CRYPTOSPORIDIUM HOMINIS: EXPERIMENTAL CHALLENGE OF HEALTHY ADULTS

Am. J. Trop. Med. Hyg., 75(5), 2006, pp. 851 857 Copyright 2006 by The American Society of Tropical Medicine and Hygiene CRYPTOSPORIDIUM HOMINIS: EXPERIMENTAL CHALLENGE OF HEALTHY ADULTS CYNTHIA L. CHAPPELL,* PABLO C. OKHUYSEN, REBECCA LANGER-CURRY, GIOVANNI WIDMER, DONNA E. AKIYOSHI, SULTAN TANRIVERDI, AND SAUL TZIPORI The University of Texas Health Science Center at Houston School of Public Health and Medical School, Houston,Texas: Tufts University Cummings School of Veterinary Medicine, North Grafton, Massachusetts Abstract. Cryptosporidium hominis causes diarrhea in humans and has been associated with community outbreaks. This study describes the infectivity, illness, and serologic response after experimental challenge of 21 healthy adult volunteers with 10 500 C. hominis (TU502) oocysts. Sixteen subjects (76.2%) had evidence of infection; the 50% infectious dose (ID 50 ) was estimated to be 10 83 oocysts using clinical and microbiologic definitions of infection, respectively. Diarrhea occurred in 40% of subjects receiving 10 oocysts with a stepwise increase to 75% in those receiving 500 oocysts. A serum IgG response was seen in those receiving more than 30 oocysts. Greatest responses were seen in volunteers with diarrhea and oocyst shedding. Volunteers with no evidence of infection had indeterminant or negative IgG responses. Cryptosporidium hominis is infectious for healthy adults (ID 50 10 oocysts) and is clinically similar to C. parvum-induced illness. In contrast to C. parvum, C. hominis elicted a serum IgG response in most infected persons. INTRODUCTION * Address correspondence to Cynthia L. Chappell, The University of Texas Houston School of Public Health, 1200 Herman Pressler Street, Suite 118A, Houston, TX 77030. E-mail: Cynthia.L. Chappell@uth.tmc.edu 851 Molecular techniques have shown that two major species, Cryptosporidium parvum and C. hominis, are responsible for most human cases of cryptosporidiosis. 1,2 Cryptosporidium hominis infections are transmitted directly or indirectly from person to person, but have also been found on rare occasions in animals. 3 5 In addition, C. hominis has been experimentally transmitted to ruminants, and sensitive polymerase chain reaction (PCR) tests have detected C. hominis subpopulations in humans and animals excreting C. parvum oocysts. 6,7 In contrast, C. parvum is transmitted zoonotically and has been identified in multiple mammalian species, including cattle, horses, sheep, goats, and other domesticated and wild species. 8 Serologic studies indicate that 25% of the U.S. population have been exposed to Cryptosporidium. 9 18 The number is even higher in developing countries or in areas with poor sanitation and drinking water quality. 11 15 Outbreaks of diarrhea from C. hominis or C. parvum are typically associated with contaminated recreational or drinking waters. 16 Both Cryptosporidium species have caused community outbreaks of diarrhea, but urban populations are more often infected with C. hominis.. 17 19 Also, C. hominis appears to predominate in most studies of persons infected with human immunodeficiency virus. 20,21 Over the past few years, experimental studies of C. parvum in healthy adults have contributed important information regarding infectivity (50% infectious dose [ID 50 ]), natural history of the disease, and host immune response. 22 26 Overall, the onset of diarrhea in the volunteers typically occurred between days 4 and 7 post-challenge and lasted for another 4 7 days. Oocyst shedding usually began on days 6 8 postchallenge, lasted for 3 8 days, and often continued for a few days after diarrhea resolved. In contrast, the infectious dose (ID 50 ) among four isolates varied from 9 to 1,042 oocysts. In further studies, volunteers who had pre-existing serum antibody to Cryptosporidium antigens showed a relative resistance to re-infection when challenged with the homologous isolate (Iowa). In this population, ID 50 s were approximately 20-fold higher than in those subjects without pre-existing specific antibodies. Since knowledge of C. hominis infections has been limited to case reports and outbreak situations, little information exists regarding C. hominis infectivity and illness, particularly in immunocompetent hosts. Furthermore, serologic response to C. hominis has not been previously studied. Thus, this experimental challenge with C. hominis oocysts is the first study to examine infectivity, illness, and the serum IgG response to homologous antigens in healthy individuals. These data are important not only to advance our understanding of C. hominis pathogenicity, but also to provide essential information for risk assessment and protection of the drinking water supply. MATERIALS AND METHODS Study population. The study population consisted of healthy adults of both sexes and all races (age range 18 50 years) recruited by advertising in the Texas Medical Center and in Houston newspapers. This study was done following inclusion and exclusion criteria and study-related procedures as previously described. 25,26 After informed consent was obtained, volunteers agreeing to participate in the study were initially serologically screened for the presence of antibodies to Cryptosporidium. Volunteers who were negative for specific IgG underwent a thorough medical examination to identify any abnormalities that might be present. The challenge study was carried out in the University Clinical Research Center at the Memorial-Hermann Hospital (Houston, TX). The study was reviewed and approved by The University of Texas Health Science Center at Houston Committee for the Protection of Human Subjects. Informed consent was obtained from all subjects prior to the initiation of the study. Serologic testing. Blood was collected prior to challenge and at days 5, 10, 30, and 45 post-oocyst challenge. Sera were separated by centrifugation, tested for pre-challenge antibodies, and stored at 80 C. Antibodies to Cryptosporidium were detected by enzyme-linked immunosorbent assay using dis-

852 CHAPPELL AND OTHERS rupted C. parvum or C. hominis oocysts as previously described. 25 Cryptosporidium parvum antigens were used to assess pre-challenge antibodies to Cryptosporidium in volunteers. Known positive and known negative control sera were run on each microtiter plate. Positive sera were defined as those with a mean absorbance (414 nm) 1.5 times the negative control (mean optical density [OD] 0.115). Cryptosporidium hominis antigens were used to examