971). Cysts are excreted in the feces and are available for ingestion and infection in the next host. The signals involved in excystation of ingested cysts in the cecum are not known. The environmental stability of E. muris is unknown. Cell-free cultivation of E. muris has not been reported.
3. CellBiology The genus Entamoeba has classically been divided into groups based on the number of nuclei in the mature cyst forms (Neal 1966). In general, nonpathogenic Entamoeba have one nucleus (E. chattoni from nonhuman primates; E. polecki from pigs and humans) or eight (E. coli from humans; E. muris from mice) nuclei per mature cyst. The pathogenic E. histolytica and nonpathogenic E. dispar of humans possess four nuclei per mature cyst. In addition, E. invadens of reptiles possess four nuclei in the mature cyst. This classification has held up to phylogenetic analysis of ribosomal DNA and protein sequences of Entamoeba (Silberman et al. 1999). This morphologic detail is important when diagnosing amoeba infection in other species. The presence of octonucleate cysts in the feces of a reptile suggests that these are E. muris that were transiently acquired from feeder mice, while quadranucleate cysts suggest parasitism with E. invadens. Like the flagellates, Entamoeba lack mitochondria, peroxisomes, and a Golgi complex. A mitochondrial-like organelle, variably termed crypton, cryptome, or mitosome, has been identified in E. histolytica (Mai et al. 1999; McFadden 2003; Tovar et al. 1999). Its function has yet to be determined. Entamoeba spp. generate energy through glycolysis. Elongated, rhomboidshaped cytoplasmic inclusions called chromatoid bodies can be seen by light microscopy in cyst stages of Entamoeba spp. These bodies are aggregates of ribosomes, and their function is unknown (Kusamrarn et al. 1975; Neal 1966). 4. Disease and Diagnosis
Disease due to the presence of E. muris in the cecum and colon of mice has not been reported. Although molecular and
528
KATHERINE
cellular analysis of E. muris has not been reported, it is likely that this species lacks the virulence factors that are well characterized in E. histolytica. These factors include lectin-binding molecules, channel-forming amoebapores, and cysteine proteases responsible for tissue destruction in cases of invasive human amoebiasis (Espinosa-Cantellano and Martinez-Palomo 2000). E. muris can be diagnosed by examining fresh wet mounts from the cecum and colon by light microscopy. If slides are kept at 37~ amoeboid movement may be observed. When cooled, organisms tend to round up and are more difficult to identify. Visualization of octonucleate cysts is enhanced by the addition of iodine to wet mounts. In histologic sections, trophozoites are round to oval in shape and vary from 8 to 30 ~tm in diameter (Fig. 21-3). They live in colonies of varying number within the mucus layer of the cecum, and rarely, anterior colon. Trophozoites stain eosinophilic with a granular and often highly vacuolated cytoplasm. Close examination of the vacuoles may reveal ingested bacteria or other protists. Cyst forms are smaller and possess a thin cell wall. Refractile chromatoidal bodies may be present in the cytoplasm. Due to plane of sectioning, mature cysts often appear to contain less than eight nuclei. 5.
Treatment, Prevention, Control
As with the intestinal flagellates, rederived and barriermaintained mice are free of E. muris. Chemotherapeutic elimination of E. muris from infected mice has not been reported,
Fig. 21-3 Entamoeba muris cysts from the cecum of a mouse. Note variably sized intracytoplasmic vacuoles. H&E; bar = 50 l.tm.
WASSON
although luminal amoebicides (iodoquinol, diloxanide furoate, paromomycin) or imidiazoles (metronidazole) used as therapy for human amoebiasis may be effective (Upcroft and Upcroft 2001). The environmental stability of E. muris cysts has not been reported. 6.
Research Implications
Interference with experimental design or reproducibility has not been reported from mice harboring E. muris. There has been tremendous interest in creating a murine model of human amoebic dysentery and invasive amoebiasis. Unfortunately, E. histolytica is not very pathogenic for mice (Ghadirian et al. 1987; Gold and Kagan 1978; Stern et al. 1984). Early reports of mouse mortality due to intracecal inoculation with E. histolytica were complicated by the fact that the organisms were co-cultured with a cocktail of potentially pathogenic bacteria (including E. coli and clostridial species) in order to generate organisms for mouse experiments (Owen 1985, 1990). Since then, methods for cultivating E. histolytica in monoand axenic media have been developed, and more mousevirulent strains isolated (Clark and Diamond 2002). Still, patent intestinal E. histolytica infection in mice depends on intracecal inoculation of xenically passaged organisms (Thompson et al. 2000). Infection is also dependent on mouse strain: 60% of cecally inoculated C3H mice developed typhlitis and remained persistently infected 18 months post-inoculation, compared with no infection or histologic lesions in BALB/c, C57BL/6, or INF-7-, ILl2-, or iNOS-knockout mice (Cieslak et al. 1992). An interesting variation on the mouse model of amoebiasis is the SCID (severe combined inmmunodeficient) mousehuman intestinal xenograft model (Seydel et al. 1997). In this model, human fetal intestinal sections are transplanted under the skin of C.B-17-Prkdc scid (SCID) mice and allowed to develop for several weeks prior to direct inoculation into the intestinal lumen with E. histolytica trophozoites. With this model, researchers have characterized the proinflammatory cytokines released by human intestinal epithelial cells in response to E. histolytica infection and have examined the virulence of genetically manipulated E. histolytica trophozoites (Zhang et al. 2003, 2004). Models of invasive amoebiasis, the most common form in humans being hepatic abscess, can be recreated by intrahepatic inoculation of trophozoites into SCID mice (Cieslak et al. 1992).
IV.
APICOMPLEXANS
A.
General Introduction
The phylum Apicomplexa is a large and diverse group of spore-forming, sexually reproducing protozoans that share
2 1.
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PROTOZOA
an anteriorly located, apical complex important for host-cell invasion. This phylum includes the coccidia (Cryptosporidium, Eimeria, Isospora, Klossiella, Sarcocystis, Toxoplasma gondii), piroplasms (Babesia), and hemoapicomplexans (Plasmodia). Apicomplexans of importance in the laboratory mouse include Eimeria spp., Sarcocystis muds, Klossiella muds, Cryptosporidium spp., and Toxoplasma gondii. Although apicomplexans infect a variety of cell types and cause distinctly different diseases in their respective hosts, they share a common basic life cycle (Levine and Ivens 1990). This life cycle starts with the oocyst (or spore form), which must undergo sporulation (meiosis) before becoming infective for the host. Once sporulated, the oocyst is composed of sporoblasts, each of which contains a specific number of invasive sporozoites, depending on the species. After the sporulated oocyst is ingested by a susceptible host, the sporozoites are released and invade the appropriate host cells. These forms replicate by multiple fission resulting in daughter merozoites (also referred to as schizozoites or trophozoites). Merozoites are released by rupture of the host cell and invade neighboring cells. This cycle of asexual replication, rupture, and re-invasion (referred to as merogony or schizogony) is repeated for one or more generations, depending on the parasite species. After asexual replication, the parasites cease replicating and differentiate into microgametes or macrogametes within the host cells. Macrogametes are fertilized by motile microgametes and develop into oocysts, which mature and rupture from the host cell to start the parasite life cycle again. Variations in this basic apicomplexan life cycle include length of time and location where sporulation occurs, number and arrangement of sporoblasts and sporozoites within the sporulated oocyst, tissue and cell preference of the sporozoites, number of generations of merogony, presence or absence of intermediate hosts, parasite host specificity, and production of cyst or pseudocyst forms within hosts. These variations will be dealt with when discussing the specific protozoans. Another common feature shared by these parasites is the apical complex, the defining criterion for inclusion in the phylum Apicomplexa. The apical complex refers to a group of organelles that are present in one or more stages of the protozoan's life cycle. These organellesmvisible by electron microscopyminclude the polar ring, conoid, micronemes, and rhoptries (Morrissette and Sibley 2002). The polar ring is a cogwheel-like, microtubule-organizing center from which microtubules radiate out of and down the length of the sporozoite. Engagement of these structures is thought to play a role in parasite gliding motility. The conoid is a retractable protuberance at the apical end of the parasite. It is composed of tightly coiled microtubules and is thought to be responsible for mechanical penetration of epithelial cell membranes by the invading sporozoite. Micronemes and rhoptries are secretory organelles containing proteins required for parasite motility, and adhesion to and invasion into host cells. Unlike the flagellates and amoebae, most apicomplexans possess
mitochondria and a Golgi complex. Most apicomplexans also possess an apicoplast, a chloroplast-like organelle homologous to the plastid of algae (Kohler et al. 1997; Roos et al. 1999). This organelle contains its own genomic material and is thought to have an endosymbiotic origin in apicomplexans similar to that of mitochondria in eukaryotic cells. Although its function is not completely understood, its presence is essential for parasite survival (Roos et al. 1999). In addition, because the genes on this episomal DNA appear prokaryotic in nature, it is an attractive target for drug development against the apicomplexans (McFadden 2003).
B. 1.
Eimeria spp.
Introduction
There is a surprisingly large body of literature regarding Eimeria infection in mice, especially when compared with other protozoal agents of rodents. Much of the early literature deals with species identification and life-cycle elucidation. Recent work focuses on Eimeria infection in mice as a model for investigating the immunopathogenesis of coccidiosis, a disease that accounts for large economic losses in the livestock industries. In general, members of the genus Eimeria are homoxenous (completing their life cycle in one host) and are host-specific (Fernando 1990; Levine and Ivens 1988). Most, but not all, are intestinal pathogens (Fernando 1990). Eighteen species of Eimeria have been described in Mus musculus, of which four of these (E. falciformis, E. vermiformis, E. papillata, and E. ferrisi) are considered pathogenic (Levine and Ivens 1990). Infection with Eimeria is rare in well-managed, laboratory mouse colonies. A recent study indicated a 26.3% infection rate with Eimeria spp. in a variety of wild rodents including Mus musculus (Franjola et al. 1995). 2.
Life Cycle
The life cycle of Eimeria spp. in mice is simple and direct, and follows the basic apicomplexan life-cycle scheme described above. Oocysts are shed in the feces and require 3 to 6 days in the environment to undergo sporulation (Levine and Ivens 1990). Sporulated oocysts contain four sporoblasts, each of which contains two infective sporozoites. After ingestion, sporozoites are released and invade the intestinal epithelium. As with other members in the genus Eimeria, those infecting mice have preferred host-cell and intestinal tract niches: E. falciformis infects the crypt epithelium of the cecum and colon (Fig. 21-4A); E. vermiformis, the crypt epithelium of the distal small intestine (Fig. 21-4B); E. papillata, the villus epithelium of the distal small intestine; and E. ferrisi, the villus epithelium of the cecum and colon (Table 21-1; Schito et al. 1996). E. falciformis undergoes four generations of asexual replication in the host, while E. vermiformis and E. ferrisi undergo three (Levine and Ivens 1990). The number
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Fig. 21-4 Eimeria spp. from the intestinal tract of mice. (A) Eimeria falciformis oocysts in the crypts of the proximal colon. (B) E. vermiformis oocysts in the crypts of the distal small intestine. (C) and (D) Mature meronts of E. falciformis and E. vermiformis, respectively. (E) E. falciformis microgametes (bottom of figure), and macrogametes (top of figure). A single macrogamete being fertilized by microgametes is present in the middle of the figure. (F) A cluster of E. vermiformis macrogametes. H&E; A and B, bar = 100 ~tm; C through F, bar = 50 ~tm.
21.
of rounds of asexual replication in E. papillata has not been reported. Sexual replication begins with differentiation of the parasites into microgametes and macrogametes. After fertilization, mature oocysts are released from host cells and shed in the feces to begin the cycle again. The infective dose for naive mice is 103 oocysts, and the prepatent period ranges from 4 to 8 days (Schito et al. 1996). Resistance to reinfection with Eimeria is dependent on the host mouse strain, immunocompetency, and initial parasite dose and species (Mahrt and Shi 1988; Mesfin and Bellamy 1979; Schito et al. 1996). 3.
Cell Biology
Eimeria species possess an apical complex, Golgi complex, mitochondria, and apicoplast (see Section IV.A; Adams and Todd 1983; Cai et al. 2003; Chobotar et al. 1975). Although the functions of the apicoplast genes are unknown, portions of its genome have been used to investigate phylogenetic relationships within the rodent lineages of Eimeria (Zhao and Duszynski 2001).
4.
531
PROTOZOA
Disease and Diagnosis
Four Eimeria species are considered pathogenic for mice: E. falciformis, E. vermiformis, E. ferrisi, and E. papillata. All result in similar clinical disease and histologic lesions. Concurrent infection with multiple species has been observed in mice (Allen and Fetterer 2002). Virulence factors have not been identified in Eimeriamdisease is due to the damage sustained by the intestinal epithelium during multiple rounds of parasite replication and rupture from the host cells. Clinical signs are dose-related and include soft, mucus-covered feces with perianal staining, depression, dehydration, and anorexia (Allen and Fetterer 2002; Blagburn and Todd 1984; Mesfin et al. 1978). In experimentally infected mice, mortality rates increase with doses of 103 and higher for E. falciformis and E. vermiformis oocysts (Blagburn and Todd 1984; Mesfin et al. 1978). Histologic lesions vary by intestinal location and include multifocal mucosal erosions of the intestinal epithelium with a pyogranulomatous cellular infiltrate and hemorrhage, submucosal congestion, and edema (Blagburn and Todd 1984; Levine and Ivens 1990; Mesfin et al. 1978). Intralesional parasites, at various stages of replication within intestinal epithelial cells, may be observed during acute infection. As infection resolves, a reduction in the intestinal villus to crypt ratio, villus shortening or crypt hyperplasia, and a nonspecific mononuclear cell infiltrate may be all that is observed (Blagburn and Todd 1984). Focal granulomas with oocysts in the lamina propria of the colon have been described in resolving E. falciformis infections (Mesfin et al. 1978). Hyperplasia of the mesenteric lymph nodes and splenic follicles may also be seen histologically (Smith and Hayday 2000a). Eimeria spp. are readily observed in hematoxylin-eosin stained histologic sections. Meronts (asexually replicating forms) are
found within intracellular, parasitophorous vacuoles. Meronts vary in number and size depending on their stage of development. Immature forms are round to indistinct in shape, while mature forms are crescent or banana-shaped (Fig. 21-4C, D). Most are uninucleate and lightly basophilic. Microgametes stain intensely basophilic, are comma-shaped, and possess two to three flagella (Fig. 21-4E). Macrogametes have a prominent nucleus and nucleolus. Refractile eosinophilic material or periodic acid-Schiff (PAS) positive staining material may be observed within the cytoplasm of the macrogamete (Fig. 21-4E, F). Oocysts can be distinguished from macrogametes by the presence of one or two refractile cell walls. Speciation of Eimeria parasites can be tentatively assigned based on the tissue and cell location of parasite infection (see Section IVB2). Eimeria spp. infection can also be diagnosed by fecal flotation, although speciation is difficult. In fresh fecal pellets, oocysts are round to ellipsoidal and vary considerably in size both within and between species. Most range in size between 18-26 x 11-24 gm (see Table 21-4; Levine and Ivens 1990). Sporulated oocysts are smaller (ranging in size between 8-14 x 5-10 gm), and contain four sporoblasts with two sporozoites each. Oocyst walls of E. falciformis and E. ferrisi are smooth and clear to light brown in color; those of E. vermiformis and E. papillata are yellowish brown and pitted or papillated (Levine and Ivens 1990). 5.
Treatment, Prevention, Control
A variety of anticoccidial drugs and coccidiostats are used in the livestock industry to control Eimeria infection (Allen and Fetterer 2002). The efficacy of some of these compounds has been examined in naturally infected mice, with varying results (Haberkorn et al. 1983). Toltrazuril (Baycox| was effective in eradicating a mixed Eimeria infection in C57BL and CFW1 mice, while amprolium, several ionophores, and sulfa-based drugs were only partially effective (Haberkorn et al. 1983). Efficacy was enhanced when some of these less effective drugs where given simultaneously (Harder and Haberkorn 1989). Autoclaving is sufficient to inactivate Eimeria oocysts. E. falciformis oocysts were found to be nonviable after 7 days at 40~ (summarized in Schneider et al. 1972). However, due to the large reproductive potential of Eimeria spp., oocyst numbers build up quickly in the environment and can be difficult to eliminate from some types of laboratory animal housing (Haberkorn et al. 1983; Wilkinson et al. 2001). Rederived and barrier-maintained mice housed in clean environments are free of coccidia. 6.
Research Implications
Outbred mice previously infected with E. falciformis were noted to be resistant to experimental infection with Salmonella abortus-ovis (Lantier et al. 1981). Survival time was found to be greater in mice previously infected with E. falciformis and
532
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experimentally inoculated with Toxoplasma gondii (Chinchilla et al. 1986). The authors of these reports suggest that Eimeria infection induces a nonspecific inflammatory response that provides protection against other pathogens, but state that the mechanisms involved are unknown. Due to immunologic stimulation and potential for severe intestinal pathology, mice infected with Eimeria spp. are unsuitable for most research involving the immune system and gastrointestinal tract. Experimental Eimeria spp. infection in mice, however, has served as a marvelous tool for investigating gut immune responses to a natural murine pathogen. The stimulus for this research is the hope of developing vaccines against these pathogens in poultry and cattle (Rose et al. 1997). Primary infection with Eimeria spp. in immunocompetent mice results in cellular and humoral immune responses and clearance of infection in approximately 3 weeks (Nash and Speer 1988; Smith and Hayday 2000a). Mice are resistant to reinfection with the same species. Experiments with genetically modified mice have allowed these responses to Eimeria spp. to be immunologically "dissected." The inability of athymic nude mice to develop immunity to Eimeria falciformis suggested that T cell responses were important in clearance of infection (Mesfin and Bellamy 1979). Using several knockout mouse models, it was later shown that major histocompatibility (MHC) II-restricted CD4+ ~[3 T cells and the production of interferon gamma were important for effective primary immune responses against Eimeria spp. (Roberts et al. 1996; Rose et al. 1989, 1992). Gamma delta (y/8) T cells, noncirculating lymphocytes found in the intestinal epithelium of mice, were found to have an immunomodulatory effect on Eimeria spp. infection. Mice deficient in 3'/8 T cell receptors have more severe intestinal pathology than mice deficient in y/8 and a/I] T cell receptors when infected with E. vermiformis (Roberts et al. 1996). The ability to mount an effective secondary immune response was not altered by the lack of y/8 T cells, however. Work done with severe combined immunodeficient-beige, perforin-knockout mice, and mice treated with monoclonal antibodies to deplete CD4+ or CD8+ T cells demonstrated that cytotoxins produced by natural killer cells and cytotoxic (CD8+) T cells play a role in immunity to secondary infection with E. papillata or E. vermiformis (Rose et al. 1992; Schito and Barta 1997). Lastly, infection of TAP1, interleukin-4, Fas ligand, and inducible nitric oxide knockout mice with E. vermiformis has demonstrated that MHC class I-restricted T cells, B cell activation, and respiratory burst activity of macrophages are not important for clearance of primary Eimeria spp. infections (Smith and Hayday 2000b). C. 1.
Sarcocystis muris
Introduction
Sarcocystis muris is a coccidial parasite of murine skeletal muscle (Miescher 1843). In the early part of the twentieth
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century, the prevalence of Sarcocystis infection in laboratory mice was reported to be between 14 and 60% (Smith 1901; Twort and Twort 1932). Although modem husbandry and management practices such as separation of species, vermin control, rederivation, and barrier housing have decreased the incidence of S. muris, this parasite is still occasionally observed in laboratory-bred mice (Tillmann et al. 1999). 2.
Life Cycle
The source of Sarcocystis muris infection in mice remained obscure until 1976, when cats were identified as the definitive host (Ruiz and Frenkel 1976). Limited cross-transmission studies demonstrate that Mus musculus is the only intermediate host for S. muris (Ruiz and Frenkel, 1976). The life cycle follows the basic scheme as described above with a few exceptions: Oocysts sporulate within the carnivore's intestinal tract and are immediately infective when released into the environment; sporulated oocysts contain two sporocysts, each of which contains four sporozoites; after ingestion of sporocysts by mice, asexual replication first occurs in the liver, then in the skeletal muscle; in the muscle, replication of parasites results in the formation of large cystic structures (sarcocysts) filled with thousands of organisms (bradyzoites). When infected skeletal muscle is consumed by the carnivore, bradyzoites are released, invade the host intestinal epithelium, and undergo sexual replication. This heteroxenous life cycle begins again with the release of sporocysts into the environment. Recent data suggest that S. muris can also be sustained within a mouse colony due to cannibalism of infected individuals (Koudela and Modry 2000). 3.
Cell Biology
Sarcocystis muris possess the standard complement of organelles found in apicomplexans, including an apical complex, Golgi complex, and mitochondria. An apicoplast organelle has been identified in S. muris by electron microscopy (Hackstein et al. 1995). Partial sequencing of this apicoplast genome revealed an herbicide-binding region, suggesting that S. muris may be susceptible to triazine chemotherapeutics such as toltrazuril (Hackstein et al. 1995).
4.
Disease and Diagnosis
Difficulty moving is the only clinical symptom associated with sarcocystosis in mice (Ruiz and Frenkel 1976). Mice of that report were experimentally inoculated, and approximately 80% of muscle fibers were infected with S. muris by 100 days post-inoculation (Ruiz and Frenkel 1976). A dose of 50 sporocysts is reported to result in widespread and severe muscle infection with S. muris in NMRI and AKR/N mice (Rommel et al. 1981). Natural infections in mice are usually not that severe (Tillmann et al. 1999; Twort and Twort 1932).
2 1.
533
PROTOZOA
Diagnosis of S. muris is made by histologic examination of skeletal muscle; organisms are occasionally observed in the myocardium (Tillmann et al. 1999; Twort and Twort 1932). Within muscle, spherical or cylindrical cysts--often several millimeters long--can be seen within myocytes. Hundreds of banana-shaped, uninucleate bradyzoites are present within the cysts. Often there is no inflammatory reaction to these structures. Bradyzoites are visible with routine hematoxylineosin and periodic acid-Schiff staining, and measure 4-6 x 14-16 ~tm (Ruiz and Frenkel 1976). Sporocysts are shed in the feces of infected cats and can be diagnosed by fecal floatation. Sporocysts measure 7.5-9 x 8.7-11.7 ~tm, contain four sporozoites, and have a smooth and colorless wall (Cawthorn and Speer 1990; Ruiz and Frenkel 1976). 5.
Prevention, Treatment, Control
There is a single report documenting elimination of S. muris from the livers of experimentally infected mice with sulfaquinoxaline and pyrimethamine (Rommel et al. 1981). It is not clear if this drug combination was effective against bradyzoite forms found in muscle. Other anticoccidial drugs were not effective (Rommel et al. 1981). Sporocysts are environmentally resistant and remain infectious for at least 119 days at 21~ in fecal flotation solutions (Smith and Frenkel 1978). Although Sarcocystis spp. have an obligatory heteroxenous life cycle (requiring definitive and intermediate hosts), direct contact with cats or cat feces is not needed to transmit S. muris to mice. Cockroaches exposed to S. muris-infected cat feces transmitted infection to naive mice for 20 days (Smith and Frenkel 1978). Cannibalism--although less efficient--will sustain S. muris in a mouse population (Koudela and Modry 2000). For these reasons, rederivation and maintenance in a clean barrier is recommended to eliminate S. muris from mouse colonies. 6.
Research Implications
Experimental infection with S. muris was found to suppress humoral and cell-mediated immune responses to vaccination with unrelated proteins, and to induce splenomegaly in mice (Gill et al. 1988a, 1988b). These nonspecific immune aberrations make infected mice unsuitable for research involving the immune system.
D. 1.
KlossieUa muris
Introduction
Klossiella muris is the least characterized apicomplexan parasite of mice. Infection in the kidneys of mice was first reported in 1889 and further described in 1902 (Smith and Johnson 1902). In a 1932 histologic survey of mice used in carcinogenesis studies, renal infection with K. muris was estimated to be 60% (Twort and Twort 1932). Infection in
wild-caught mice has been reported as high as 93% (Rosenmann and Morrison 1975). Under current laboratory animal housing conditions, infection with K. muris is seldom reported. 2.
Life Cycle
Infection in mice begins with ingestion of sporocysts and uptake into the portal vascular system (Yang and Grice 1964; Smith and Johnson 1902). Sporozoites are released, circulate through the vascular system, and preferentially invade the endothelium of the glomeruli, where they undergo several rounds of asexual replication (Fig. 21-5A). Parasites have also been observed in capillaries and arterioles of the lungs, liver, spleen, and thymus (Twort and Twort 1932). At some point, asexually replicating forms rupture from the glomeruli into Bowman's space and pass to the renal tubules. Parasites invade the tubular epithelium and differentiate into microand macrogametes. Fertilization results in the formation of sporonts, each of which contains as many as 30 sporoblasts and a residual body (Fig. 21-5B; Levine and Ivens 1990; Smith and Johnson 1902; Yang and Grice 1964). Sporoblasts mature into sporocysts, each of which in turn contains 25 to 35 crescent-shaped sporozoites. The life cycle is completed when infective sporocysts are released from ruptured host cells, pass into the urinary bladder, and are shed into the environment. A homoxenous life cycle has been inferred due to the large number of infected mice diagnosed histologically after remaining caged together for several months (Smith and Johnson 1902). 3.
Cell Biology
Ultrastructural examination of Klossiella life-cycle stages and organelles has not been reported. They presumably possess components of the apical complex found in other apicomplexans. The means by which they identify and invade host cells, generate energy, and acquire nutrients for replication have not been investigated. 4.
Disease and Diagnosis
Clinical disease has not been reported with K. muris infection in mice. Diagnosis is usually made histologically and is dependent on identifying replicating forms within the glomerular endothelium or renal tubular epithelium (Smith and Johnson 1902; Yang and Grice 1964). Asexually replicating forms (merozoites) often cause distention of endothelial cells and may appear as multiple minute (< 0.5 ~tm) bodies surrounded by clear halos within a parasitophorous vacuole in the cell (Yang and Grice 1964). Sporocysts are subspherical, measuring 16 x 13 ~tm, and are located in the renal tubule epithelium (Levine and Ivens 1990; Yang and Grice 1964). Multiple banana-shaped sporozoites may be seen budding within each sporocyst (Levine and Ivens 1990; Yang and Grice 1964). Heavily parasitized kidneys show evidence of nonsuppurative
534
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Fig. 21-5 Klossiella muris from the kidneys of a mouse. (A) Merozoites replicating in endothelial cells of a glomerulus. (B) Sporont within renal tubular epithelial cell; note budding sporoblasts and residual body. (C) High magnification of mature sporocysts in the renal tubular epithelium. (D) Low magnification of K. muris infection in the renal medulla of a mouse. H&E; A through D, final magnification unknown (Images courtesy of S. W. Barthold).
interstitial nephritis with tubular degeneration. Often, there is no inflammatory response associated with intracellular stages (Yang and Grice 1964). Organisms are periodic acid-Schiff negative but readily identifiable by routine hematoxylin-eosin staining (Fig. 21-5C, D; Yang and Grice 1964). Clinical diagnosis of Klossiella equi by examination of urine has been described and should be possible for diagnosis in mice (Reppas and Collins 1995). However, the authors of that report note that sporocysts were destroyed by fecal flotation solutions and that organisms were apparent only after examining the pellet of centrifuged urine samples (Reppas and Collins 1995).
5.
Prevention, Treatment, Control
Treatment of endemic K. muris infection with coccidiostats reduced the histologic incidence of infection in mice from 93. to 23% (Rosenmann and Morrison 1975). Unfortunately, that report does not specify the compound, dose, or route used. Infection does not occur in rederived, barrier-maintained mouse colonies.
6.
Research Implications
A single report documents the effects of K. muris infection in mice on research results (Rosenmann and Morrison 1975). Endemically infected mice exhibited decreased oxygen consumption and endurance when compared with uninfected controls. The authors suggest that under certain environmental conditions, K. muris infection impairs the metabolic capabilities of infected mice.
E. 1.
Toxoplasma gondii
Introduction
Toxoplasma gondii was first described in 1908, in Ctenodactylus gundii (or "gundi"), a guinea pig-like rodent found in North Africa (Nicolle and Manceaux 1908). Although T. gondii was subsequently shown to cause disease in a variety of animals around the world, it took 50 years to identify the cat as the definitive host (Hutchison 1965). Hutchison correctly
21.
surmised that cats excreted infective forms in their stools. However, concurrent Isospora spp. infection in his experimental cats obscured identification of T. gondii oocysts on fecal examination. In addition, the presence of Toxocara cati ova in the feces led him to conclude that T. gondii was transmitted in conjunction with this common nematode of cats (Hutchison 1965). This was quickly rectified once parasite-free cats were used for transmission experiments (Sheffield and Melton 1969). Mice of the genus Mus are but one of several hundred mammals (including humans), birds, and reptiles since identified as intermediate hosts in the complex life cycle and biology of this protozoal parasite. Natural infection of most animals (including mice) with T. gondii is subclinical. Because of the pervasiveness of this parasite and its role in human and animal disease on a global basis, there is a tremendous amount of published research on various aspects of toxoplasmosis. Mice have played an integral role in understanding the virulence and pathogenesis of this organism. 2.
535
PROTOZOA
Life Cycle
The life cycle of T. gondii is perhaps the most complicated of all the apicomplexans of mice. Oocysts are shed in the feces of infected cat s, and sporulate in the environment after several days. Sporulated oocysts contain two sporocysts with four sporozoites each and can be differentiated on fecal floatation from Isospora spp. by size: T. gondii oocysts measure 12 x 10 ~m, while those of Isospora spp. are larger: I. rivolta measure 22 x 17 and I. felis 42 x 35 ~tm (Foyet 2001). After ingestion, sporozoites are released in the small intestine of intermediate hosts. In experimentally infected mice, sporozoites can be identified in cells of the ileal lamina propria as early as 2 hours post-inoculation (PI; Dubey, Speer et al. 1997). After invading a suitable host cell, sporozoites differentiate into tachyzoites and replicate asexually for an infinite number of generations. These rapidly replicating ("tachy-" = fast) asexual forms are responsible for "acute" toxoplasmosis. From the small intestine, parasites invade additional tissues after being transported by blood or lymph. Unlike other apicomplexans, T. gondii can infect virtually any nucleated cell in the host. In experimentally infected mice, tachyzoites can be identified in the mesenteric lymph nodes 8 hours PI (Dubey et al. 1997). By day 3 PI, parasites are present in the lungs, spleen, and kidneys; by day 4, in the heart and liver; by day 6 in the pancreas and brain; and by day 7, in the skeletal muscle (Dubey et al. 1997). If the host survives acute infection, most parasites in these peripheral locations are cleared by the immune response. This immune pressure is thought to stimulate tachyzoites to differentiate into bradyzoites within a protective tissue cyst wall. These cyst forms are typically found in the central nervous system, or skeletal or cardiac muscle (Dubey et al. 1997). In experimentally infected mice, tissue cysts are observed by day 15 PI (Dubey et al. 1997). These slowly replicating ("brady-"= slow) asexual forms are responsible for "chronic" toxoplasmosis in
intermediate hosts. Ingestion of tissue cysts by felids results in release of bradyzoites, invasion into intestinal epithelial cells, and differentiation into the sexually replicating forms of the parasite. As with the other coccidians, fertilization of macrogametes by microgametes results in the formation of oocysts and completion of the life cycle. Although the life cycle of T. gondii is considered heteroxenous, mice can sustain T. gondii infection through cannibalism or congenital transmission, in addition to ingestion of sporulated oocysts. Stage conversion between tachyzoites, bradyzoites, and oocysts can be demonstrated in vitro (Dubey et al. 1998). However, in vivo generation and collection of oocyst or tissue cysts from cats or mice, respectively, are usually required to generate sufficient numbers of organisms for experimental work. 3.
Cell Biology
T. gondii possesses an apical complex, mitochondrion, Golgi complex, and apicoplast similar to the other apicomplexans (see Section IV. A). The large complement of secretory organelles (micronemes and rhoptries of the apical complex, dense granules) and their functions are best--though not completelyiunderstood in T. gondii. The evolution of this extensive network of secretory organelles is thought to be an adaptation to an obligate intracellular lifestyle. Micronemes release proteins involved in the early stages of tachyzoite attachment and invasion into host cells (Carruthers et al. 1999). Depletion of microneme proteins results in transient loss of tachyzoite infectivity. Rhoptries are distinctive, club-shaped organelles numbering 8 to 16 per tachyzoite. Rhoptry proteins are secreted later in the host-cell invasion process; some are also incorporated into the parasitophorous vacuole that encloses tachyzoites once within the host cell (Carruthers and Sibley 1997). Dense-granule proteins are secreted late in the invasion process. These proteins insert in the parasitophorous membrane and are thought to be involved with nutrient acquisition and importation from the host cytoplasm to the replicating tachyzoites (Carruthers 1999). Formation of these organelles, and the biogenesis, trafficking, processing, storage, and secretion of their respective protein cargos suggest that an intricate and complex network of secretory pathways are present in apicomplexans. These pathways must be executed in a precise and sequential manner for parasite survival. A second set of unique organelles found in T. gondii are the acidocalcisomes (Moreno and Zhong 1996). First identified in Trypanosoma cruzi, these structures have since been identified in other trypanosomatids, apicomplexans, algae, slime molds, and the bacterium Agrobacterium tumifaciens (Docampo et al. 1995; Docampo and Moreno 2001; Seufferheld et al. 2003). As the name implies, these organelles contain acidified stores of calcium and other elemental minerals. Although the exact function of acidocalcisomes is not known--and may vary between species--several functions have been proposed. These include as an energy source, an intracellular calcium store for
536
parasite signaling pathways, a mechanism for intracellular pH homeostasis, and for parasite osmoregulation (Docampo and Moreno 1999). In addition, since acidocalcisomes are not present in mammalian cells, they represent an attractive target for antiprotozoal therapies.
4.
Disease and Diagnosis
T. gondii infection in mice is subclinical with minimal gross necropsy lesions (Perrin 1942). Virulence of experimental infection is dependent on mouse strain, T. gondii-type strain, stage of parasite (sporulated oocysts being more pathogenic than bradyzoites, which in turn are more pathogenic than tachyzoites), and parasite dose (Araujo et al. 1976; Suzuki et al. 1995). Swiss Webster mice inoculated with the mildly pathogenic type III strain of sporulated oocysts remained clinically normal until 3 weeks PI, at which time they appeared unthrifty, lost weight, and developed paralysis (Dubey et al. 1997). Mice euthanized at early time points had enlarged and edematous mesenteric lymph nodes. The ileum was congested and edematous, with small white pinpoint foci visible through the serosa. Necrosis was the prominent histologic lesion, with the organs involved dependent on the time point of infection at which mice were euthanized. Necrosis and infarction of the ileal lamina propria and mesenteric lymph nodes were seen in the first days of infection. Focal hepatitis and myocarditis with mixed leukocytic infiltration were observed during the first week of infection. Interstitial pneumonia with intralesional parasites developed during the second week of infection. Intra- and extracellular tachyzoites were seen in brain parenchyma as early as 9 days PI. By 87 days after inoculation, bradyzoite-filled tissue cysts with little associated inflammation were present in most mouse brains (Dubey et al. 1997). Diagnosis of toxoplasmosis in mice is primarily by histology. Individual tachy- and bradyzoites are difficult to identify in histologic sections, but tissue cysts may be observed in the CNS, myocardium, or skeletal muscle (Fig. 21-6A). Cysts are spherical and of variable size, with a thin wall that is argyrophilic and faintly periodic acid-Schiff (PAS) positive (Frenkel 1956). Cysts are packed with PAS positive, fusiform bradyzoites. Unstained impression smears of brain material can also be screened for the presence of T. gondii tissue cysts (Dubey et al. 1998). In intermediate hosts with patent parasitemia, individual tachyzoites may be identified in the lamina propria of the small intestine, endothelium of the small intestine or lungs, or within leukocytes on peripheral blood smears. Pre-cyst intracellular clusters of tachyzoites may also be observed in heart and skeletal muscles (Fig. 21-6B). Tachyzoites have a centrally located nucleus and measure 2-3 x 6-7 ~tm (Moller 1968). Tachyzoites exhibit a gliding motility that is usually only appreciated in tissue culture systems. T. gondii infection in mice can also be diagnosed by PCR and serology (James et al. 1994).
KATHERINE 5.
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Treatment, Prevention, Control
Because of a short shedding period and subclinical signs, cats with T. gondii infection usually do not receive treatment (Dubey 1994). Acute toxoplasmosis in intermediate hosts is amenable to chemotherapeutic intervention, usually a combination of pyrimethamine and sulfa drugs (Wilson et al. 2003). Tissue cyst forms are resistant to treatment, however. Experimentally infected mice treated with sulfonamides for 15 days had a lower mortality rate than those treated with chlortetracycline or left untreated (Frenkel 1956). In general, treatment of T. gondii infection in intermediate hosts is not warranted. Prevention of infection in rodent colonies relies on separation of species and elimination of potential transport vectors from the environment (Chinchilla et al. 1994). Because T. gondii can also be sustained through cannibalism and congenital infection, rederivation and barrier maintenance are recommended for naturally infected mouse colonies (Beverley 1959). As with other apicomplexans, the oocysts of T. gondii are remarkably hardy. Oocysts retain infectivity when stored at 4~ for 54 months or 0~ for 13 months; oocysts lose infectivity at temperatures above 40~ (Dubey 1998). Oocysts also survive exposure to 5.25% sodium hypochlorite (bleach) solution (Dubey et al. 1997). This method is used to "sterilize" oocysts harvested from cat feces prior to tissue culture or mouse inoculation. Tachyzoites and bradyzoites are less environmentally stable, although tissue cysts (filled with bradyzoites) retain infectivity when stored at 4~ for several months (Dubey 1997). Autoclaving or heat treatment at 70~ for 10 minutes will inactivate T. gondii oocysts from bedding, caging, and other equipment (Dubey 1994). 6.
Research Implications
Interference with research due to endemic toxoplasmosis has not been reported for mice. Several studies examining the behavioral effects of experimental T. gondii infection in mice have been published, however, and suggest that subclinical infection would interfere with behavioral phenotyping of mice. Experimentally and congenitally infected mice exhibited decreased motor coordination, spent less time grooming and exploring novel areas, and spent more time running on a home-cage wheel than uninfected control mice (Hay et al. 1983, 1984, 1985). The authors attributed these behavioral abnormalities to subclinical T. gondii-induced encephalitis in the otherwise clinically healthy mice, and suggested that this behavior increased their chance of predation (and therefore perpetuation of toxoplasmosis) by cats (Hay et al. 1985). More recent experimental mouse work has focused on three areas of importance in human toxoplasmosis: ocular, encephalitic, and congenital toxoplasmosis. Ocular toxoplasmosis in humans is thought to result from reactivation of congenital T. gondii infection, and occurs in both immunocompetent and
21.
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537
Fig. 21-6 Toxoplasma gondii tissue cyst in the cerebellum (A) and tachyzoites in the heart muscle (B) of a wallaby. Note the difference in shape and size of bradyzoites within the mature cyst compared to the intracellular tachyzoites. (C) Cryptosporidium parvum in the small intestine of a mouse. (D) Higher magnification of C. parvum. Note the variation in size of the merozoites and their intracellular but extracytoplasmic location in the enterocytes. H&E; A, B and D, bar = 50 ~m; C, bar = 100 ~tm (Images A, B courtesy of S. Tunev).
538
immunosuppressed individuals. Ocular toxoplasmosis causes a recurrent and progressive, necrotizing retinochoroiditis that eventually results in blindness of the affected eye (Gutierrez 2000). The disease can be experimentally reproduced in mice by intraperitoneal injection of tissue cysts or by intraocular injection of tachyzoites (Hu et al. 1999; Lyons et al. 2001). Using the mouse model, researchers have shown that T. gondii infection results in an upregulation of both inflammatory and anti-inflammatory mediators of the immune system and in apoptotic pathways in ocular tissues (Hu et al. 1999; Lyons et al. 2001). These responses may serve as a mechanism to minimize inflammation in the otherwise immune-privileged eye by eliminating parasite-infected cells while preserving uninfected tissues. As with ocular disease, toxoplasmic encephalitis results from reactivation of a congenital or subclinical infection. Toxoplasmic encephalitis tends to occur in profoundly immunosuppressed individuals and generally carries a poor prognosis (Suzuki 2002). A murine model of reactivated toxoplasmosis can be created by inoculating virulent strains of T. gondii into sulfadiazine-treated mice. Antibiotic treatment suppresses--but does not eradicate--parasite infection. This prevents development of acute toxoplasmosis while allowing the infection to establish itself in brain and muscle tissues. "Reactivation" of infection (renewed proliferation of tachyzoites) and development of fatal encephalitis occur when antibiotics are discontinued. Using resistant, susceptible, and genetically modified mouse strains, researchers have demonstrated that the source of gamma interferon (INF-'f), an immune mediator critical for preventing toxoplasmic encephalitis, is produced by T cells and an as-of-yet unidentified population of non-T cells within the brain (Kang and Suzuki 2001). This non-T cell source of INF-), appears critical for host resistance against T. gondii encephalitis. Mice have also been key to determining the genetics of resistance to toxoplasmic encephalitis. Mice with the d haplotype (e.g., BALB/c mice) in the D region of the major histocompatibility complex (MHC) are resistant, while those with the b or k haplotype (C57BL/6 mice) are susceptible (Suzuki et al. 1994). This correlates with what is seen in human infection. Humans with the HLA-DQ3 haplotype of the MHC are more susceptible to toxoplasmic encephalitis than those with the HLA-DQ 1 haplotype. Mice engineered to express human HLA-DQ1 had a greater degree of protection against toxoplasmic encephalitis compared with those expressing human HLA-DQ3 (Mack et al. 1999). Lastly, mice have been used as a model to investigate aspects of congenital transmission of toxoplasmosis. BALB/c mice infected with tissue cysts on day 12 of gestation--but not 8 weeks prior to breeding, or at 8 weeks prior to and on day 12 of gestation--gave birth to T. gondii-infected pups (Roberts and Alexander 1992). Pups had histologic evidence of toxoplasmic encephalitis, and some developed ocular disease at 2 months of age. Using this model, researchers have demonstrated the importance of natural killer cells--an important
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component of the innate immune response--in protecting the fetus from congenital transmission of T. gondii (Abou-Bacar et al. 2004). This model has also been useful for evaluating the efficacy of candidate vaccines against toxoplasmosis (Roberts et al. 1994). This na'fve pregnant mouse model replicates several of the epidemiologic and clinical features of human congenital toxoplasmosis. These include infection rates that are highest in fetuses born to mothers exposed to T. gondii for the first time during pregnancy; protection of fetuses against infection if mothers have been previously exposed to T. gondii; and a high incidence of ophthalmic and neurologic disorders in congenitally infected infants (Gutierrez 2000).
E 1.
Cryptosporidium muris
Introduction
Cryptosporidium muris is the type species for a genus of parasites that were considered biomedical curiosities until the advent of HIV and AIDS (Current and Garcia 1991). Shortly after he identified C. muris in the gastric glands of mice, Ernest Tyzzer went on to identify and describe Cryptosporidium parvum, a second species that resided in the small intestine of mice (Tyzzer 1910, 1912). It wasn't until 70 years after Tyzzer's first descriptions that Cryptosporidium spp. were recognized as a cause of self-limiting diarrhea in immunocompetent humans and of severe life-threatening diarrhea in immunocompromised patients (Peterson 1992). There are now 13 validated Cryptosporidium species described, and most of these infect a wide range of hosts (Table 21-2; Xiao et al. 2004). Humans are the primary host for C. hominis but can be infected with various isolates of C. parvum and C. muris (Palmer et al. 2003; Xiao et al. 2004). Cryptosporidial infection of contemporary mouse colonies is seldom reported; natural infection with C. muris and/or C. parvum is occasionally observed in wild rodent populations (Bajer et al. 2003; Klesius et al. 1986; Torres et al. 2000).
2.
Life Cycle
Tyzzer's 1910 light microscopic description of the life cycle and morphology of C. muris is remarkably accurate (Tyzzer 1910). Despite the parasite's small size, he determined that the parasite had asexual and sexual stages, formed a unique organ of attachment to the host cell, produced two types of oocysts, and was likely related to coccidia (Tyzzer 1910). Seventy-five years later, electron microscopy confirmed Tyzzer's findings, with few exceptions (Current and Reese 1986). Like Eimeria, Cryptosporidium spp. are homoxenous and follow the general apicomplexan life cycle described in Section IV. A. Sporulated oocysts are shed in feces. Once consumed, four sporozoites excyst and invade the gastrointestinal epithelium. These develop within a parasitophorous vacuole in the host cell
2 1.
into type I merozoites. Type I merozoites replicate asexually, rupture from the host cell, and re-invade neighboring epithelium, developing into either type I or type II merozoites. Type I merozoites continue the asexual replication cycle, while type II merozoites develop into micro- or macrogametes. After fertilization by microgametes, macrogametes develop either into thick-walled or thin-walled oocysts. Oocysts undergo sporulation and contain four "naked" sporozoites prior to rupture from the host cell (there are no sporocysts). Thick-walled oocysts are shed as fully infective parasites into the environment; thin-walled oocysts excyst while still in the host, infect new gastrointestinal epithelial cells, and initiate the asexual replication cycle again (Tyzzer 1910). The autoinfective ability of type II merozoites and thin-walled oocysts can result in an overwhelming parasitism despite a relatively small, initial inoculum, particularly in the immunocompromised host (Current and Garcia 1991). C. muris infects and replicates in the gastric mucosa, while C. parvum infects small intestinal epithelium (Current and Reese 1986; Tyzzer 1910). The complete life cycle of Cryptosporidium spp. has yet to be replicated in vitro. Oocysts for in vitro experimentation are usually purified from the feces of experimentally inoculated neonatal ruminants or suckling mice (Arrowood 2002; Current 1990). Oocysts can be induced to excyst upon exposure to sodium hypochlorite (bleach) or trypsin and bile salts, and the released sporozoites will infect a number of different cell culture lines and replicate asexually (Arrowood 2002). However, production of significant numbers of mature oocysts in vitro has not been reported (Arrowood 2002). 3.
539
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Cell Biology
Cryptosporidium spp. possess an apical complex, complete with preconoid rings, conoid, rhoptries, and micronemes, which warrants their inclusion in the phylum Sporozoa (Current and Reese 1986). However, they lack a Golgi complex, mitochondrion, and the apicoplast present in other coccidia (Abrahamsen et al. 2004; Current and Reese 1986; Zhu et al. 2000). In addition, they inhabit a unique intracellular, extracytoplasmic location in their host cells (Current and Reese 1986; Goebel and Brandler 1982). At the interface between the parasite and host-cell cytoplasm, a unique "feeding organelle" can be seen by electron microscopy (Current and Reese 1986). This organelle is present during asexual and sexual parasite replication, and is composed of a portion of the parasitophorous vacuole membrane and a microfilamentous sheet (Current and Reese 1986). As the name implies, this organelle is thought to be important in nutrient transport from the host cytoplasm to the developing parasite (Current and Reese 1986). Recent work demonstrates that this organelle is composed, in part, of a membrane transporter protein that belongs in the ATP-binding cassette protein superfamily. This superfamily of proteins regulates the flow of diverse compounds across cell membranes (Perkins, Riojas et al. 1999). Recent publication of the
C parvum genome reveals that these parasites also possess several "plant-like" and bacterial-type enzymes that are either absent or divergent from those found in mammals (Abrahamsen et al. 2004). These proteins are likely to be good candidates in the search for drug targets against human and animal cryptosporidiosis.
4.
Disease and Diagnosis
Suckling mice infected with C muris or C. parvum are reported to grow more slowly and are less active than their uninfected littermates (Current and Reese 1986; Tyzzer 1910). Neonatal mice are susceptible to experimental infection with C. parvum up until 14 days of age, at which time they clear and remain resistant to infection (Novak and Sterling 1991). Clinical signs of C. muris infection in immunocompetent adult mice are not reported, and it is not clear if an age-related clearance of infection occurs for this species (McDonald et al. 1996; Tyzzer 1910). Experimental infection of immunosuppressed mice results in persistent infection with clinical signs of weight loss and sticky stools (Perryman and Bjorneby 1991; Ungar et al. 1990). Diagnosis is made by histologic examination of gastrointestinal tract tissue. Dilation of gastric glands and minimal lymphoid infiltrate are seen in mice infected with C. muris (Tyzzer 1910). In heavy C. parvum infection, blunting and fusion of intestinal villi, crypt hyperplasia, and moderate lymphocytic-plasmacytic infiltration of the underlying lamina propria are seen (Current and Reese 1986). In athymic nude mice, C. parvum can also be seen infecting the hepatobiliary tree and pancreatic ducts (Ungar et al. 1990). Small, round, basophilic-staining parasites may be observed by routine hemoxylin-eosin staining of histologic sections of the gastrointestinal mucosa. Life-cycle stages are difficult to differentiate by light microscopy due to their small size. Meronts and oocysts of C. muris measure 5 x 7 ~tm and are located on the luminal surface of gastric epithelium (Tyzzer 1910). Similar stages of C. parvum measure 5 ~tm or less in diameter and are found "decorating" the brush border of ileal enterocytes (Fig. 21-6C, D; Current and Reese 1986). These differences in size and tissue location can be used to differentiate between the two species in mice (Perryman 1990). Diagnosis can also be made by fecal flotation. Oocysts are spherical or ellipsoidal, with a smooth colorless wall and a faint suture line running down one-half the oocyst length. C. muris oocysts measure 7-8 x 5-6.5 ~tm; those of C. parvum are smaller, measuring 4.5-5 x 4-5 ~tm (Levine and Ivens 1990). 5.
Prevention, Treatment, Control
In immunocompetent animals and humans, infection with Cryptosporidium spp. is self-limiting. Although treatment of laboratory mice for natural infection with C. muris or C. parvum
has not been reported, suckling and immunosuppressed mice have been a useful in vivo model to test anticryptosporidial
540
compounds (see Blagbum and Soave 1997 for a summary of literature). Fifteen common anticoccidial compounds (including amprolium and sulfaquinoxaline) were found to be ineffective in protecting neonatal mice against infection with C. parvum (Angus et al. 1994). Lack of the apicoplast organelle in Cryptosporidium spp. has been hypothesized as one reason why these parasites are not susceptible to chemotherapeutics effective in other apicomplexans (Abrahamsen et al. 2004; Zhu et al. 2000). In addition to a lack of effective drug treatment, Cryptosporidium oocysts remain infective in chlorinated water (Carpenter et al. 1999; Korich et al. 1990). Oocysts are also remarkably resistant to common disinfectants. In an experiment examining the infectivity of oocysts after 18 hours incubation in various agents, only those treated with 10% formalin or 5% ammonia were noninfective for the suckling mouse model (Campbell et al. 1982). Steam sterilization, pasteurization, and ethylene oxide exposure for 24 hours will inactivate oocysts (Fayer et al. 1996; Harp et al. 1996). However, oocysts retain their infectivity when stored in 2% potassium dichromate solution at 4~ for greater than 6 months (Arrowood 2002). Because of a lack of treatment, resistance to common disinfectants, the environmental stability of oocysts, and demonstrated zoonotic potential of Cryptosporidium spp., endemically infected laboratory mouse colonies should be rederived and barrier maintained. 6. ResearchImplications Interference with research results due to natural infection in mice with C. muris has not been reported. In addition, C. parvum is thought to be primarily a pathogen of ruminants and humans, and is not likely to be identified in barrier-maintained mouse colonies (Klesius et al. 1986; Xiao et al. 2004). On the other hand, suckling mice have been useful for generating small amounts of pure isolates of Cryptosporidium parvum oocysts for in vitro work (Current 1990). Although Cryptosporidium spp. are not pathogenic for mice, work done in immunocompromised mouse models has helped elucidate the immune mechanisms responsible for clearance of infection. Similar to infection with Eimeria spp., clearance of C. muris and C. parvum infection was found to be dependent on production of interferon gamma and presence of CD4+ intraepithelial T cells (Chen et al. 1993; Culshaw et al. 1997; McDonald et al. 1996). The course of infection in mice with C. muris is also dependent on major histocompatibility complex (MHC) types. BALB/c mice (with the H-2d MHC type) were shown to produce fewer oocysts and to recover from infection with C. muris faster than BALB/B (H-2b) or BALB/K (H-2k) mice (Davami et al. 1997). This was attributed to a difference in interleukin-4 secretion by splenocytes between these strains of mice (Davami et al. 1997). Lastly, patent infection with Cryptosporidium spp. appears to depend on the host intestinal microflora. Axenic C.B-17/IcrTac-Prkdc sCid mice
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shed oocysts sooner when infected with C. parvum than their cohorts bearing intestinal flora (Harp et al. 1992). The authors speculate that resistance to initial infection is due to competition by endogenous microflora for preferred intestinal niches.
V. A.
MICROSPORIDIA
Encephalitozoon cuniculi
1. Introduction Mice can be naturally and experimentally infected with Encephalitozoon cuniculi, the type species for a burgeoning
collection of parasitic organisms that are more closely related to fungi than protozoans. Although frequently associated with rabbits and mice, E. cuniculi is a much more promiscuous organism than originally thought and has been identified as a pathogen in hamsters and guinea pigs, squirrel monkeys and cotton-top tamarins, carnivores (including fox and mink), domestic dogs and cats, goats, sheep, pigs, horses, muskrats, and shrews; and is a suspected cause of equine abortion (PattersonKane et al. 2003; Reetz et al. 2004; Wasson and Peper 2000; Wasson and Zbka 2003). It is also a pathogen of immunocompromised and immunocompetent humans (De Groote et al. 1995a; Mertens et al. 1997; van Gool et al. 2004). E. cuniculi infection was first described in rabbits in 1922 and in mice two years later (Cowdry and Nicholson 1924; Wright and Craighead 1922). E. cuniculi was reported in laboratory mice frequently thereafter, often in conjunction with additional parasite infestations including fleas, lice, pinworms, cysticerci, K. muris, S. muris, and T. gondii, and subclinical bacterial and--in all likelihoodmviral infections (Innes et al. 1962; Lackey et al. 1953; Morris et al. 1956; Perrin 1942; Perrin 1943; Twort and Twort 1932). In several of these reports, granulomatous hepatitis or meningoencephalitis associated with endemic E. cuniculi confounded attempts to develop murine models of human viral hepatitis or encephalitis, respectively. It was estimated that E. cuniculi was prevalent in 15 to 50% of laboratory mouse colonies at this time (Innes et al. 1962; Twort and Twort 1932). This older literature variably refers to this organism as "haplosporidia," "mouse ascitic agent," Nosema muris, E. muris, or E. rabii (Lainson et al. 1964; Morris et al. 1956; Twort and Twort 1932). By 1964, the name Nosema cuniculi was proposed, to reflect its morphologic similarity by light microscopy to the familiar Nosema spp. parasites of insects (Weiser 1964). However, ultrastructural comparison of E. cuniculi with Nosema spp. confirmed the former's status as a separate genus and reaffirmed its original name as given by Levaditi et al. 1923 (Levaditi et al. 1923; Pakes et al. 1975; Sprague and Vernick 1971). Until 1985, E. cuniculi was considered the only microsporidial parasite of mammals. With the arrival of HIV and AIDS, six generamrepresenting approximately
2 1.
12 speciesmhave since been identified as human pathogens (Franzen and Muller 1999). Mice appear to be natural hosts for E. cuniculi only. However, they can be experimentally infected with several of these newly described species. And although rare, E. cuniculi infection is still reported in contemporary mouse colonies (El Nass et al. 1998). 2.
Life Cycle
E. cuniculi is an obligate intracellular parasite with a simple and direct life cycle, and exists outside the host as an environmentally resistant spore (Wasson and Peper 2000). The initial site of E. cuniculi infection in mice is not known but is thought to be the intestinal epithelium. Once ingested, E. cuniculi infects susceptible host cells through the deployment of a preformed polar tube (discussed below). This polar tube is extruded from the spore, penetrates a susceptible host-cell membrane, and deposits the spore's infectious sporoplasm in the host-cell cytoplasm. The sporoplasm proliferates, resulting in the production of numerous plasmodial cells (also referred to as meronts) within a parasitophorous vacuole. These cells are amorphous with an indistinct nucleus and a simple cytoplasm when examined by transmission electron microscopy. Plasmodial cells develop into sporonts as the cell-limiting membrane (plasmalemma) becomes electron dense and the nucleus more distinct. Sporonts undergo binary fission to produce sporoblasts, which are no longer capable of dividing. Sporoblasts begin depositing organelles, including the polar tube and its associated polaroplast membrane, endoplasmic reticulum, the posterior vacuole, ribosomes, and electron dense bodies. Mature spores can be distinguished from sporoblasts by the presence of a thick spore wall, distinct cross sections through the coiled polar tube, and increased density of the spore cytoplasm. As nascent spores mature and increase in number, the host cell becomes distended and ruptures. Spores are released into the surrounding tissues to infect neighboring cells. E. cuniculi is capable of infecting a variety of host organs, although the mechanisms by which dissemination to these sites occur are not clear. In naturally infected, immunocompetent mice, parasites and associated inflammation are usually observed in the CNS and kidneys. Spores are shed in the environment through urine to complete the life cycle. E. cuniculi is also easily propagated in tissue culture, resulting in release of fully infective spores from a variety of mammalian cell lines and generating sufficient numbers of parasites for in vitro work (Visvesvara 2002).
3.
541
PROTOZOA
Cell Biology
Microsporidia contain a curious mix of eukaryotic, prokaryotelike, and novel organelles. Like all eukaryotic organisms, they possess a membrane-bound nucleus, endoplasmic reticulum, Golgi apparatus, and membrane-bound vesicles. Although they lack mitochondria, a mitochondrial remnantmtermed
mitosomemhas been identified (Williams et al. 2002). The function of the mitosome is not clear, as microsporidia lack the mitochondrial enzymes associated with aerobic metabolism. All life stages of microsporidia are replete with ribosomes. These ribosomes are smaller than typical eukaryotic ribosomes, with a sedimentation rate and molecular organization similar to those found in prokaryotes. The defining criterion for inclusion into the phylum Microspora is possession of the polar tube and its associated structures. Also referred to as a polar filament, this unique apparatus is the means by which parasites invade and infect susceptible host cells. The polar tube is preformed and coiled around the sporoplasm within mature microsporidial spores. This tube is attached at the anterior end of the spore by the means of an anchoring disk and is surrounded by lamellar and vesicular membranes (also referred to as the polaroplast). At the opposite end of the spore is a membrane-bound posterior vacuole. After activation by some yet-to-be-determined signal (or signals), the polar tube is forcefully ejected from the spore, uncoiling and penetrating a susceptible host cell in the process. Once fully ejected, the spore's sporoplasm (containing the nucleus and abundant ribosomes) travels through the tube and is deposited within the host-cell cytoplasm. The function of the polaroplast is not clear, but it may contribute to the limiting membrane of the discharged sporoplasm (Weidner et al. 1984). It is also not clear what role the posterior vacuole plays. However, this vacuole is seen to swell by electron microscopy during the course of polar tube extrusion and may provide additional osmotic force to propel the sporoplasm through the polar tube and into the host cytoplasm (Lom and Vavra 1963). Although the mechanisms behind this sequence of events are still being worked out, it appears that calcium signaling and water flux across the spore wall play key roles in the activation of this novel parasite invasion apparatus (Frixione et al. 1997; Weidner and Byrd 1982). 4.
Disease and Diagnosis
E. cuniculi does not appear to be transmitted vertically in mice, nor are neonates more susceptible to infection than adults (Liu et al. 1988). Infection in immunocompetent mice is subclinical, although susceptibility to infection will vary by strain. C57BL/6, DBA, and 129/J mice had a higher parasite burden and depressed antibody responses to intraperitoneal inoculation with E. cuniculi when compared with BALB/c mice (Niederkorn et al. 1981). In immunologically impaired or immunosuppressed mice, however, overt disease may be seen. Naturally infected, immunocompetent mice treated twice weekly with corticosteroids developed abdominal distention and ascites. Microscopic examination of ascites fluid, peripheral blood smears, and brain, liver, spleen, and kidneys tissues revealed clusters of intracellular E. cuniculi organisms after 5 weeks of this immunosuppressive regime (Bismanis 1969). Experimentally infected C 5 7 B L / 6 - H f h l l nu mice develop ascites,
542
or a chronic wasting syndrome characterized by lethargy, anorexia, dehydration, and death within 2 to 4 weeks of parasite inoculation (Didier et al. 1994). Diagnosis of encephalitozoonosis is made by serology, histopathologic examination of tissues, and PCR. Serologic screening is a routine and useful method for monitoring E. cuniculi in immunocompetent mouse colonies. Histologically, parasites may be observed in a variety of tissues, but lesions are most frequently identified in the brain and kidneys. In contrast to the lesions seen in the CNS of endemically E. cuniculi-infected rabbits, those present in mice are nongranulomatous. In infected immunocompetent mice, a diffuse, nonsuppurative meningoencephalitis, and multifocal aggregates of activated microglia, lymphocytes, and macrophages, often with a perivascular distribution, are seen in the CNS. In addition, diffuse lymphocytic interstitial nephritis is observed. In these mice, parasites are scarce and may be difficult to identify. In immunosuppressed mice, E. cuniculi infection tends to be extensive and disseminated. Multiple necrotic foci with abundant intralesional and extralesional parasites have been described in the brain, heart, lungs, liver, spleen, adrenals, kidneys, pancreas, intestine, and the visceral and parietal peritoneal tissues of mice (Didier et al. 1994). Parasites are best observed in tissue sections with special stains (Fig. 21-7A, B). E. cuniculi spores stain positive with a Brown and Brenn's (B&B) tissue Gram stain, and bright red with PAS stain (Wasson and Peper 2000). Mature spore forms pick up stain the best--proliferative forms of the parasite are
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more difficult to identify. Spores are rod-shaped and measure 1.5 x 2.5 ~tm. With B&B, spores often appear to have a clear vacuole at one end. With PAS, this vacuole appears as a deeply red staining granule. Other internal organelles are not visible by light microscopy. On occasion, it may be necessary to discriminate E. cuniculi from T. gondii infection, particularly in mice with CNS lesions. While T. gondii stains PAS positive, it does not stain with B&B. In addition, intracellular clusters of E. cuniculi spores can be distinguished from bradyzoitefilled T. gondii cysts by the presence of a cyst wall in the latter. Cyst walls stain faintly PAS positive and are argyrophilic (Frenkel 1956). Urinalysis for detection of E. cuniculi spores is theoretically possible but is more often used to monitor parasite shedding in experimentally infected mice. Molecular-based methods are available for diagnosis of encephalitozoonosis and may be useful when dealing with immunosuppressed mice and/or inconclusive histopathologic results (Didier et al. 1995). 5. Prevention, Treatment, Control
Because of its subclinical nature and insidious effect on animal-based biomedical research, E. cuniculi-infected colonies should be depopulated and repopulated with clean stock. Alternatively, rederivation can be performed. Although E. cuniculi is not as prevalent in contemporary mouse colonies as in the earlier part of the twentieth century, investigators, mouse biologists, and diagnosticians should remain vigilant.
Fig. 21-7 Encephalitozooncuniculi in the kidneyof a mouse. Clusters of spores are present within intracytoplasmicparasitophorousvacuoles. (A) H&E. (B) B&B Gram stain. A and B, bar = 50 t.tm.
2 1.
PROTOZOA
E. cuniculi infects a variety of cell types and animal species,
and may inadvertently be transmitted through the use of contaminated biologics in mice. Despite the handful of reports suggesting clinical improvement with the use of albendazole in humans with E. cuniculi infection, therapeutic efficacy has not been demonstrated in mice (De Groote et al. 1995b; Fournier et al. 2000; Koudela et al. 1994). Although E. cuniculi infection in humans has been documented, direct zoonotic transmission from animals to humans has not been demonstrated (Glaser et al. 1994). Human infections are likely obtained from water sources or the environment. In laboratory environments, Encephalitozoon spp. have been shown to be inactivated by 2% Lysol, 10% formalin, 70% ethanol, 10% bleach, quaternary ammonium compounds, and by boiling (Santillana-Hayat et al. 2002; Shadduck and Polley 1978). For this reason, steam sterilization of equipment and contaminated animal bedding, as well as routine disinfection of surfaces, should prevent environmental buildup of E. cuniculi spores in animal facilities. 6.
Research Implications
As alluded to in the introduction, interference with research results due to subclinical E. cuniculi infection is well documented in rodents. The inflammatory lesions commonly observed in the CNS of infected mice were often misinterpreted as lesions due to experimental inoculation with rabies virus, herpes simplex, cercopithecine herpesvirus 1, poliomyelitis virus, hepatitis B virus, syphilis, psittacosis, scrub typhus, and toxoplasmosis (reviewed in Wasson and Peper 2000). In addition to infectious disease work, E. cuniculi also interfered with early cancer research. Rats were found to be resistant to tumor development when inoculated with E. cuniculi-infected rat sarcoma cells (Petri 1965). E. cuniculi-infected mice were shown to have significantly reduced tumor growth and prolonged survival rates compared in uninfected controls when inoculated with different strains of solid or ascites-producing murine tumor cell lines (Arison et al. 1966; Niederkorn 1985). Nonspecific resistance to tumor growth in these mice may in part be due to enhanced natural killer cell activity--and associated increase in gamma interferon (INF-y) production--secondary to E. cuniculi infection (Niederkorn et al. 1983). Experimentally, E. cuniculi-infected C57BL/6 mice cleared lymphoma tumor cells from the lungs faster when compared to natural killer cell-deficient beige mice and infected, cyclophosphamide-treated C57BL/6 mice (both of which have reduced levels of INF-y). In addition, infected C57BL/6 mice were significantly more resistant to pulmonary tumor formation with B 16F10 melanoma cells when compared with uninfected, age-matched controls (Niederkorn 1985). The immune response to E. cuniculi infection in mice is now being elucidated from these early observations of enhanced tumor resistance. The importance of INF-~t was reaffirmed when INF-y depleted and INF-y knockout mice were shown to be highly susceptible to E. cuniculi
543
infection when compared to unmanipulated or wild-type controls (Khan and Moretto 1999). INF-y production by natural killer cells and gamma delta T lymphocytes is thought to activate CD8+ cytotoxic T lymphocytes, which in turn results in parasite killing. Evidence for this was shown in CD8 knockout mice, which were more susceptible to E. cuniculi infection compared to CD4 knockout mice and wild-type controls (Khan et al. 1999; Moretto et al. 2000).
ACKNOWLEDGMENTS This work was supported in part by NIH grant AI49735. The author wishes to thank Drs. Stephen Barthold, Nicole Baumgarth, Howard Gelberg, Stefan Tunev, and Mr. Robert Munn, for translation of articles from German and Russian, and for assistance with histopathology antd figures.
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I.
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
II.
Flagellates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
520
A.
Giardia muris . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
520
1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
520
2. Life C y c l e and M o r p h o l o g y . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
520
3. Cell Biology
520
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4. Disease and D i a g n o s i s
B.
...................................
521
6. R e s e a r c h Implications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
521
Spironucleus muris .........................................
523
1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
523
2. Life Cycle and M o r p h o l o g y . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. Cell Biology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
523
4. D i s e a s e and Diagnosis
524
...................................
524 524
1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
524
...................................
5. Treatment, Prevention, Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
III.
524
Tritrichomonas muris .......................................
4. Disease and D i a g n o s i s
E.
524
6. R e s e a r c h Implications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
2. Life C y c l e and M o r p h o l o g y . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. Cell Biology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
D.
521
5. Prevention, Treatment, Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
5. Prevention, Treatment, Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C.
518
524 525 525 525
6. R e s e a r c h Implications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
525
O t h e r Intestinal Flagellates
525
Trypanosoma musculi
..................................
......................................
525
1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
525
2. Life Cycle and M o r p h o l o g y . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. Cell Biology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
526
4. Disease and D i a g n o s i s
526
...................................
526
5. Treatment, Control, R e s e a r c h Implications . . . . . . . . . . . . . . . . . . . Amoebae ....................................................
527
A.
527
Entamoeba muris ..........................................
527
1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
527
2. Life Cycle and M o r p h o l o g y . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
527
3. Cell Biology
527
THE MOUSE IN BIOMEDICAL RESEARCH, 2ND EDITION
...........................................
Copyright 9 2007, 1980, Elsevier Inc. All rights reserved.
517
518
KATHERINE 4. D i s e a s e and D i a g n o s i s
...................................
5. Treatment, Prevention, Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6. R e s e a r c h Implications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV.
528
A.
General Introduction
528
B.
E i m e r i a spp . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
529
1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
529
C.
2. Life C y c l e . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
529
3. Cell Biology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
531
4. Disease and D i a g n o s i s
531
...................................
D.
531
Sarcocystis m u r i s . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
532
1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
532
2. Life Cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
532
3. Cell Biology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
532
4. Disease and D i a g n o s i s
532
...................................
533
Klossiella m u r i s . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
533
1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
533
2. Life C y c l e . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
533
3. Cell Biology
533
...........................................
533 534
6. R e s e a r c h Implications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
534
Toxoplasma gondii .. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
534
1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
534
2. Life Cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
535 535
3. Cell Biology
........................................... ...................................
536
5. Treatment, Prevention, Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
536
6. R e s e a r c h Implications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
536
Cryptosporidium muris
538
.....................................
1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
538
2. Life C y c l e . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
538
3. Cell Biology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
539
4. Disease and D i a g n o s i s
539
...................................
5. Prevention, Treatment, Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
539
6. R e s e a r c h Implications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
540
Microsporidia A.
...................................
5. Prevention, Treatment, Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
4. Disease and D i a g n o s i s
F.
533
6. R e s e a r c h Implications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
4. D i s e a s e and D i a g n o s i s
E.
531
6. R e s e a r c h Implications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
5. Prevention, Treatment, Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
................................................
E n c e p h a l i t o z o o n cuniculi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
540 540
2. Life C y c l e . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
540 541
3. Cell B i o l o g y
541
1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ...........................................
4. Disease and D i a g n o s i s
I.
528 528
5. Treatment, Prevention, Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
V.
527
Apicomplexans ............................................... .......................................
...................................
541
5. Prevention, Treatment, Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
542
6. R e s e a r c h Implications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
543
Acknowledgments ..................................................
543
References
543
.....................................................
INTRODUCTION
The Kingdom Protozoa contains a diverse collection of single-celled eukaryotic organisms that in many ways bridge the gap between the plant and animal worlds. Protozoa may be free living or parasitic, may be capable of photosynthesis, or may be thriving in a microaerophilic environment. Others can be considered extremophiles, living in ice, hydrothermal vents, or sulfur-emitting fumaroles. A relatively small number
WASSON
of these organisms are parasitic for mammals. Although there is no universal agreement on the correct terminology for these organisms, the parasitic protozoa of mammals fall into one of four phyla: the amoebae (Sarcodina), the flagellates (Mastigophora), the apicomplexans (Sporozoa, or the coccidia), and the ciliates (Ciliophora). Of these, mice can be naturally infected with members from the first three phyla; there are no known reports of murine-specific ciliates. A fifth phylum, the Microspora (which includes Encephalitozoon cuniculi), is now considered a member of the Kingdom Fungi (see Table 21-1 for
21.
519
PROTOZOA TABLE 21-1
COMPARISON OF MURINE PROTOZOANS a Phylum
Genus and species
Diagnostic stage(s)
Diagnostic test(s)
Size ( ~ t m )
Description
Amoeba
Entamoeba muris
Cyst
Fecal float, smear
9-19 diameter
Trophozoite
Fecal smear
12-30 diameter
Cryptosporidium spp.
Oocyst
Fecal float
5• 7
Eimeria falciformis
Oocyst
Fecal float
14-27 • 11-24
E. ferrisi
Oocyst
Fecal float
12-22 • 11-18
E. papillata
Oocyst
Fecal float
18-26 • 16-24
E. vermiformis
Oocyst
Fecal float
18-26 • 15-21
Klossiella muris
Sporocyst
Urinary centrifugation
16 x 13
Sarcocystis muris Toxoplasma gondii
Bradyzoite-filled cyst Cyst (intermediate host) Oocyst (Cat)
Histology Histology
4-6 • 14-16 Variable
Fecal float, smear
11-14 • 9-11
Tachyzoite
Histology
2-3 • 6-7
Bradyzoite
Histology
Giardia muris
Cyst Trophozoite
Fecal float, smear Fecal smear
15 • 17 7-13 • 5-10
SpironucIeus muris
Cyst
Fecal float
7• 4
Trophozoite
Fecal smear
10-15 x 3-4
Tritrichomonas muris
Trophozoite
Fecal smear
16-19 • 7-9
Trypanosoma musculi Encephalitozoon cuniculi
Trypomastigote Spore
Blood smear Histology
2-3 • 16-34 1.5 • 2.5
Round; 8 nuclei; lumen of cecum and ascending colon Pleomorphic, 1 nucleus; lumen of cecum and ascending colon Round to ellipsoid; gastrointestinal mucosa; intracellular but extracytoplasmic Round to ellipsoid; smooth; clear to light brown; crypt epithelium of cecum and colon Round; smooth; clear to light brown; villus epithelium of cecum and colon Round to ellipsoid; papillated; yellowish brown; villus epithelium of distal small intestine Ellipsoidal; pitted; yellowish brown; crypt epithelium of distal small intestine Round; glomerular endothelium and renal tubule epithelium Banana-shaped; myocytes Bradyzoites within cysts 5-8 • 1-2; prominent cyst wall; CNS, muscle, other tissues Round to subspherical; small intestinal epithelium Crescent shaped; within variety of nucleated cells Fusiform shaped; found within tissue cysts in variety of tissues Ellipsoid; 4 nuclei; lumen of small intestine Bilaterally symmetrical, pear-shaped; two nuclei; 4 pairs of flagella; closely associated with small intestinal villus brush border; "falling leaf' motility Ellipsoid; 2 nuclei; "Easter egg"; lumen of small intestinal Ellipsoid, tapered; found in mucus layer of small intestinal villus and crypts; "zig-zag" motility Pear-shaped; lumen of cecum and colon; "rolling" motility Elongated, vermiform shaped Rod-shaped; free or within parasitophorous vacuoles in brain, renal tubules
Apicomplexans
Flagellates
Microsporidia
aSee text for references.
a summary). Because of E. cuniculi's historical classification as a protozoan, and because of its importance as a pathogen in laboratory mice, this organism will be included in this discussion of murine protozoa. Not all of these organisms are pathogenic for mice. Indeed, the flagellate Tritrichomonas muris and the amoeba Entamoeba muris can be considered commensals of the murine large intestine. Others, for example, the flagellate Giardia muris and the apicomplexan Cryptosporidium muris, tend to cause disease in immunologically na'fve, neonatal mice. Susceptibility to these organisms tends to have a mouse
strain and gender predisposition, and it is expected that genetically engineered mice will vary in their susceptibility to disease as well. Lastly, several of the parasitic protozoa of mammals were first recognized and described in mice, and mice continue to be instrumental in understanding the pathology, immune response to, clearance, and treatment of these infections. Although parasitic infections are rare in well-managed facilities utilizing rederived and barrier-maintained mice, surveillance for and familiarization with these organisms in mouse colonies are still warranted.
520
KATHERINE
II.
A.
FLAGELLATES
WASSON
G. lamblia, or unspeciated Giardia (Franjola et al. 1995; Sogayar and Yoshida 1995).
Giard& muris 2.
1. Introduction
Life Cycle and Morphology
Genus
Species
Primary host
Giardia
muris agilis ardeae lamblia (duodenalis, intestinalis) microti psittaci
Rodents Amphibians Herons Mammals including humans
G. muris, as with all Giardia species, has a simple and direct life cycle. Infection is acquired through the ingestion of environmentally resistant cysts. Experimental infections indicate that the minimal infectious dose of G. muris in outbred Swiss or athymic nude mice is 10 spores (range, 5 to 20), and the prepatent period prior to fecal shedding is 8 days (range, 5-14; Roberts-Thomas et al. 1976; Stachan and Kunstyr 1983). Cysts undergo excystation upon exposure to gastric pH and secretions from the upper gastrointestinal tract. Upon excystation, two trophozoites are released into the small intestine. Trophozoites are bilaterally symmetrical and pear-shaped, possess four pairs of flagella, and measure 7-13 x 5-10 ~tm. Two nuclei and the median body often give trophozoites a "ghost" face appearance under light microscopy. Trophozoites are found in the upper small intestine where they attach to intestinal epithelial cells by means of a ventral "adhesive" disk. Trophozoites replicate by binary fission to produce two identical daughter trophozoites. As trophozoites are sloughed into the intestinal lumen toward the large bowel, a certain percentage will encyst and be excreted as cysts into the environment. The length of time that cysts are shed in the feces is dependent on the dose of the original inoculum, host immune status, mouse strain, and possibly gender (Belosevic and Faubert 1983; Belosevic et al. 1984; Daniels and Belosevic 1995; Heyworth 1988). Cysts are ellipsoid in shape, possess four nuclei (two trophozoites), and measure 15 • 17 ~tm. To provide organisms for in vitro studies, G. muris trophozoites can be collected from the duodenum and jejunum of infected mice (or other infected hosts) and isolated in axenic culture media (Tillotson et al. 1991). Trophozoites can also be generated by harvesting cysts from fecal pellets and inducing them to excyst in the presence of trypsin and low pH (Schaefer et al. 1984).
Voles, muskrats Parrots
3.
muris andersoni baileyi canis felis galli hominis meleagridis molnari parvum serpentis saurophilium wrairi
Rodents Cattle Chickens, turkeys Dogs Cats Finches, chickens Primates including humans Turkeys Fish Cattle, sheep, goats Snakes, lizards Lizards Guinea pigs
Mice are natural hosts to Giardia muris and can be experimentally infected with G. lamblia. G. lamblia (also referred to as G. intestinalis, or G. duodenalis; Thompson et al. 2000) is a waterborne pathogen and an important cause of human diarrhea on a global basis (Adam 2001). Although the classification and nomenclature of Giardia is still evolving, it is generally agreed that six distinct species exist and that G. lamblia can be further subdivided into two major genotypes (Table 21-2; Adam 2001). The virulence and zoonotic potential between species and genotypes varies. Among the flagellates found in the intestines of mice, only G. muris is considered a primary pathogen of mice (Sebesteny 1969). Giardiasis in laboratory-reared mice should be nonexistent under proper rederivation, husbandry, and management standards. Colonies of G. muris-infected mice still exist, however. G. muris was diagnosed in 6.9% of "white" laboratory mice, and co-infection with other murine intestinal protists was noted (Jalili et al. 1995). Giardia infection is also common in wild rodent populations, which can serve as potential sources of infection for laboratory mice. Recent field studies of wild rodents (including Mus musculus and Rattus rattus) indicated that 14.3 to 100% of the various populations harbored G. muris,
TABLE 21-2
VALIDATED GIARDIA AND CRYPTOSPORIDIUM SPECIESb
Cryptosporidium
bAdapted from Adam (2001) and Xiao et al (2004).
Cell Biology
Giardia spp. possess two membrane-bound nuclei, a complex cytoskeleton, and a Golgi-like membrane system. However, they lack several features found in other eukaryotes, including nucleoli, mitochondria, enzymes involved in oxidative phosphorylation, and most enzymes involved in amino acid and nucleoside synthesis. As a result, Giardia are anaerobic and must forage for amino acids, purines, and pyrimidines from the intestinal milieu. The nuclei of Giardia are unique in that both are transcriptionally active and replicate at the same time. A second unique feature of Giardia is the ventral disk. This cytoskeletal structure allows the trophozoite to attach to intestinal epithelial cells of the duodenum and jejunum, preventing premature exit from the host. It is composed of
21.
several cytoskeletal proteins, including [3-tubulins (Parsons 1995). The median body, located on the midline, is another unique component of the trophozoite's cytoskeleton system. This structure is thought to be the assembly site for microtubule bundles to be incorporated into the ventral disk (Meng et al. 1996). The morphology of the median body can be used to distinguish between some of the Giardia species: G. lamblia has one or two transverse median bodies that are shaped like hammer claws; G. muris has one small, rounded median body (Adam 2001). Giardia lack mitochondria and the ability to generate energy through aerobic metabolism. However, identification of mitochondria-like heat-shock protein genes and mitochondrial remnant organelles ("mitosomes") suggests that Giardia may have lost this organelle during the course of adapting to a parasitic lifestyle (Roger et al. 1998; Tovar et al. 2003). Lastly, Giardia trophozoites are capable of antigenic variation by expressing a family of cysteine-rich, immunodominant proteins across their surfaces known as variable surface proteins (Aggarwal et al. 1988). The ability to undergo antigenic variation is likely a means of evading the host immune response and may give the organisms a survival advantage in differing intestinal microenvironments (Adam 2001). 4.
521
PROTOZOA
Disease and Diagnosis
G. muris infections generally do not cause clinical signs in immunocompetent mice. Immunocompromised mice may exhibit weight loss and failure to thrive. These clinical signs are also observed in mice infected as weanlings, presumably due to a less mature immune system at this age (Buret et al. 1990). Unlike giardial infection in other mammals, diarrhea is not a clinical feature of disease in mice (Eckmann and Gillin 2001). Giardia clearance, as well as acquired immunity to the parasite, is primarily dependent on IgA production (Langford et al. 2002). Cellular immune responses and innate immunity factors such as defensins and host microflora also contribute to disease resistance and clearance (Singer and Nash 1999, 2000). G. muris can be diagnosed by identifying the characteristic trophozoites or cysts in feces by light microscopy. A "falling leaf' motility may be observed in fresh, unstained wet-mount samples taken from the duodenum or jejunum. Wright or Giemsa stains can be used to enhance the "ghost" face appearance of trophozoites. Cyst stages can be identified on fecal flotation or smears, and differentiated from other cysts or oocysts by their size (Table 21-1). Four nuclei may be seen on trichrome- or iodine-stained preparations of cyst forms. Owing to irregular shedding of cysts, several fecal examinations may need to be performed prior to ruling out an antemortem diagnosis of giardiasis. Histologically, trophozoites are found attached to the brush border of epithelial cells of the duodenum and jejunum. Trophozoites are lightly eosinophilic, crescentshaped on longitudinal view, and noninvasive. Careful examination under oil immersion may reveal the presence of
the ventral disk (Fig. 21-1A). There may be mild reduction in the villus:crypt ratio, and mild to moderate lymphocyte infiltration in the underlying mucosa and lamina propria (MacDonald and Ferguson 1978). Organisms need to be differentiated from Spironucleus muris, which tend to localize in the distal small intestine and are often found packed in intestinal crypts (Brett and Cox 1982, Owen et al. 1979). Enzyme-linked immunosorbent assays, flourescence antibody assays, and molecular-based methods are also available for diagnosis of giardiasis (Garcia et al. 1992; Nash et al. 1987; Sedinova et al. 2003). These tests are used for the detection of human giardial infection, and several are available in kit format. Use of these or comparable diagnostic kits for screening rodent colonies has not been reported. 5.
Prevention, Treatment, Control
G. muris is readily transmitted between mice and other rodent species (Belosevic et al. 1986a; Kunstyr et al. 1992; Saxe 1954). Therefore, rederivation is the treatment of choice for eliminating Giardia spp. from rodent colonies. Metronidazole and albendazole are the drugs of choice for treating humans infected with G. lamblia (Gardner and Hill 2001). The efficacy of these compounds may not be complete in mice (Bemrick 1963; Oxberry et al. 1994). Treatment with oral metronidazole resulted in only a 58.3% cure rate in mice naturally infected with G. muris, based on histologic examination of the small intestine for the presence of trophozoites (Cruz et al. 1997). Cyst forms shed into the environment are resistant to chlorine and ozone exposure, but can be inactivated by autoclaving of bedding and caging material (Lane and Lloyd 2002).
6.
Research Implications
Subtle alterations in intestinal epithelial cell kinetics, brush border enzyme composition, and cytokine production have been documented in immunocompetent mice with G. muris infection. Although villus height remained the same, a higher turnover rate of intestinal epithelial cells was observed in mice naturally co-infected with G. muris and Spironucleus muris when compared to control mice (MacDonald and Ferguson 1978). Significant decreases in the brush border enzymes lactase, sucrase, trehalase, and maltase were documented in C57BL/6 mice by day 10 of infection with G. muris (Daniels and Belosevic 1992). Reduced production of the epithelial cytokine IL-6 was reported in CD-1 mice (Scott et al. 2000). In addition, an increased epithelial lymphocyte infiltration of the small intestine has been observed with resolving Giardia infection (Brett and Cox 1982). All of these factors make endemically infected mice unsuitable for research involving intestinal physiology. A decreased T cell response has also been reported in mice experimentally infected with G. muris (Brett 1983). This was documented by a decreased ability of infected mice to mount
522
KATHERINE
WASSON
Fig. 21-1 Intestinal flagellates of mice. (A) Two Giardia spp. trophozoites in the upper small intestine of a mouse. Note the pear-shaped trophozoite with two nuclei and posterior flagella, and its close association to the intestinal brush border; the second trophozoite is oriented longitudinally, demonstrating the cup shape of the ventral adhesive disk. (B) Spironucleus muris trophozoites in the distal small intestine of a mouse; trophozoites are small and tapered, and associated with the mucus layer. (C) Tritrichomonas muris and (D) T. minuta in the ceca of mice; note the foamy cytoplasm and larger size of T. muris compared with T. minuta. H&E; A through D, bar = 50 ~tm.
2 1.
523
PROTOZOA
an immune response to sheep red blood cells. T cell responses returned to normal with resolution of infection (Brett 1983). Not surprisingly, exogenous administration of the immunosuppressive drugs cortisone and cyclosporin A resulted in increased cyst shedding and prolonged time to clearance of infection (Nair et al. 1981; Belosevic et al. 1986b). In the case of cortisone, recrudescence of subclinical infection could also be induced (Nair et al. 1981). These results suggest that administration of immunosuppressive agents to mice on experiment may exacerbate occult G. muris infection. A murine model of giardiasis was first described in 1976, and although it does not replicate all the features of human giardiasis, mice have been valuable in understanding the pathophysiology of this disease (Roberts-Thomas et al. 1976). Choice of mouse to use for infection studies should take into account strain, major histocompatibility complex haplotype, and perhaps gender. Early work demonstrated that C57BL/6, C57BL/10, and DBA/2 strains have a longer prepatent period, lower fecal cyst output, and faster resolution of experimental G. muris infection than A/J, BALB/c, or C3H/He strains of mice (Belosevic et al. 1984; Daniels and Belosevic 1992). The differences in the time course of infection were hypothesized to be due to differences in production of IgG2a and gamma interferon (7-INF) inherent in the different mouse strains. A more virulent giardial infection could be induced in C57BL/10 mice by the administration of anti-y-INF antibodies (Venkatesan et al. 1996). No gender differences were observed in the number of cysts shed in feces in BALB/c mice during acute infection with G. muris (Heyworth 1988); however, female C57BL/6 mice cleared G. muris infection by 18 days post-inoculation, while male mice of the same strain continued to shed cysts in the feces beyond 60 days (Daniels and Belosevic 1995). Murine haplotype also plays a role in clearing infection. BALB mice of the H-2b haplotype shed G. muris cysts for a longer period of time than BALB mice of the H-2d or H-2k haplotypes (Venkatesan et al. 1993). The commercial source of mice should also be considered. Isogenic mice from two different vendors were found to have different susceptibilities to experimental G. lamblia infection (Singer and Nash 1999). The inherent resistance in mice from one vendor could be overcome when the intestinal flora was altered with the antibiotic neomycin. Resistance in these mice could also be overcome when housed with susceptible mice from the second vendor (Singer and Nash 1999). These results suggest that differences in the endogenous microflora and fauna can affect both experimental and natural Giardia infection.
B. 1.
Spironucleus muris
Introduction
Originally called Hexamita muris, Spironucleus muris has been renamed to differentiate the exclusively parasitic ("Spironucleus")
from the usually free-living ("Hexamita") members of this genus (Brugerolle et al. 1980). Recent surveys of laboratory and wild mouse populations suggest that the prevalence of S. muris ranges from 4.1 to 38.6% (Franjola et al. 1995; Jalili et al. 1995). Spironucleus muris is probably represented by subspecies with differing host preferences and infectivity. In several separate transfaunation studies, isolates of S. muris from mice and golden Syrian hamsters were infectious for the reciprocal host; isolates obtained from a rat and European hamster were not infectious for mice; and in a third study, isolates from a mouse and rat were infectious for hamsters and rats (Saxe 1954; Schagemann et al. 1990; Sunstyr et al. 1993). Regardless of host specificity, these organisms are likely commensals in their respective rodent hosts (Baker et al. 1998; Sebesteny 1969). Several clinical reports from the 1970s describe "outbreaks" of spironucleosis in laboratory mice, with mortality rates ranging from 20 to 50%. Clinical signs included chronic wasting in athymic nude and thymectomized mice (Boorman et al. 1973); or depression, distended abdomens, "sticky stools," and nonbloody, catarrhal enteritis in weanling age mice (Flatt et al. 1978; Wagner et al. 1974). Large numbers of Spironucleus muris trophozoites, and occasionally G. muris cysts, were observed at necropsy. Thirty years later, descriptions of these "outbreaks" are reminiscent of disease seen with enteric mouse hepatitis virus (MHV), transmissible murine colonic hyperplasia (Citrobacter rodentium), Tyzzer's disease (Clostridium piliforme), and/or salmonellosis rather than primary infection with S. muris. Today, large numbers of S. muris organisms are frequently observed secondary to underlying disease, often MHV infection (Percy and Barthold 2001). A diagnosis of spironucleosis should prompt the clinician to search mouse colonies for an underlying infectious agent or disease condition. 2.
Life Cycle and Morphology
S. muris has a simple and direct life cycle. Infection is initiated with the ingestion of cysts. A single trophozoite is released from S. muris upon excystation in the upper gastrointestinal tract (Schagemann et al. 1990). Trophozoites replicate by longitudinal binary fission. Trophozoites have a slender, tapered body measuring 10-15 x 3-4 ~tm. They possess four pairs of flagella and two nuclei, but lack the specialized cytoskeletal features seen in Giardia spp. (Brugerolle et al. 1980). Trophozoites encyst as they move down the intestinal tract. Cysts measure 7 • 4 ~tm and resemble "Easter eggs" due to the presence of flagella under the cyst membrane, giving the cysts a banded appearance (Kunstyr !977; Kunstyr et al. 1977). The minimal infective dose is one cyst, and the prepatent period has been reported between 2 and 8 days for mice (Kunstyr 1977; Stachan and Kunstyr 1983). In vitro culture and manipulation of S. muris have not been reported; organisms are passaged in flagellate-free mice for experimental manipulation (Schagemann et al. 1990).
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KATHERINE
3. Cell Biology Relatively little is known regarding the cellular and molecular biology of S. muris. Like Giardia, they are early eukaryotic organisms, possessing a membrane-bound nucleus, but lacking mitochondria and, presumably, the enzymes involved in aerobic metabolism (Brugerolle et al. 1980). S. muris lacks the ventral disk and median body observed in Giardia (Januschka et al. 1988). Trophozoites feed on luminal flora, as evidenced by the presence of bacteria in digestive vacuoles seen on electron microscopy (Brugerolle et al. 1980). An inverse ratio of Giardia and Spironucleus trophozoites has been observed in naturally infected mice, suggesting that these organisms compete for similar resources in the host (Sebesteny 1969). 4.
Disease and Diagnosis
Early descriptions of disease due to spironucleosis must be interpreted with caution, as S. muris is likely a "facultative pathogen" in mice infected with other disease agents (Kunstyr et al. 1977). In contemporary mouse colonies, heavy S. muris burdens are associated with enteric mouse hepatitis virus infection (Percy and Barthold 2001). Spironucleosis has also been observed in otherwise pathogen-free, genetically engineered mice with underlying immune alterations. Experimental S. muris infection of athymic nude mice resulted in abdominal distention, failure to thrive, and death after 2 to 3 months of infection (Kunstyr et al. 1977). A more recent experimental report noted a lack of clinical disease and intestinal pathology in a variety of inbred strains of mice orally inoculated with S. muris cysts (Baker et al. 1998). Diagnosis of S. muris depends on observing trophozoites or cysts in fecal smears or histopathology. Trophozoites have a fast, zigzag movement on fresh wet-mount samples taken from the distal small intestine and colon (Sebesteny 1969). The nuclei are best visualized with Giemsa or hematoxylin stain (Sebesteny 1969). Cysts are found in the large intestine and feces, and can be differentiated from Giardia cysts by their smaller size and "Easter egg" appearance (Kunstyr et al. 1977). On histologic sections, organisms are located in the distal small intestine and are usually found in the mucus layer or packed within intestinal crypts (Wagner et al. 1974; Brugerolle et al. 1980). Invasion of the intestinal mucosa and lamina propria by organisms has also been reported (Flatt et al. 1978). S. muris trophozoites appear as smudgy, lightly eosinophilic structures with hematoxylin-eosin stain (Fig. 21-1B). Under 100X oil immersion, they appear as plump banana-shaped organisms, with a single mid-body nucleus visible. Diagnostic immunologic or molecular techniques have not been reported for S. muris. 5.
WASSON
dimetronidazole, metronidazole, or tinidazole was ineffective in eliminating cyst shedding in mice (Kunstyr et al. 1977; Sebesteny 1969). Albendazole, a microtubule inhibitor with some efficacy against Giardia infection, had minimal effect on the ultrastructure of S. muris (Oxberry et al. 1994). Cysts are susceptible to several common disinfectants, fixatives (70% ethanol, household bleach, aldehyde-based compounds), and temperatures above 45~ suggesting that standard laboratory animal husbandry and management procedures will control cysts in the environment (Kunstyr and Ammerpohl 1978). 6. ResearchImplications S. muris infection complicates research with experimental intestinal flagellate infection (Owen et al. 1979). In that report, mice colonized with S. muris were less susceptible to experimental G. muris infection. Alterations in intestinal epithelial cell kinetics were observed in mice co-infected with S. muris and G. muris (MacDonald and Ferguson 1978). A decreased villus to crypt ratio, and decreased T cell-dependent immune response to sheep red blood cells, have been observed in S. muris-infected mice (Brett 1983; Brett and Cox 1982).
C.
Tritrichomonas muris
1. Introduction Tritrichomonas muris is a nonpathogenic flagellate of mice, rats, hamsters, and other rodents. Early literature refers to this organism as Trichomonas muris, Trichomonas cricetus, or Tritrichomonas cricetus. However, these were determined to be synonyms for morphologically identical organisms found in a variety of laboratory and wild rodents (Daniel et al. 1971; Honigberg 1963). An exhaustive transfaunation experiment demonstrated the ease with which several trichomonads (including T. muris and Pentatrichomonas hominis) could be established in and transmitted between mice, rats, and hamsters (Saxe 1954). T. muris is related to the more pathogenic Trichomonas vaginalis and Tritrichomonas foetus of humans and cattle, respectively. These latter organisms are sexually transmitted protists that live in the genitourinary tracts of their hosts, causing reproductive disorders (BonDurant 1997; Petrin et al. 1998). T. muris resides in the cecum and colon of rodents, where it is considered a component of the normal fauna (Sebesteny 1969). Recent surveys of laboratory and wild colonies of mice demonstrate a 29.6 to 47.4% prevalence of the organism in the large intestines (Franjola et al. 1995; Stachan and Kunstyr 1983). Although little is known of T. muris, much of its life cycle and cell biology can be inferred from work done on more pathogenic family members.
Prevention, Treatment, Control
S. muris is readily transmitted between rodents (Saxe 1954).
Rederivation and fostering on flagellate-free dams will eliminate S. muris from mouse colonies. Chemotherapy using
2. LifeCycle and Morphology T. muris has a simple and direct life cycle, and exists as motile trophozoites within the host. Pseudocyst forms of
2 1.
T. vaginalis and other trichomonads have been observed and are thought to represent degenerating trophozoites responding to unfavorable environmental conditions (Honigberg 1963; Petrin et al. 1998). The minimal infectious dose of T. muris "pseudocysts" for mice is 5, and the prepatent period is 10 days (Stachan and Kunstyr 1983). Trophozoites are pear- or teardropshaped, measuring 16-19 • 7-9 ~tm (Selukaite 1977; Fig. 21-1C). Trophozoites replicate by binary fission. Trophozoites reside in the cecum and colon but have been reported in the stomach and small intestines, likely as a result of recent ingestion (Koyama et al. 1987; Selukaite 1977). Newborn hamsters, and probably other rodent pups as well, are colonized by T. muris within a week after birth (Mattern and Daniel 1908). Cell-free cultivation of T. muris has not been reported; trophozoites have been harvested from mono-infected rodents for experimental manipulation (Saxe 1954).
3.
Cell Biology
All trichomonads possess a single membrane-bound nucleus, three anterior flagella, and a fourth posterior flagellum that forms the "backbone" of the undulating membrane. The undulating membrane runs partway down the length of T. muris and can be visualized in less motile trophozoites under light microscopy (Osada 1962). The axostyle is the cytoskeletal structure that appears to give the organism a "backbone." It originates at the nucleus and tapers to a tail-like appendage at the distal end of the parasite (Daniel et al. 1971). In T. vaginalis, the axostyle is thought to be an attachment organ on vaginal epithelial cells (Petrin et al. 1998). On electron microscopy, a comb-like structure called the costa can be seen anchoring the base of the undulating membrane (Daniel et al. 1971). The parabasal body is a collection of flattened cisterns similar in structure to the Golgi complex in higher eukaryotes. Trichomonads lack mitosomes but possess double-membrane bound granules that metabolize carbohydrates and produce ATP and hydrogen, and are referred to as hydrogenosomes (Petrin et al. 1998). 4.
525
PROTOZOA
Disease and Diagnosis
No disease has been attributed to T. muris in rodents. Impressive numbers of trophozoites can be seen in the cecal lumen of some mice in the absence of disease (Fig. 21-1C). Diagnosis can be made by examining fresh wet mounts from the cecum and colon by light microscopy. Organisms can be identified by their "rolling" or "quivering" movement (Petrin et al. 1998). Staining wet mounts with Giemsa or iodine enhances the appearance of the nucleus and undulating membrane.
indicator of breach in barrier maintenance of otherwise pathogen-free mice. Chemotherapeutic elimination of T. muris has not been reported. The lack of T. muris cyst forms implies that standard husbandry and management practices will eliminate organisms from the environment.
6.
Interference with research results due to the presence of T. muris in mice has not been reported. Due to the health significance of T. vaginalis in the human population, there has been interest in using the laboratory mouse as a model for this sexually transmitted disease. Patent, long-term infection requires pretreatment of female mice with estrogen and intravaginal doses of Lactobacillus spp. prior to the introduction of T. vaginalis organisms (McGrory and Garber 1992). Using this model, investigators have demonstrated that IgA antibodies protect against severe infection with T. vaginalis and that T-lymphocytes are important in parasite clearance in infected women (Paintlia et al. 2002).
D.
Rederived and barrier-maintained mice are free of T. muris. The ease with which T. muris can be transmitted between mice and other rodents suggests that trichomonads can be used as an
Other Intestinal Flagellates
In addition to the above-mentioned protists, several other nonpathogenic flagellates have been identified in the intestines of the laboratory mouse. Most of these can be identified according to their morphology and size. Trichomonas minuta (4-9 x 2-5 l.tm) and Trichomonas wenyoni (4-16 x 2.5-6 l.tm) are smaller in size and have a less vacuolated or "foamy" cytoplasm compared to T. muris (Levine, 1973b). T. minuta has a more prominent and eosinophilic axostyle than T. muris (Fig. 21-1D). Chilomastix bettencourti is a cyst-forming flagellate found in the cecum of mice, rats, and hamsters. Trophozoites are asymmetrically piriform and measure 8-20 x 7-8 l.tm; cysts are lemon-shaped and measure 6.5-9 x 5.5-7 l.tm (Nie 1948). They possess three anterior flagella and a short, posteriorly directed flagellum. A distinctive, pouch-like cytostome can be seen in both forms, and is thought to be a feeding organelle (Levine 1973a). Octomitus pulcher is a bilaterally symmetrical flagellate that morphologically resembles Giardia spp. They measure 6-10 x 3-7 ~tm and possess six anterior and two posterior flagella, as well as two anteriorly located nuclei. O. pulcher is found in the cecum of mice, rats, hamsters, and other wild rodents (Gabel 1954; Levine 1973a).
E. 1.
5. Treatment, Prevention, Control
Research Implications
Trypanosoma musculi
Introduction
Trypanosoma musculi is a nonpathogenic hemoflagellate parasite of wild mice living in the Mediterranean basin, West Africa, and Central America. T. musculi is host-specific for Mus musculus (Krampitz 1969). First described in 1909, this
526
KATHERINE
WASSON
organism is also referred to as T. duttoni in the older literature (Kendall 1906). A relatively recent survey identified T. musculi in 3.8% of wild house mice in the Arabian peninsula (Molan and Hussein 1988). There are no published reports of natural infection with this parasite in laboratory-raised mice.
recta of the kidneys, and suggested that mice can be persistently infected for life (Viens et al. 1972). Subsequent work has shown that capillaries of the vasa recta provide nutrients and an "immunologically" privileged site for T. musculi trypomastigotes (Monroy and Dusanic 1997).
2.
3.
Life Cycle and Morphology
The life cycle and morphology of T. musculi is similar to that of other African trypanosomes. T. musculi is transmitted by fleas, including the Oriental rat flea, the Northern rat flea, and the mouse flea (Xenopsylla cheopis, Nosopsyllus spp., and Leptosylla segnis, respectively). Mice become infected by ingesting infected fleas or flea feces containing trypanosomes. Mice become parasitemic, during which time trypomastigotes replicate by multiple fission, and can be identified in peripheral blood smears. Trypomastigotes have an elongated, vermiform shape and measure 2 to 3 ktm in width and 10 to 34 ~m in length (Taliaferro and Pavlinova 1936; Fig. 21-2). Parasitemia lasts 2 to 3 weeks, after which time organisms are difficult to identify in blood smears. The life cycle is perpetuated when fleas consume a blood meal from parasitemic mice. The peak parasite burden varies considerably in immunocompetent strains of mice (Derothe et al. 1999). Mice develop immunity and are resistant to reinfection with T. musculi. However, Viens et al. demonstrated that a small percentage of parasites persisted (and continued replicating) in the vasa
Trypanosomes possess a membrane-bound nucleus and a single flagellum. Trypanosomes also possess a single mitochondrion referred to as a kinetoplast. In addition to performing metabolic functions as in other eukaryotic organisms, this organelle has a unique mitochondrial genome organization and function (McFadden 2003). Indeed, several unique RNA processing mechanisms were first described in the kinetoplast of trypanosomes (Gott and Nilsen 2003). These include transsplicing and RNA editing. The DNA in the kinetoplast is organized in 20 to 50 "maxicircle" DNA segments and 5000 to 10,000 small circular DNA segments. The large content of DNA in this organelle contributes to its intense staining. Instead of the conventional linear method of transferring genomic information from DNA to RNA (transcription), and then to protein (translation), various transcripts of RNA from different areas of the kinetoplast genome can be ligated together prior to being translated (trans-splicing). The ability to perform trans-splicing is the mechanism behind the phenomenon of antigenic variation in African trypanosomes, a major means by which parasites evade the host immune system. In addition to modifying RNA by ligating pieces together, sections of transcribed RNA can be altered by the insertion or deletion of uridine residues in order to create messenger RNA that will code for functional proteins (RNA editing). Once thought to be unique to the kinetoplastid parasites, trans-splicing has since been identified in parasitic and free-living nematodes, and RNA-editing has been described in plant mitochondria and chloroplasts. 4.
Fig. 21-2 Trypanosoma sp. in a peripheral blood smear from a cow. Note the kinetoplast, undulating membrane, and single flagellum. Similar features would be seen in T. musculi from mice. H&E; bar - 50 gm (Image courtesy of H. Gelberg).
Cell Biology
Disease and Diagnosis
T. musculi is relatively nonpathogenic for immunocompetent mice. Mice develop mild anemia, splenomegaly, hepatomegaly, and lymph node hyperplasia that resolve after one month (Hirokawa et al. 1981). Mice experimentally infected between the fourth and fifteenth day of gestation, however, develop fatal parasitemia, with large numbers of replicating trypomastigotes present in the maternal vessels of the placenta (Krampitz 1969). Athymic nude mice develop persistent infection, while splenectomized mice develop fatal parasitemia, when experimentally inoculated with T. musculi trypomastigotes (Rank et al. 1977; Taliaferro and Pavlinova 1936). The trypomastigote forms of T. musculi are morphologically similar to those of other mammalian trypanosomes. When stained with Giemsa, they have a large, red, centrally placed nucleus and a smaller, posteriorly located kinetoplast. The cytoplasm
2 1.
527
PROTOZOA
stains blue. An undulating membrane can be seen by light microscopy running the length of the trypomastigote. The membrane continues past the main body of the parasite as a free, anteriorly located flagellum. Infection is best diagnosed on peripheral blood smears. Trypomastigotes may also be observed in histologic sections of spleen, liver, and kidneys (Hirokawa et al. 1981). 5. Treatment, Control, Research Implications
Because of the self-limiting nature of T. musculi infection, treatment of infected mice has not been reported. Experimental infection in laboratory mice has been proposed as an in vivo model for screening chemotherapeutics against Chagas' disease in humans (Jennings and Gray 1982). However, laboratory mice can be experimentally infected with the etiologic agents of both human American and African trypanosomiasis (T. cruzi and T. brucei spp., respectively). Infection in mice with these organisms mimics many of the pathologic and immunologic features of chronic human infection (Kennedy 1999; Marinho et al. 2004). Consequently, T. musculi infection in mice is rarely reported as a model for human trypanosomiasis. In areas where T. musculi is endemic in wild mouse populations, prevention of infection in the laboratory setting involves appropriate vermin and insect control. There are no published reports of natural T. musculi infection confounding experimental data obtained from mice.
III.
A.
AMOEBAE
Entamoeba muris
1. Introduction The genus Entamoeba is composed of a diverse group of parasitic and free-living, single-celled organisms with an amoeboid mode of locomotion (Silberman et al. 1999). Most Entamoeba are nonpathogenic. Entamoeba muris is the only amoeba identified in laboratory mice and is considered a commensal inhabitant of the cecum. The older literature refers to this organism as Amoeba muris, Entamoeba muris decumani, Councilmania muris, Councilmania decumani, Endamoeba ratti, or Entamoeba coli var ratti. Based on
morphology, these terms are now considered synonyms for E. muris (Neal 1950). Surveys of laboratory and wild mouse populations demonstrate a prevalence of E. muris between 5 and 55% (Franjola et al. 1995; Jalili et al. 1995; Livingston 2004; Pruss 1960). Organisms morphologically identical to E. muris are also identified in laboratory and wild rats, and hamsters (Pruss 1960). E. muris is related to the more pathogenic E. histolytica and E. invadens of humans and reptiles, respectively.
2. LifeCycle and Morphology Entamoeba spp. have a simple and direct life cycle. Trophozoites are found in the cecum and anterior colon, where they feed on bacteria, protozoa, and other luminal material (Lin 1971). They are pleomorphic on wet mounts, round or ovoid in histologic sections, and possess a single nucleus (Neal 1950). The pseudopod, an ectoplasmic extension distinct from the endoplasm (cytoplasm), and a trailing uropod may be observed in wet-mount preparations and account for the amoeboid motion of these organisms. They lack cilia, flagella, or other organized cytoskeletal structures. The cytoplasm contains vacuoles of varying size. Trophozoites replicate by binary fission. Encystation occurs in the cecum, with mature cysts possessing eight nuclei and measuring 9 to 20 ktm in diameter. Immature or "precyst" stages may possess four nuclei. Amoebae that pass from the cecum without encysting do not survive outside the host (Lin 1971). Cysts are excreted in the feces and are available for ingestion and infection in the next host. The signals involved in excystation of ingested cysts in the cecum are not known. The environmental stability of E. muris is unknown. Cell-free cultivation of E. muris has not been reported.
3. CellBiology The genus Entamoeba has classically been divided into groups based on the number of nuclei in the mature cyst forms (Neal 1966). In general, nonpathogenic Entamoeba have one nucleus (E. chattoni from nonhuman primates; E. polecki from pigs and humans) or eight (E. coli from humans; E. muris from mice) nuclei per mature cyst. The pathogenic E. histolytica and nonpathogenic E. dispar of humans possess four nuclei per mature cyst. In addition, E. invadens of reptiles possess four nuclei in the mature cyst. This classification has held up to phylogenetic analysis of ribosomal DNA and protein sequences of Entamoeba (Silberman et al. 1999). This morphologic detail is important when diagnosing amoeba infection in other species. The presence of octonucleate cysts in the feces of a reptile suggests that these are E. muris that were transiently acquired from feeder mice, while quadranucleate cysts suggest parasitism with E. invadens. Like the flagellates, Entamoeba lack mitochondria, peroxisomes, and a Golgi complex. A mitochondrial-like organelle, variably termed crypton, cryptome, or mitosome, has been identified in E. histolytica (Mai et al. 1999; McFadden 2003; Tovar et al. 1999). Its function has yet to be determined. Entamoeba spp. generate energy through glycolysis. Elongated, rhomboidshaped cytoplasmic inclusions called chromatoid bodies can be seen by light microscopy in cyst stages of Entamoeba spp. These bodies are aggregates of ribosomes, and their function is unknown (Kusamrarn et al. 1975; Neal 1966). 4. Disease and Diagnosis
Disease due to the presence of E. muris in the cecum and colon of mice has not been reported. Although molecular and
528
KATHERINE
cellular analysis of E. muris has not been reported, it is likely that this species lacks the virulence factors that are well characterized in E. histolytica. These factors include lectin-binding molecules, channel-forming amoebapores, and cysteine proteases responsible for tissue destruction in cases of invasive human amoebiasis (Espinosa-Cantellano and Martinez-Palomo 2000). E. muris can be diagnosed by examining fresh wet mounts from the cecum and colon by light microscopy. If slides are kept at 37~ amoeboid movement may be observed. When cooled, organisms tend to round up and are more difficult to identify. Visualization of octonucleate cysts is enhanced by the addition of iodine to wet mounts. In histologic sections, trophozoites are round to oval in shape and vary from 8 to 30 ~tm in diameter (Fig. 21-3). They live in colonies of varying number within the mucus layer of the cecum, and rarely, anterior colon. Trophozoites stain eosinophilic with a granular and often highly vacuolated cytoplasm. Close examination of the vacuoles may reveal ingested bacteria or other protists. Cyst forms are smaller and possess a thin cell wall. Refractile chromatoidal bodies may be present in the cytoplasm. Due to plane of sectioning, mature cysts often appear to contain less than eight nuclei. 5.
Treatment, Prevention, Control
As with the intestinal flagellates, rederived and barriermaintained mice are free of E. muris. Chemotherapeutic elimination of E. muris from infected mice has not been reported,
Fig. 21-3 Entamoeba muris cysts from the cecum of a mouse. Note variably sized intracytoplasmic vacuoles. H&E; bar = 50 l.tm.
WASSON
although luminal amoebicides (iodoquinol, diloxanide furoate, paromomycin) or imidiazoles (metronidazole) used as therapy for human amoebiasis may be effective (Upcroft and Upcroft 2001). The environmental stability of E. muris cysts has not been reported. 6.
Research Implications
Interference with experimental design or reproducibility has not been reported from mice harboring E. muris. There has been tremendous interest in creating a murine model of human amoebic dysentery and invasive amoebiasis. Unfortunately, E. histolytica is not very pathogenic for mice (Ghadirian et al. 1987; Gold and Kagan 1978; Stern et al. 1984). Early reports of mouse mortality due to intracecal inoculation with E. histolytica were complicated by the fact that the organisms were co-cultured with a cocktail of potentially pathogenic bacteria (including E. coli and clostridial species) in order to generate organisms for mouse experiments (Owen 1985, 1990). Since then, methods for cultivating E. histolytica in monoand axenic media have been developed, and more mousevirulent strains isolated (Clark and Diamond 2002). Still, patent intestinal E. histolytica infection in mice depends on intracecal inoculation of xenically passaged organisms (Thompson et al. 2000). Infection is also dependent on mouse strain: 60% of cecally inoculated C3H mice developed typhlitis and remained persistently infected 18 months post-inoculation, compared with no infection or histologic lesions in BALB/c, C57BL/6, or INF-7-, ILl2-, or iNOS-knockout mice (Cieslak et al. 1992). An interesting variation on the mouse model of amoebiasis is the SCID (severe combined inmmunodeficient) mousehuman intestinal xenograft model (Seydel et al. 1997). In this model, human fetal intestinal sections are transplanted under the skin of C.B-17-Prkdc scid (SCID) mice and allowed to develop for several weeks prior to direct inoculation into the intestinal lumen with E. histolytica trophozoites. With this model, researchers have characterized the proinflammatory cytokines released by human intestinal epithelial cells in response to E. histolytica infection and have examined the virulence of genetically manipulated E. histolytica trophozoites (Zhang et al. 2003, 2004). Models of invasive amoebiasis, the most common form in humans being hepatic abscess, can be recreated by intrahepatic inoculation of trophozoites into SCID mice (Cieslak et al. 1992).
IV.
APICOMPLEXANS
A.
General Introduction
The phylum Apicomplexa is a large and diverse group of spore-forming, sexually reproducing protozoans that share
2 1.
529
PROTOZOA
an anteriorly located, apical complex important for host-cell invasion. This phylum includes the coccidia (Cryptosporidium, Eimeria, Isospora, Klossiella, Sarcocystis, Toxoplasma gondii), piroplasms (Babesia), and hemoapicomplexans (Plasmodia). Apicomplexans of importance in the laboratory mouse include Eimeria spp., Sarcocystis muds, Klossiella muds, Cryptosporidium spp., and Toxoplasma gondii. Although apicomplexans infect a variety of cell types and cause distinctly different diseases in their respective hosts, they share a common basic life cycle (Levine and Ivens 1990). This life cycle starts with the oocyst (or spore form), which must undergo sporulation (meiosis) before becoming infective for the host. Once sporulated, the oocyst is composed of sporoblasts, each of which contains a specific number of invasive sporozoites, depending on the species. After the sporulated oocyst is ingested by a susceptible host, the sporozoites are released and invade the appropriate host cells. These forms replicate by multiple fission resulting in daughter merozoites (also referred to as schizozoites or trophozoites). Merozoites are released by rupture of the host cell and invade neighboring cells. This cycle of asexual replication, rupture, and re-invasion (referred to as merogony or schizogony) is repeated for one or more generations, depending on the parasite species. After asexual replication, the parasites cease replicating and differentiate into microgametes or macrogametes within the host cells. Macrogametes are fertilized by motile microgametes and develop into oocysts, which mature and rupture from the host cell to start the parasite life cycle again. Variations in this basic apicomplexan life cycle include length of time and location where sporulation occurs, number and arrangement of sporoblasts and sporozoites within the sporulated oocyst, tissue and cell preference of the sporozoites, number of generations of merogony, presence or absence of intermediate hosts, parasite host specificity, and production of cyst or pseudocyst forms within hosts. These variations will be dealt with when discussing the specific protozoans. Another common feature shared by these parasites is the apical complex, the defining criterion for inclusion in the phylum Apicomplexa. The apical complex refers to a group of organelles that are present in one or more stages of the protozoan's life cycle. These organellesmvisible by electron microscopyminclude the polar ring, conoid, micronemes, and rhoptries (Morrissette and Sibley 2002). The polar ring is a cogwheel-like, microtubule-organizing center from which microtubules radiate out of and down the length of the sporozoite. Engagement of these structures is thought to play a role in parasite gliding motility. The conoid is a retractable protuberance at the apical end of the parasite. It is composed of tightly coiled microtubules and is thought to be responsible for mechanical penetration of epithelial cell membranes by the invading sporozoite. Micronemes and rhoptries are secretory organelles containing proteins required for parasite motility, and adhesion to and invasion into host cells. Unlike the flagellates and amoebae, most apicomplexans possess
mitochondria and a Golgi complex. Most apicomplexans also possess an apicoplast, a chloroplast-like organelle homologous to the plastid of algae (Kohler et al. 1997; Roos et al. 1999). This organelle contains its own genomic material and is thought to have an endosymbiotic origin in apicomplexans similar to that of mitochondria in eukaryotic cells. Although its function is not completely understood, its presence is essential for parasite survival (Roos et al. 1999). In addition, because the genes on this episomal DNA appear prokaryotic in nature, it is an attractive target for drug development against the apicomplexans (McFadden 2003).
B. 1.
Eimeria spp.
Introduction
There is a surprisingly large body of literature regarding Eimeria infection in mice, especially when compared with other protozoal agents of rodents. Much of the early literature deals with species identification and life-cycle elucidation. Recent work focuses on Eimeria infection in mice as a model for investigating the immunopathogenesis of coccidiosis, a disease that accounts for large economic losses in the livestock industries. In general, members of the genus Eimeria are homoxenous (completing their life cycle in one host) and are host-specific (Fernando 1990; Levine and Ivens 1988). Most, but not all, are intestinal pathogens (Fernando 1990). Eighteen species of Eimeria have been described in Mus musculus, of which four of these (E. falciformis, E. vermiformis, E. papillata, and E. ferrisi) are considered pathogenic (Levine and Ivens 1990). Infection with Eimeria is rare in well-managed, laboratory mouse colonies. A recent study indicated a 26.3% infection rate with Eimeria spp. in a variety of wild rodents including Mus musculus (Franjola et al. 1995). 2.
Life Cycle
The life cycle of Eimeria spp. in mice is simple and direct, and follows the basic apicomplexan life-cycle scheme described above. Oocysts are shed in the feces and require 3 to 6 days in the environment to undergo sporulation (Levine and Ivens 1990). Sporulated oocysts contain four sporoblasts, each of which contains two infective sporozoites. After ingestion, sporozoites are released and invade the intestinal epithelium. As with other members in the genus Eimeria, those infecting mice have preferred host-cell and intestinal tract niches: E. falciformis infects the crypt epithelium of the cecum and colon (Fig. 21-4A); E. vermiformis, the crypt epithelium of the distal small intestine (Fig. 21-4B); E. papillata, the villus epithelium of the distal small intestine; and E. ferrisi, the villus epithelium of the cecum and colon (Table 21-1; Schito et al. 1996). E. falciformis undergoes four generations of asexual replication in the host, while E. vermiformis and E. ferrisi undergo three (Levine and Ivens 1990). The number
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Fig. 21-4 Eimeria spp. from the intestinal tract of mice. (A) Eimeria falciformis oocysts in the crypts of the proximal colon. (B) E. vermiformis oocysts in the crypts of the distal small intestine. (C) and (D) Mature meronts of E. falciformis and E. vermiformis, respectively. (E) E. falciformis microgametes (bottom of figure), and macrogametes (top of figure). A single macrogamete being fertilized by microgametes is present in the middle of the figure. (F) A cluster of E. vermiformis macrogametes. H&E; A and B, bar = 100 ~tm; C through F, bar = 50 ~tm.
21.
of rounds of asexual replication in E. papillata has not been reported. Sexual replication begins with differentiation of the parasites into microgametes and macrogametes. After fertilization, mature oocysts are released from host cells and shed in the feces to begin the cycle again. The infective dose for naive mice is 103 oocysts, and the prepatent period ranges from 4 to 8 days (Schito et al. 1996). Resistance to reinfection with Eimeria is dependent on the host mouse strain, immunocompetency, and initial parasite dose and species (Mahrt and Shi 1988; Mesfin and Bellamy 1979; Schito et al. 1996). 3.
Cell Biology
Eimeria species possess an apical complex, Golgi complex, mitochondria, and apicoplast (see Section IV.A; Adams and Todd 1983; Cai et al. 2003; Chobotar et al. 1975). Although the functions of the apicoplast genes are unknown, portions of its genome have been used to investigate phylogenetic relationships within the rodent lineages of Eimeria (Zhao and Duszynski 2001).
4.
531
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Disease and Diagnosis
Four Eimeria species are considered pathogenic for mice: E. falciformis, E. vermiformis, E. ferrisi, and E. papillata. All result in similar clinical disease and histologic lesions. Concurrent infection with multiple species has been observed in mice (Allen and Fetterer 2002). Virulence factors have not been identified in Eimeriamdisease is due to the damage sustained by the intestinal epithelium during multiple rounds of parasite replication and rupture from the host cells. Clinical signs are dose-related and include soft, mucus-covered feces with perianal staining, depression, dehydration, and anorexia (Allen and Fetterer 2002; Blagburn and Todd 1984; Mesfin et al. 1978). In experimentally infected mice, mortality rates increase with doses of 103 and higher for E. falciformis and E. vermiformis oocysts (Blagburn and Todd 1984; Mesfin et al. 1978). Histologic lesions vary by intestinal location and include multifocal mucosal erosions of the intestinal epithelium with a pyogranulomatous cellular infiltrate and hemorrhage, submucosal congestion, and edema (Blagburn and Todd 1984; Levine and Ivens 1990; Mesfin et al. 1978). Intralesional parasites, at various stages of replication within intestinal epithelial cells, may be observed during acute infection. As infection resolves, a reduction in the intestinal villus to crypt ratio, villus shortening or crypt hyperplasia, and a nonspecific mononuclear cell infiltrate may be all that is observed (Blagburn and Todd 1984). Focal granulomas with oocysts in the lamina propria of the colon have been described in resolving E. falciformis infections (Mesfin et al. 1978). Hyperplasia of the mesenteric lymph nodes and splenic follicles may also be seen histologically (Smith and Hayday 2000a). Eimeria spp. are readily observed in hematoxylin-eosin stained histologic sections. Meronts (asexually replicating forms) are
found within intracellular, parasitophorous vacuoles. Meronts vary in number and size depending on their stage of development. Immature forms are round to indistinct in shape, while mature forms are crescent or banana-shaped (Fig. 21-4C, D). Most are uninucleate and lightly basophilic. Microgametes stain intensely basophilic, are comma-shaped, and possess two to three flagella (Fig. 21-4E). Macrogametes have a prominent nucleus and nucleolus. Refractile eosinophilic material or periodic acid-Schiff (PAS) positive staining material may be observed within the cytoplasm of the macrogamete (Fig. 21-4E, F). Oocysts can be distinguished from macrogametes by the presence of one or two refractile cell walls. Speciation of Eimeria parasites can be tentatively assigned based on the tissue and cell location of parasite infection (see Section IVB2). Eimeria spp. infection can also be diagnosed by fecal flotation, although speciation is difficult. In fresh fecal pellets, oocysts are round to ellipsoidal and vary considerably in size both within and between species. Most range in size between 18-26 x 11-24 gm (see Table 21-4; Levine and Ivens 1990). Sporulated oocysts are smaller (ranging in size between 8-14 x 5-10 gm), and contain four sporoblasts with two sporozoites each. Oocyst walls of E. falciformis and E. ferrisi are smooth and clear to light brown in color; those of E. vermiformis and E. papillata are yellowish brown and pitted or papillated (Levine and Ivens 1990). 5.
Treatment, Prevention, Control
A variety of anticoccidial drugs and coccidiostats are used in the livestock industry to control Eimeria infection (Allen and Fetterer 2002). The efficacy of some of these compounds has been examined in naturally infected mice, with varying results (Haberkorn et al. 1983). Toltrazuril (Baycox| was effective in eradicating a mixed Eimeria infection in C57BL and CFW1 mice, while amprolium, several ionophores, and sulfa-based drugs were only partially effective (Haberkorn et al. 1983). Efficacy was enhanced when some of these less effective drugs where given simultaneously (Harder and Haberkorn 1989). Autoclaving is sufficient to inactivate Eimeria oocysts. E. falciformis oocysts were found to be nonviable after 7 days at 40~ (summarized in Schneider et al. 1972). However, due to the large reproductive potential of Eimeria spp., oocyst numbers build up quickly in the environment and can be difficult to eliminate from some types of laboratory animal housing (Haberkorn et al. 1983; Wilkinson et al. 2001). Rederived and barrier-maintained mice housed in clean environments are free of coccidia. 6.
Research Implications
Outbred mice previously infected with E. falciformis were noted to be resistant to experimental infection with Salmonella abortus-ovis (Lantier et al. 1981). Survival time was found to be greater in mice previously infected with E. falciformis and
532
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experimentally inoculated with Toxoplasma gondii (Chinchilla et al. 1986). The authors of these reports suggest that Eimeria infection induces a nonspecific inflammatory response that provides protection against other pathogens, but state that the mechanisms involved are unknown. Due to immunologic stimulation and potential for severe intestinal pathology, mice infected with Eimeria spp. are unsuitable for most research involving the immune system and gastrointestinal tract. Experimental Eimeria spp. infection in mice, however, has served as a marvelous tool for investigating gut immune responses to a natural murine pathogen. The stimulus for this research is the hope of developing vaccines against these pathogens in poultry and cattle (Rose et al. 1997). Primary infection with Eimeria spp. in immunocompetent mice results in cellular and humoral immune responses and clearance of infection in approximately 3 weeks (Nash and Speer 1988; Smith and Hayday 2000a). Mice are resistant to reinfection with the same species. Experiments with genetically modified mice have allowed these responses to Eimeria spp. to be immunologically "dissected." The inability of athymic nude mice to develop immunity to Eimeria falciformis suggested that T cell responses were important in clearance of infection (Mesfin and Bellamy 1979). Using several knockout mouse models, it was later shown that major histocompatibility (MHC) II-restricted CD4+ ~[3 T cells and the production of interferon gamma were important for effective primary immune responses against Eimeria spp. (Roberts et al. 1996; Rose et al. 1989, 1992). Gamma delta (y/8) T cells, noncirculating lymphocytes found in the intestinal epithelium of mice, were found to have an immunomodulatory effect on Eimeria spp. infection. Mice deficient in 3'/8 T cell receptors have more severe intestinal pathology than mice deficient in y/8 and a/I] T cell receptors when infected with E. vermiformis (Roberts et al. 1996). The ability to mount an effective secondary immune response was not altered by the lack of y/8 T cells, however. Work done with severe combined immunodeficient-beige, perforin-knockout mice, and mice treated with monoclonal antibodies to deplete CD4+ or CD8+ T cells demonstrated that cytotoxins produced by natural killer cells and cytotoxic (CD8+) T cells play a role in immunity to secondary infection with E. papillata or E. vermiformis (Rose et al. 1992; Schito and Barta 1997). Lastly, infection of TAP1, interleukin-4, Fas ligand, and inducible nitric oxide knockout mice with E. vermiformis has demonstrated that MHC class I-restricted T cells, B cell activation, and respiratory burst activity of macrophages are not important for clearance of primary Eimeria spp. infections (Smith and Hayday 2000b). C. 1.
Sarcocystis muris
Introduction
Sarcocystis muris is a coccidial parasite of murine skeletal muscle (Miescher 1843). In the early part of the twentieth
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century, the prevalence of Sarcocystis infection in laboratory mice was reported to be between 14 and 60% (Smith 1901; Twort and Twort 1932). Although modem husbandry and management practices such as separation of species, vermin control, rederivation, and barrier housing have decreased the incidence of S. muris, this parasite is still occasionally observed in laboratory-bred mice (Tillmann et al. 1999). 2.
Life Cycle
The source of Sarcocystis muris infection in mice remained obscure until 1976, when cats were identified as the definitive host (Ruiz and Frenkel 1976). Limited cross-transmission studies demonstrate that Mus musculus is the only intermediate host for S. muris (Ruiz and Frenkel, 1976). The life cycle follows the basic scheme as described above with a few exceptions: Oocysts sporulate within the carnivore's intestinal tract and are immediately infective when released into the environment; sporulated oocysts contain two sporocysts, each of which contains four sporozoites; after ingestion of sporocysts by mice, asexual replication first occurs in the liver, then in the skeletal muscle; in the muscle, replication of parasites results in the formation of large cystic structures (sarcocysts) filled with thousands of organisms (bradyzoites). When infected skeletal muscle is consumed by the carnivore, bradyzoites are released, invade the host intestinal epithelium, and undergo sexual replication. This heteroxenous life cycle begins again with the release of sporocysts into the environment. Recent data suggest that S. muris can also be sustained within a mouse colony due to cannibalism of infected individuals (Koudela and Modry 2000). 3.
Cell Biology
Sarcocystis muris possess the standard complement of organelles found in apicomplexans, including an apical complex, Golgi complex, and mitochondria. An apicoplast organelle has been identified in S. muris by electron microscopy (Hackstein et al. 1995). Partial sequencing of this apicoplast genome revealed an herbicide-binding region, suggesting that S. muris may be susceptible to triazine chemotherapeutics such as toltrazuril (Hackstein et al. 1995).
4.
Disease and Diagnosis
Difficulty moving is the only clinical symptom associated with sarcocystosis in mice (Ruiz and Frenkel 1976). Mice of that report were experimentally inoculated, and approximately 80% of muscle fibers were infected with S. muris by 100 days post-inoculation (Ruiz and Frenkel 1976). A dose of 50 sporocysts is reported to result in widespread and severe muscle infection with S. muris in NMRI and AKR/N mice (Rommel et al. 1981). Natural infections in mice are usually not that severe (Tillmann et al. 1999; Twort and Twort 1932).
2 1.
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Diagnosis of S. muris is made by histologic examination of skeletal muscle; organisms are occasionally observed in the myocardium (Tillmann et al. 1999; Twort and Twort 1932). Within muscle, spherical or cylindrical cysts--often several millimeters long--can be seen within myocytes. Hundreds of banana-shaped, uninucleate bradyzoites are present within the cysts. Often there is no inflammatory reaction to these structures. Bradyzoites are visible with routine hematoxylineosin and periodic acid-Schiff staining, and measure 4-6 x 14-16 ~tm (Ruiz and Frenkel 1976). Sporocysts are shed in the feces of infected cats and can be diagnosed by fecal floatation. Sporocysts measure 7.5-9 x 8.7-11.7 ~tm, contain four sporozoites, and have a smooth and colorless wall (Cawthorn and Speer 1990; Ruiz and Frenkel 1976). 5.
Prevention, Treatment, Control
There is a single report documenting elimination of S. muris from the livers of experimentally infected mice with sulfaquinoxaline and pyrimethamine (Rommel et al. 1981). It is not clear if this drug combination was effective against bradyzoite forms found in muscle. Other anticoccidial drugs were not effective (Rommel et al. 1981). Sporocysts are environmentally resistant and remain infectious for at least 119 days at 21~ in fecal flotation solutions (Smith and Frenkel 1978). Although Sarcocystis spp. have an obligatory heteroxenous life cycle (requiring definitive and intermediate hosts), direct contact with cats or cat feces is not needed to transmit S. muris to mice. Cockroaches exposed to S. muris-infected cat feces transmitted infection to naive mice for 20 days (Smith and Frenkel 1978). Cannibalism--although less efficient--will sustain S. muris in a mouse population (Koudela and Modry 2000). For these reasons, rederivation and maintenance in a clean barrier is recommended to eliminate S. muris from mouse colonies. 6.
Research Implications
Experimental infection with S. muris was found to suppress humoral and cell-mediated immune responses to vaccination with unrelated proteins, and to induce splenomegaly in mice (Gill et al. 1988a, 1988b). These nonspecific immune aberrations make infected mice unsuitable for research involving the immune system.
D. 1.
KlossieUa muris
Introduction
Klossiella muris is the least characterized apicomplexan parasite of mice. Infection in the kidneys of mice was first reported in 1889 and further described in 1902 (Smith and Johnson 1902). In a 1932 histologic survey of mice used in carcinogenesis studies, renal infection with K. muris was estimated to be 60% (Twort and Twort 1932). Infection in
wild-caught mice has been reported as high as 93% (Rosenmann and Morrison 1975). Under current laboratory animal housing conditions, infection with K. muris is seldom reported. 2.
Life Cycle
Infection in mice begins with ingestion of sporocysts and uptake into the portal vascular system (Yang and Grice 1964; Smith and Johnson 1902). Sporozoites are released, circulate through the vascular system, and preferentially invade the endothelium of the glomeruli, where they undergo several rounds of asexual replication (Fig. 21-5A). Parasites have also been observed in capillaries and arterioles of the lungs, liver, spleen, and thymus (Twort and Twort 1932). At some point, asexually replicating forms rupture from the glomeruli into Bowman's space and pass to the renal tubules. Parasites invade the tubular epithelium and differentiate into microand macrogametes. Fertilization results in the formation of sporonts, each of which contains as many as 30 sporoblasts and a residual body (Fig. 21-5B; Levine and Ivens 1990; Smith and Johnson 1902; Yang and Grice 1964). Sporoblasts mature into sporocysts, each of which in turn contains 25 to 35 crescent-shaped sporozoites. The life cycle is completed when infective sporocysts are released from ruptured host cells, pass into the urinary bladder, and are shed into the environment. A homoxenous life cycle has been inferred due to the large number of infected mice diagnosed histologically after remaining caged together for several months (Smith and Johnson 1902). 3.
Cell Biology
Ultrastructural examination of Klossiella life-cycle stages and organelles has not been reported. They presumably possess components of the apical complex found in other apicomplexans. The means by which they identify and invade host cells, generate energy, and acquire nutrients for replication have not been investigated. 4.
Disease and Diagnosis
Clinical disease has not been reported with K. muris infection in mice. Diagnosis is usually made histologically and is dependent on identifying replicating forms within the glomerular endothelium or renal tubular epithelium (Smith and Johnson 1902; Yang and Grice 1964). Asexually replicating forms (merozoites) often cause distention of endothelial cells and may appear as multiple minute (< 0.5 ~tm) bodies surrounded by clear halos within a parasitophorous vacuole in the cell (Yang and Grice 1964). Sporocysts are subspherical, measuring 16 x 13 ~tm, and are located in the renal tubule epithelium (Levine and Ivens 1990; Yang and Grice 1964). Multiple banana-shaped sporozoites may be seen budding within each sporocyst (Levine and Ivens 1990; Yang and Grice 1964). Heavily parasitized kidneys show evidence of nonsuppurative
534
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Fig. 21-5 Klossiella muris from the kidneys of a mouse. (A) Merozoites replicating in endothelial cells of a glomerulus. (B) Sporont within renal tubular epithelial cell; note budding sporoblasts and residual body. (C) High magnification of mature sporocysts in the renal tubular epithelium. (D) Low magnification of K. muris infection in the renal medulla of a mouse. H&E; A through D, final magnification unknown (Images courtesy of S. W. Barthold).
interstitial nephritis with tubular degeneration. Often, there is no inflammatory response associated with intracellular stages (Yang and Grice 1964). Organisms are periodic acid-Schiff negative but readily identifiable by routine hematoxylin-eosin staining (Fig. 21-5C, D; Yang and Grice 1964). Clinical diagnosis of Klossiella equi by examination of urine has been described and should be possible for diagnosis in mice (Reppas and Collins 1995). However, the authors of that report note that sporocysts were destroyed by fecal flotation solutions and that organisms were apparent only after examining the pellet of centrifuged urine samples (Reppas and Collins 1995).
5.
Prevention, Treatment, Control
Treatment of endemic K. muris infection with coccidiostats reduced the histologic incidence of infection in mice from 93. to 23% (Rosenmann and Morrison 1975). Unfortunately, that report does not specify the compound, dose, or route used. Infection does not occur in rederived, barrier-maintained mouse colonies.
6.
Research Implications
A single report documents the effects of K. muris infection in mice on research results (Rosenmann and Morrison 1975). Endemically infected mice exhibited decreased oxygen consumption and endurance when compared with uninfected controls. The authors suggest that under certain environmental conditions, K. muris infection impairs the metabolic capabilities of infected mice.
E. 1.
Toxoplasma gondii
Introduction
Toxoplasma gondii was first described in 1908, in Ctenodactylus gundii (or "gundi"), a guinea pig-like rodent found in North Africa (Nicolle and Manceaux 1908). Although T. gondii was subsequently shown to cause disease in a variety of animals around the world, it took 50 years to identify the cat as the definitive host (Hutchison 1965). Hutchison correctly
21.
surmised that cats excreted infective forms in their stools. However, concurrent Isospora spp. infection in his experimental cats obscured identification of T. gondii oocysts on fecal examination. In addition, the presence of Toxocara cati ova in the feces led him to conclude that T. gondii was transmitted in conjunction with this common nematode of cats (Hutchison 1965). This was quickly rectified once parasite-free cats were used for transmission experiments (Sheffield and Melton 1969). Mice of the genus Mus are but one of several hundred mammals (including humans), birds, and reptiles since identified as intermediate hosts in the complex life cycle and biology of this protozoal parasite. Natural infection of most animals (including mice) with T. gondii is subclinical. Because of the pervasiveness of this parasite and its role in human and animal disease on a global basis, there is a tremendous amount of published research on various aspects of toxoplasmosis. Mice have played an integral role in understanding the virulence and pathogenesis of this organism. 2.
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Life Cycle
The life cycle of T. gondii is perhaps the most complicated of all the apicomplexans of mice. Oocysts are shed in the feces of infected cat s, and sporulate in the environment after several days. Sporulated oocysts contain two sporocysts with four sporozoites each and can be differentiated on fecal floatation from Isospora spp. by size: T. gondii oocysts measure 12 x 10 ~m, while those of Isospora spp. are larger: I. rivolta measure 22 x 17 and I. felis 42 x 35 ~tm (Foyet 2001). After ingestion, sporozoites are released in the small intestine of intermediate hosts. In experimentally infected mice, sporozoites can be identified in cells of the ileal lamina propria as early as 2 hours post-inoculation (PI; Dubey, Speer et al. 1997). After invading a suitable host cell, sporozoites differentiate into tachyzoites and replicate asexually for an infinite number of generations. These rapidly replicating ("tachy-" = fast) asexual forms are responsible for "acute" toxoplasmosis. From the small intestine, parasites invade additional tissues after being transported by blood or lymph. Unlike other apicomplexans, T. gondii can infect virtually any nucleated cell in the host. In experimentally infected mice, tachyzoites can be identified in the mesenteric lymph nodes 8 hours PI (Dubey et al. 1997). By day 3 PI, parasites are present in the lungs, spleen, and kidneys; by day 4, in the heart and liver; by day 6 in the pancreas and brain; and by day 7, in the skeletal muscle (Dubey et al. 1997). If the host survives acute infection, most parasites in these peripheral locations are cleared by the immune response. This immune pressure is thought to stimulate tachyzoites to differentiate into bradyzoites within a protective tissue cyst wall. These cyst forms are typically found in the central nervous system, or skeletal or cardiac muscle (Dubey et al. 1997). In experimentally infected mice, tissue cysts are observed by day 15 PI (Dubey et al. 1997). These slowly replicating ("brady-"= slow) asexual forms are responsible for "chronic" toxoplasmosis in
intermediate hosts. Ingestion of tissue cysts by felids results in release of bradyzoites, invasion into intestinal epithelial cells, and differentiation into the sexually replicating forms of the parasite. As with the other coccidians, fertilization of macrogametes by microgametes results in the formation of oocysts and completion of the life cycle. Although the life cycle of T. gondii is considered heteroxenous, mice can sustain T. gondii infection through cannibalism or congenital transmission, in addition to ingestion of sporulated oocysts. Stage conversion between tachyzoites, bradyzoites, and oocysts can be demonstrated in vitro (Dubey et al. 1998). However, in vivo generation and collection of oocyst or tissue cysts from cats or mice, respectively, are usually required to generate sufficient numbers of organisms for experimental work. 3.
Cell Biology
T. gondii possesses an apical complex, mitochondrion, Golgi complex, and apicoplast similar to the other apicomplexans (see Section IV. A). The large complement of secretory organelles (micronemes and rhoptries of the apical complex, dense granules) and their functions are best--though not completelyiunderstood in T. gondii. The evolution of this extensive network of secretory organelles is thought to be an adaptation to an obligate intracellular lifestyle. Micronemes release proteins involved in the early stages of tachyzoite attachment and invasion into host cells (Carruthers et al. 1999). Depletion of microneme proteins results in transient loss of tachyzoite infectivity. Rhoptries are distinctive, club-shaped organelles numbering 8 to 16 per tachyzoite. Rhoptry proteins are secreted later in the host-cell invasion process; some are also incorporated into the parasitophorous vacuole that encloses tachyzoites once within the host cell (Carruthers and Sibley 1997). Dense-granule proteins are secreted late in the invasion process. These proteins insert in the parasitophorous membrane and are thought to be involved with nutrient acquisition and importation from the host cytoplasm to the replicating tachyzoites (Carruthers 1999). Formation of these organelles, and the biogenesis, trafficking, processing, storage, and secretion of their respective protein cargos suggest that an intricate and complex network of secretory pathways are present in apicomplexans. These pathways must be executed in a precise and sequential manner for parasite survival. A second set of unique organelles found in T. gondii are the acidocalcisomes (Moreno and Zhong 1996). First identified in Trypanosoma cruzi, these structures have since been identified in other trypanosomatids, apicomplexans, algae, slime molds, and the bacterium Agrobacterium tumifaciens (Docampo et al. 1995; Docampo and Moreno 2001; Seufferheld et al. 2003). As the name implies, these organelles contain acidified stores of calcium and other elemental minerals. Although the exact function of acidocalcisomes is not known--and may vary between species--several functions have been proposed. These include as an energy source, an intracellular calcium store for
536
parasite signaling pathways, a mechanism for intracellular pH homeostasis, and for parasite osmoregulation (Docampo and Moreno 1999). In addition, since acidocalcisomes are not present in mammalian cells, they represent an attractive target for antiprotozoal therapies.
4.
Disease and Diagnosis
T. gondii infection in mice is subclinical with minimal gross necropsy lesions (Perrin 1942). Virulence of experimental infection is dependent on mouse strain, T. gondii-type strain, stage of parasite (sporulated oocysts being more pathogenic than bradyzoites, which in turn are more pathogenic than tachyzoites), and parasite dose (Araujo et al. 1976; Suzuki et al. 1995). Swiss Webster mice inoculated with the mildly pathogenic type III strain of sporulated oocysts remained clinically normal until 3 weeks PI, at which time they appeared unthrifty, lost weight, and developed paralysis (Dubey et al. 1997). Mice euthanized at early time points had enlarged and edematous mesenteric lymph nodes. The ileum was congested and edematous, with small white pinpoint foci visible through the serosa. Necrosis was the prominent histologic lesion, with the organs involved dependent on the time point of infection at which mice were euthanized. Necrosis and infarction of the ileal lamina propria and mesenteric lymph nodes were seen in the first days of infection. Focal hepatitis and myocarditis with mixed leukocytic infiltration were observed during the first week of infection. Interstitial pneumonia with intralesional parasites developed during the second week of infection. Intra- and extracellular tachyzoites were seen in brain parenchyma as early as 9 days PI. By 87 days after inoculation, bradyzoite-filled tissue cysts with little associated inflammation were present in most mouse brains (Dubey et al. 1997). Diagnosis of toxoplasmosis in mice is primarily by histology. Individual tachy- and bradyzoites are difficult to identify in histologic sections, but tissue cysts may be observed in the CNS, myocardium, or skeletal muscle (Fig. 21-6A). Cysts are spherical and of variable size, with a thin wall that is argyrophilic and faintly periodic acid-Schiff (PAS) positive (Frenkel 1956). Cysts are packed with PAS positive, fusiform bradyzoites. Unstained impression smears of brain material can also be screened for the presence of T. gondii tissue cysts (Dubey et al. 1998). In intermediate hosts with patent parasitemia, individual tachyzoites may be identified in the lamina propria of the small intestine, endothelium of the small intestine or lungs, or within leukocytes on peripheral blood smears. Pre-cyst intracellular clusters of tachyzoites may also be observed in heart and skeletal muscles (Fig. 21-6B). Tachyzoites have a centrally located nucleus and measure 2-3 x 6-7 ~tm (Moller 1968). Tachyzoites exhibit a gliding motility that is usually only appreciated in tissue culture systems. T. gondii infection in mice can also be diagnosed by PCR and serology (James et al. 1994).
KATHERINE 5.
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Treatment, Prevention, Control
Because of a short shedding period and subclinical signs, cats with T. gondii infection usually do not receive treatment (Dubey 1994). Acute toxoplasmosis in intermediate hosts is amenable to chemotherapeutic intervention, usually a combination of pyrimethamine and sulfa drugs (Wilson et al. 2003). Tissue cyst forms are resistant to treatment, however. Experimentally infected mice treated with sulfonamides for 15 days had a lower mortality rate than those treated with chlortetracycline or left untreated (Frenkel 1956). In general, treatment of T. gondii infection in intermediate hosts is not warranted. Prevention of infection in rodent colonies relies on separation of species and elimination of potential transport vectors from the environment (Chinchilla et al. 1994). Because T. gondii can also be sustained through cannibalism and congenital infection, rederivation and barrier maintenance are recommended for naturally infected mouse colonies (Beverley 1959). As with other apicomplexans, the oocysts of T. gondii are remarkably hardy. Oocysts retain infectivity when stored at 4~ for 54 months or 0~ for 13 months; oocysts lose infectivity at temperatures above 40~ (Dubey 1998). Oocysts also survive exposure to 5.25% sodium hypochlorite (bleach) solution (Dubey et al. 1997). This method is used to "sterilize" oocysts harvested from cat feces prior to tissue culture or mouse inoculation. Tachyzoites and bradyzoites are less environmentally stable, although tissue cysts (filled with bradyzoites) retain infectivity when stored at 4~ for several months (Dubey 1997). Autoclaving or heat treatment at 70~ for 10 minutes will inactivate T. gondii oocysts from bedding, caging, and other equipment (Dubey 1994). 6.
Research Implications
Interference with research due to endemic toxoplasmosis has not been reported for mice. Several studies examining the behavioral effects of experimental T. gondii infection in mice have been published, however, and suggest that subclinical infection would interfere with behavioral phenotyping of mice. Experimentally and congenitally infected mice exhibited decreased motor coordination, spent less time grooming and exploring novel areas, and spent more time running on a home-cage wheel than uninfected control mice (Hay et al. 1983, 1984, 1985). The authors attributed these behavioral abnormalities to subclinical T. gondii-induced encephalitis in the otherwise clinically healthy mice, and suggested that this behavior increased their chance of predation (and therefore perpetuation of toxoplasmosis) by cats (Hay et al. 1985). More recent experimental mouse work has focused on three areas of importance in human toxoplasmosis: ocular, encephalitic, and congenital toxoplasmosis. Ocular toxoplasmosis in humans is thought to result from reactivation of congenital T. gondii infection, and occurs in both immunocompetent and
21.
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537
Fig. 21-6 Toxoplasma gondii tissue cyst in the cerebellum (A) and tachyzoites in the heart muscle (B) of a wallaby. Note the difference in shape and size of bradyzoites within the mature cyst compared to the intracellular tachyzoites. (C) Cryptosporidium parvum in the small intestine of a mouse. (D) Higher magnification of C. parvum. Note the variation in size of the merozoites and their intracellular but extracytoplasmic location in the enterocytes. H&E; A, B and D, bar = 50 ~m; C, bar = 100 ~tm (Images A, B courtesy of S. Tunev).
538
immunosuppressed individuals. Ocular toxoplasmosis causes a recurrent and progressive, necrotizing retinochoroiditis that eventually results in blindness of the affected eye (Gutierrez 2000). The disease can be experimentally reproduced in mice by intraperitoneal injection of tissue cysts or by intraocular injection of tachyzoites (Hu et al. 1999; Lyons et al. 2001). Using the mouse model, researchers have shown that T. gondii infection results in an upregulation of both inflammatory and anti-inflammatory mediators of the immune system and in apoptotic pathways in ocular tissues (Hu et al. 1999; Lyons et al. 2001). These responses may serve as a mechanism to minimize inflammation in the otherwise immune-privileged eye by eliminating parasite-infected cells while preserving uninfected tissues. As with ocular disease, toxoplasmic encephalitis results from reactivation of a congenital or subclinical infection. Toxoplasmic encephalitis tends to occur in profoundly immunosuppressed individuals and generally carries a poor prognosis (Suzuki 2002). A murine model of reactivated toxoplasmosis can be created by inoculating virulent strains of T. gondii into sulfadiazine-treated mice. Antibiotic treatment suppresses--but does not eradicate--parasite infection. This prevents development of acute toxoplasmosis while allowing the infection to establish itself in brain and muscle tissues. "Reactivation" of infection (renewed proliferation of tachyzoites) and development of fatal encephalitis occur when antibiotics are discontinued. Using resistant, susceptible, and genetically modified mouse strains, researchers have demonstrated that the source of gamma interferon (INF-'f), an immune mediator critical for preventing toxoplasmic encephalitis, is produced by T cells and an as-of-yet unidentified population of non-T cells within the brain (Kang and Suzuki 2001). This non-T cell source of INF-), appears critical for host resistance against T. gondii encephalitis. Mice have also been key to determining the genetics of resistance to toxoplasmic encephalitis. Mice with the d haplotype (e.g., BALB/c mice) in the D region of the major histocompatibility complex (MHC) are resistant, while those with the b or k haplotype (C57BL/6 mice) are susceptible (Suzuki et al. 1994). This correlates with what is seen in human infection. Humans with the HLA-DQ3 haplotype of the MHC are more susceptible to toxoplasmic encephalitis than those with the HLA-DQ 1 haplotype. Mice engineered to express human HLA-DQ1 had a greater degree of protection against toxoplasmic encephalitis compared with those expressing human HLA-DQ3 (Mack et al. 1999). Lastly, mice have been used as a model to investigate aspects of congenital transmission of toxoplasmosis. BALB/c mice infected with tissue cysts on day 12 of gestation--but not 8 weeks prior to breeding, or at 8 weeks prior to and on day 12 of gestation--gave birth to T. gondii-infected pups (Roberts and Alexander 1992). Pups had histologic evidence of toxoplasmic encephalitis, and some developed ocular disease at 2 months of age. Using this model, researchers have demonstrated the importance of natural killer cells--an important
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component of the innate immune response--in protecting the fetus from congenital transmission of T. gondii (Abou-Bacar et al. 2004). This model has also been useful for evaluating the efficacy of candidate vaccines against toxoplasmosis (Roberts et al. 1994). This na'fve pregnant mouse model replicates several of the epidemiologic and clinical features of human congenital toxoplasmosis. These include infection rates that are highest in fetuses born to mothers exposed to T. gondii for the first time during pregnancy; protection of fetuses against infection if mothers have been previously exposed to T. gondii; and a high incidence of ophthalmic and neurologic disorders in congenitally infected infants (Gutierrez 2000).
E 1.
Cryptosporidium muris
Introduction
Cryptosporidium muris is the type species for a genus of parasites that were considered biomedical curiosities until the advent of HIV and AIDS (Current and Garcia 1991). Shortly after he identified C. muris in the gastric glands of mice, Ernest Tyzzer went on to identify and describe Cryptosporidium parvum, a second species that resided in the small intestine of mice (Tyzzer 1910, 1912). It wasn't until 70 years after Tyzzer's first descriptions that Cryptosporidium spp. were recognized as a cause of self-limiting diarrhea in immunocompetent humans and of severe life-threatening diarrhea in immunocompromised patients (Peterson 1992). There are now 13 validated Cryptosporidium species described, and most of these infect a wide range of hosts (Table 21-2; Xiao et al. 2004). Humans are the primary host for C. hominis but can be infected with various isolates of C. parvum and C. muris (Palmer et al. 2003; Xiao et al. 2004). Cryptosporidial infection of contemporary mouse colonies is seldom reported; natural infection with C. muris and/or C. parvum is occasionally observed in wild rodent populations (Bajer et al. 2003; Klesius et al. 1986; Torres et al. 2000).
2.
Life Cycle
Tyzzer's 1910 light microscopic description of the life cycle and morphology of C. muris is remarkably accurate (Tyzzer 1910). Despite the parasite's small size, he determined that the parasite had asexual and sexual stages, formed a unique organ of attachment to the host cell, produced two types of oocysts, and was likely related to coccidia (Tyzzer 1910). Seventy-five years later, electron microscopy confirmed Tyzzer's findings, with few exceptions (Current and Reese 1986). Like Eimeria, Cryptosporidium spp. are homoxenous and follow the general apicomplexan life cycle described in Section IV. A. Sporulated oocysts are shed in feces. Once consumed, four sporozoites excyst and invade the gastrointestinal epithelium. These develop within a parasitophorous vacuole in the host cell
2 1.
into type I merozoites. Type I merozoites replicate asexually, rupture from the host cell, and re-invade neighboring epithelium, developing into either type I or type II merozoites. Type I merozoites continue the asexual replication cycle, while type II merozoites develop into micro- or macrogametes. After fertilization by microgametes, macrogametes develop either into thick-walled or thin-walled oocysts. Oocysts undergo sporulation and contain four "naked" sporozoites prior to rupture from the host cell (there are no sporocysts). Thick-walled oocysts are shed as fully infective parasites into the environment; thin-walled oocysts excyst while still in the host, infect new gastrointestinal epithelial cells, and initiate the asexual replication cycle again (Tyzzer 1910). The autoinfective ability of type II merozoites and thin-walled oocysts can result in an overwhelming parasitism despite a relatively small, initial inoculum, particularly in the immunocompromised host (Current and Garcia 1991). C. muris infects and replicates in the gastric mucosa, while C. parvum infects small intestinal epithelium (Current and Reese 1986; Tyzzer 1910). The complete life cycle of Cryptosporidium spp. has yet to be replicated in vitro. Oocysts for in vitro experimentation are usually purified from the feces of experimentally inoculated neonatal ruminants or suckling mice (Arrowood 2002; Current 1990). Oocysts can be induced to excyst upon exposure to sodium hypochlorite (bleach) or trypsin and bile salts, and the released sporozoites will infect a number of different cell culture lines and replicate asexually (Arrowood 2002). However, production of significant numbers of mature oocysts in vitro has not been reported (Arrowood 2002). 3.
539
PROTOZOA
Cell Biology
Cryptosporidium spp. possess an apical complex, complete with preconoid rings, conoid, rhoptries, and micronemes, which warrants their inclusion in the phylum Sporozoa (Current and Reese 1986). However, they lack a Golgi complex, mitochondrion, and the apicoplast present in other coccidia (Abrahamsen et al. 2004; Current and Reese 1986; Zhu et al. 2000). In addition, they inhabit a unique intracellular, extracytoplasmic location in their host cells (Current and Reese 1986; Goebel and Brandler 1982). At the interface between the parasite and host-cell cytoplasm, a unique "feeding organelle" can be seen by electron microscopy (Current and Reese 1986). This organelle is present during asexual and sexual parasite replication, and is composed of a portion of the parasitophorous vacuole membrane and a microfilamentous sheet (Current and Reese 1986). As the name implies, this organelle is thought to be important in nutrient transport from the host cytoplasm to the developing parasite (Current and Reese 1986). Recent work demonstrates that this organelle is composed, in part, of a membrane transporter protein that belongs in the ATP-binding cassette protein superfamily. This superfamily of proteins regulates the flow of diverse compounds across cell membranes (Perkins, Riojas et al. 1999). Recent publication of the
C parvum genome reveals that these parasites also possess several "plant-like" and bacterial-type enzymes that are either absent or divergent from those found in mammals (Abrahamsen et al. 2004). These proteins are likely to be good candidates in the search for drug targets against human and animal cryptosporidiosis.
4.
Disease and Diagnosis
Suckling mice infected with C muris or C. parvum are reported to grow more slowly and are less active than their uninfected littermates (Current and Reese 1986; Tyzzer 1910). Neonatal mice are susceptible to experimental infection with C. parvum up until 14 days of age, at which time they clear and remain resistant to infection (Novak and Sterling 1991). Clinical signs of C. muris infection in immunocompetent adult mice are not reported, and it is not clear if an age-related clearance of infection occurs for this species (McDonald et al. 1996; Tyzzer 1910). Experimental infection of immunosuppressed mice results in persistent infection with clinical signs of weight loss and sticky stools (Perryman and Bjorneby 1991; Ungar et al. 1990). Diagnosis is made by histologic examination of gastrointestinal tract tissue. Dilation of gastric glands and minimal lymphoid infiltrate are seen in mice infected with C. muris (Tyzzer 1910). In heavy C. parvum infection, blunting and fusion of intestinal villi, crypt hyperplasia, and moderate lymphocytic-plasmacytic infiltration of the underlying lamina propria are seen (Current and Reese 1986). In athymic nude mice, C. parvum can also be seen infecting the hepatobiliary tree and pancreatic ducts (Ungar et al. 1990). Small, round, basophilic-staining parasites may be observed by routine hemoxylin-eosin staining of histologic sections of the gastrointestinal mucosa. Life-cycle stages are difficult to differentiate by light microscopy due to their small size. Meronts and oocysts of C. muris measure 5 x 7 ~tm and are located on the luminal surface of gastric epithelium (Tyzzer 1910). Similar stages of C. parvum measure 5 ~tm or less in diameter and are found "decorating" the brush border of ileal enterocytes (Fig. 21-6C, D; Current and Reese 1986). These differences in size and tissue location can be used to differentiate between the two species in mice (Perryman 1990). Diagnosis can also be made by fecal flotation. Oocysts are spherical or ellipsoidal, with a smooth colorless wall and a faint suture line running down one-half the oocyst length. C. muris oocysts measure 7-8 x 5-6.5 ~tm; those of C. parvum are smaller, measuring 4.5-5 x 4-5 ~tm (Levine and Ivens 1990). 5.
Prevention, Treatment, Control
In immunocompetent animals and humans, infection with Cryptosporidium spp. is self-limiting. Although treatment of laboratory mice for natural infection with C. muris or C. parvum
has not been reported, suckling and immunosuppressed mice have been a useful in vivo model to test anticryptosporidial
540
compounds (see Blagbum and Soave 1997 for a summary of literature). Fifteen common anticoccidial compounds (including amprolium and sulfaquinoxaline) were found to be ineffective in protecting neonatal mice against infection with C. parvum (Angus et al. 1994). Lack of the apicoplast organelle in Cryptosporidium spp. has been hypothesized as one reason why these parasites are not susceptible to chemotherapeutics effective in other apicomplexans (Abrahamsen et al. 2004; Zhu et al. 2000). In addition to a lack of effective drug treatment, Cryptosporidium oocysts remain infective in chlorinated water (Carpenter et al. 1999; Korich et al. 1990). Oocysts are also remarkably resistant to common disinfectants. In an experiment examining the infectivity of oocysts after 18 hours incubation in various agents, only those treated with 10% formalin or 5% ammonia were noninfective for the suckling mouse model (Campbell et al. 1982). Steam sterilization, pasteurization, and ethylene oxide exposure for 24 hours will inactivate oocysts (Fayer et al. 1996; Harp et al. 1996). However, oocysts retain their infectivity when stored in 2% potassium dichromate solution at 4~ for greater than 6 months (Arrowood 2002). Because of a lack of treatment, resistance to common disinfectants, the environmental stability of oocysts, and demonstrated zoonotic potential of Cryptosporidium spp., endemically infected laboratory mouse colonies should be rederived and barrier maintained. 6. ResearchImplications Interference with research results due to natural infection in mice with C. muris has not been reported. In addition, C. parvum is thought to be primarily a pathogen of ruminants and humans, and is not likely to be identified in barrier-maintained mouse colonies (Klesius et al. 1986; Xiao et al. 2004). On the other hand, suckling mice have been useful for generating small amounts of pure isolates of Cryptosporidium parvum oocysts for in vitro work (Current 1990). Although Cryptosporidium spp. are not pathogenic for mice, work done in immunocompromised mouse models has helped elucidate the immune mechanisms responsible for clearance of infection. Similar to infection with Eimeria spp., clearance of C. muris and C. parvum infection was found to be dependent on production of interferon gamma and presence of CD4+ intraepithelial T cells (Chen et al. 1993; Culshaw et al. 1997; McDonald et al. 1996). The course of infection in mice with C. muris is also dependent on major histocompatibility complex (MHC) types. BALB/c mice (with the H-2d MHC type) were shown to produce fewer oocysts and to recover from infection with C. muris faster than BALB/B (H-2b) or BALB/K (H-2k) mice (Davami et al. 1997). This was attributed to a difference in interleukin-4 secretion by splenocytes between these strains of mice (Davami et al. 1997). Lastly, patent infection with Cryptosporidium spp. appears to depend on the host intestinal microflora. Axenic C.B-17/IcrTac-Prkdc sCid mice
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shed oocysts sooner when infected with C. parvum than their cohorts bearing intestinal flora (Harp et al. 1992). The authors speculate that resistance to initial infection is due to competition by endogenous microflora for preferred intestinal niches.
V. A.
MICROSPORIDIA
Encephalitozoon cuniculi
1. Introduction Mice can be naturally and experimentally infected with Encephalitozoon cuniculi, the type species for a burgeoning
collection of parasitic organisms that are more closely related to fungi than protozoans. Although frequently associated with rabbits and mice, E. cuniculi is a much more promiscuous organism than originally thought and has been identified as a pathogen in hamsters and guinea pigs, squirrel monkeys and cotton-top tamarins, carnivores (including fox and mink), domestic dogs and cats, goats, sheep, pigs, horses, muskrats, and shrews; and is a suspected cause of equine abortion (PattersonKane et al. 2003; Reetz et al. 2004; Wasson and Peper 2000; Wasson and Zbka 2003). It is also a pathogen of immunocompromised and immunocompetent humans (De Groote et al. 1995a; Mertens et al. 1997; van Gool et al. 2004). E. cuniculi infection was first described in rabbits in 1922 and in mice two years later (Cowdry and Nicholson 1924; Wright and Craighead 1922). E. cuniculi was reported in laboratory mice frequently thereafter, often in conjunction with additional parasite infestations including fleas, lice, pinworms, cysticerci, K. muris, S. muris, and T. gondii, and subclinical bacterial and--in all likelihoodmviral infections (Innes et al. 1962; Lackey et al. 1953; Morris et al. 1956; Perrin 1942; Perrin 1943; Twort and Twort 1932). In several of these reports, granulomatous hepatitis or meningoencephalitis associated with endemic E. cuniculi confounded attempts to develop murine models of human viral hepatitis or encephalitis, respectively. It was estimated that E. cuniculi was prevalent in 15 to 50% of laboratory mouse colonies at this time (Innes et al. 1962; Twort and Twort 1932). This older literature variably refers to this organism as "haplosporidia," "mouse ascitic agent," Nosema muris, E. muris, or E. rabii (Lainson et al. 1964; Morris et al. 1956; Twort and Twort 1932). By 1964, the name Nosema cuniculi was proposed, to reflect its morphologic similarity by light microscopy to the familiar Nosema spp. parasites of insects (Weiser 1964). However, ultrastructural comparison of E. cuniculi with Nosema spp. confirmed the former's status as a separate genus and reaffirmed its original name as given by Levaditi et al. 1923 (Levaditi et al. 1923; Pakes et al. 1975; Sprague and Vernick 1971). Until 1985, E. cuniculi was considered the only microsporidial parasite of mammals. With the arrival of HIV and AIDS, six generamrepresenting approximately
2 1.
12 speciesmhave since been identified as human pathogens (Franzen and Muller 1999). Mice appear to be natural hosts for E. cuniculi only. However, they can be experimentally infected with several of these newly described species. And although rare, E. cuniculi infection is still reported in contemporary mouse colonies (El Nass et al. 1998). 2.
Life Cycle
E. cuniculi is an obligate intracellular parasite with a simple and direct life cycle, and exists outside the host as an environmentally resistant spore (Wasson and Peper 2000). The initial site of E. cuniculi infection in mice is not known but is thought to be the intestinal epithelium. Once ingested, E. cuniculi infects susceptible host cells through the deployment of a preformed polar tube (discussed below). This polar tube is extruded from the spore, penetrates a susceptible host-cell membrane, and deposits the spore's infectious sporoplasm in the host-cell cytoplasm. The sporoplasm proliferates, resulting in the production of numerous plasmodial cells (also referred to as meronts) within a parasitophorous vacuole. These cells are amorphous with an indistinct nucleus and a simple cytoplasm when examined by transmission electron microscopy. Plasmodial cells develop into sporonts as the cell-limiting membrane (plasmalemma) becomes electron dense and the nucleus more distinct. Sporonts undergo binary fission to produce sporoblasts, which are no longer capable of dividing. Sporoblasts begin depositing organelles, including the polar tube and its associated polaroplast membrane, endoplasmic reticulum, the posterior vacuole, ribosomes, and electron dense bodies. Mature spores can be distinguished from sporoblasts by the presence of a thick spore wall, distinct cross sections through the coiled polar tube, and increased density of the spore cytoplasm. As nascent spores mature and increase in number, the host cell becomes distended and ruptures. Spores are released into the surrounding tissues to infect neighboring cells. E. cuniculi is capable of infecting a variety of host organs, although the mechanisms by which dissemination to these sites occur are not clear. In naturally infected, immunocompetent mice, parasites and associated inflammation are usually observed in the CNS and kidneys. Spores are shed in the environment through urine to complete the life cycle. E. cuniculi is also easily propagated in tissue culture, resulting in release of fully infective spores from a variety of mammalian cell lines and generating sufficient numbers of parasites for in vitro work (Visvesvara 2002).
3.
541
PROTOZOA
Cell Biology
Microsporidia contain a curious mix of eukaryotic, prokaryotelike, and novel organelles. Like all eukaryotic organisms, they possess a membrane-bound nucleus, endoplasmic reticulum, Golgi apparatus, and membrane-bound vesicles. Although they lack mitochondria, a mitochondrial remnantmtermed
mitosomemhas been identified (Williams et al. 2002). The function of the mitosome is not clear, as microsporidia lack the mitochondrial enzymes associated with aerobic metabolism. All life stages of microsporidia are replete with ribosomes. These ribosomes are smaller than typical eukaryotic ribosomes, with a sedimentation rate and molecular organization similar to those found in prokaryotes. The defining criterion for inclusion into the phylum Microspora is possession of the polar tube and its associated structures. Also referred to as a polar filament, this unique apparatus is the means by which parasites invade and infect susceptible host cells. The polar tube is preformed and coiled around the sporoplasm within mature microsporidial spores. This tube is attached at the anterior end of the spore by the means of an anchoring disk and is surrounded by lamellar and vesicular membranes (also referred to as the polaroplast). At the opposite end of the spore is a membrane-bound posterior vacuole. After activation by some yet-to-be-determined signal (or signals), the polar tube is forcefully ejected from the spore, uncoiling and penetrating a susceptible host cell in the process. Once fully ejected, the spore's sporoplasm (containing the nucleus and abundant ribosomes) travels through the tube and is deposited within the host-cell cytoplasm. The function of the polaroplast is not clear, but it may contribute to the limiting membrane of the discharged sporoplasm (Weidner et al. 1984). It is also not clear what role the posterior vacuole plays. However, this vacuole is seen to swell by electron microscopy during the course of polar tube extrusion and may provide additional osmotic force to propel the sporoplasm through the polar tube and into the host cytoplasm (Lom and Vavra 1963). Although the mechanisms behind this sequence of events are still being worked out, it appears that calcium signaling and water flux across the spore wall play key roles in the activation of this novel parasite invasion apparatus (Frixione et al. 1997; Weidner and Byrd 1982). 4.
Disease and Diagnosis
E. cuniculi does not appear to be transmitted vertically in mice, nor are neonates more susceptible to infection than adults (Liu et al. 1988). Infection in immunocompetent mice is subclinical, although susceptibility to infection will vary by strain. C57BL/6, DBA, and 129/J mice had a higher parasite burden and depressed antibody responses to intraperitoneal inoculation with E. cuniculi when compared with BALB/c mice (Niederkorn et al. 1981). In immunologically impaired or immunosuppressed mice, however, overt disease may be seen. Naturally infected, immunocompetent mice treated twice weekly with corticosteroids developed abdominal distention and ascites. Microscopic examination of ascites fluid, peripheral blood smears, and brain, liver, spleen, and kidneys tissues revealed clusters of intracellular E. cuniculi organisms after 5 weeks of this immunosuppressive regime (Bismanis 1969). Experimentally infected C 5 7 B L / 6 - H f h l l nu mice develop ascites,
542
or a chronic wasting syndrome characterized by lethargy, anorexia, dehydration, and death within 2 to 4 weeks of parasite inoculation (Didier et al. 1994). Diagnosis of encephalitozoonosis is made by serology, histopathologic examination of tissues, and PCR. Serologic screening is a routine and useful method for monitoring E. cuniculi in immunocompetent mouse colonies. Histologically, parasites may be observed in a variety of tissues, but lesions are most frequently identified in the brain and kidneys. In contrast to the lesions seen in the CNS of endemically E. cuniculi-infected rabbits, those present in mice are nongranulomatous. In infected immunocompetent mice, a diffuse, nonsuppurative meningoencephalitis, and multifocal aggregates of activated microglia, lymphocytes, and macrophages, often with a perivascular distribution, are seen in the CNS. In addition, diffuse lymphocytic interstitial nephritis is observed. In these mice, parasites are scarce and may be difficult to identify. In immunosuppressed mice, E. cuniculi infection tends to be extensive and disseminated. Multiple necrotic foci with abundant intralesional and extralesional parasites have been described in the brain, heart, lungs, liver, spleen, adrenals, kidneys, pancreas, intestine, and the visceral and parietal peritoneal tissues of mice (Didier et al. 1994). Parasites are best observed in tissue sections with special stains (Fig. 21-7A, B). E. cuniculi spores stain positive with a Brown and Brenn's (B&B) tissue Gram stain, and bright red with PAS stain (Wasson and Peper 2000). Mature spore forms pick up stain the best--proliferative forms of the parasite are
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WASSON
more difficult to identify. Spores are rod-shaped and measure 1.5 x 2.5 ~tm. With B&B, spores often appear to have a clear vacuole at one end. With PAS, this vacuole appears as a deeply red staining granule. Other internal organelles are not visible by light microscopy. On occasion, it may be necessary to discriminate E. cuniculi from T. gondii infection, particularly in mice with CNS lesions. While T. gondii stains PAS positive, it does not stain with B&B. In addition, intracellular clusters of E. cuniculi spores can be distinguished from bradyzoitefilled T. gondii cysts by the presence of a cyst wall in the latter. Cyst walls stain faintly PAS positive and are argyrophilic (Frenkel 1956). Urinalysis for detection of E. cuniculi spores is theoretically possible but is more often used to monitor parasite shedding in experimentally infected mice. Molecular-based methods are available for diagnosis of encephalitozoonosis and may be useful when dealing with immunosuppressed mice and/or inconclusive histopathologic results (Didier et al. 1995). 5. Prevention, Treatment, Control
Because of its subclinical nature and insidious effect on animal-based biomedical research, E. cuniculi-infected colonies should be depopulated and repopulated with clean stock. Alternatively, rederivation can be performed. Although E. cuniculi is not as prevalent in contemporary mouse colonies as in the earlier part of the twentieth century, investigators, mouse biologists, and diagnosticians should remain vigilant.
Fig. 21-7 Encephalitozooncuniculi in the kidneyof a mouse. Clusters of spores are present within intracytoplasmicparasitophorousvacuoles. (A) H&E. (B) B&B Gram stain. A and B, bar = 50 t.tm.
2 1.
PROTOZOA
E. cuniculi infects a variety of cell types and animal species,
and may inadvertently be transmitted through the use of contaminated biologics in mice. Despite the handful of reports suggesting clinical improvement with the use of albendazole in humans with E. cuniculi infection, therapeutic efficacy has not been demonstrated in mice (De Groote et al. 1995b; Fournier et al. 2000; Koudela et al. 1994). Although E. cuniculi infection in humans has been documented, direct zoonotic transmission from animals to humans has not been demonstrated (Glaser et al. 1994). Human infections are likely obtained from water sources or the environment. In laboratory environments, Encephalitozoon spp. have been shown to be inactivated by 2% Lysol, 10% formalin, 70% ethanol, 10% bleach, quaternary ammonium compounds, and by boiling (Santillana-Hayat et al. 2002; Shadduck and Polley 1978). For this reason, steam sterilization of equipment and contaminated animal bedding, as well as routine disinfection of surfaces, should prevent environmental buildup of E. cuniculi spores in animal facilities. 6.
Research Implications
As alluded to in the introduction, interference with research results due to subclinical E. cuniculi infection is well documented in rodents. The inflammatory lesions commonly observed in the CNS of infected mice were often misinterpreted as lesions due to experimental inoculation with rabies virus, herpes simplex, cercopithecine herpesvirus 1, poliomyelitis virus, hepatitis B virus, syphilis, psittacosis, scrub typhus, and toxoplasmosis (reviewed in Wasson and Peper 2000). In addition to infectious disease work, E. cuniculi also interfered with early cancer research. Rats were found to be resistant to tumor development when inoculated with E. cuniculi-infected rat sarcoma cells (Petri 1965). E. cuniculi-infected mice were shown to have significantly reduced tumor growth and prolonged survival rates compared in uninfected controls when inoculated with different strains of solid or ascites-producing murine tumor cell lines (Arison et al. 1966; Niederkorn 1985). Nonspecific resistance to tumor growth in these mice may in part be due to enhanced natural killer cell activity--and associated increase in gamma interferon (INF-y) production--secondary to E. cuniculi infection (Niederkorn et al. 1983). Experimentally, E. cuniculi-infected C57BL/6 mice cleared lymphoma tumor cells from the lungs faster when compared to natural killer cell-deficient beige mice and infected, cyclophosphamide-treated C57BL/6 mice (both of which have reduced levels of INF-y). In addition, infected C57BL/6 mice were significantly more resistant to pulmonary tumor formation with B 16F10 melanoma cells when compared with uninfected, age-matched controls (Niederkorn 1985). The immune response to E. cuniculi infection in mice is now being elucidated from these early observations of enhanced tumor resistance. The importance of INF-~t was reaffirmed when INF-y depleted and INF-y knockout mice were shown to be highly susceptible to E. cuniculi
543
infection when compared to unmanipulated or wild-type controls (Khan and Moretto 1999). INF-y production by natural killer cells and gamma delta T lymphocytes is thought to activate CD8+ cytotoxic T lymphocytes, which in turn results in parasite killing. Evidence for this was shown in CD8 knockout mice, which were more susceptible to E. cuniculi infection compared to CD4 knockout mice and wild-type controls (Khan et al. 1999; Moretto et al. 2000).
ACKNOWLEDGMENTS This work was supported in part by NIH grant AI49735. The author wishes to thank Drs. Stephen Barthold, Nicole Baumgarth, Howard Gelberg, Stefan Tunev, and Mr. Robert Munn, for translation of articles from German and Russian, and for assistance with histopathology antd figures.
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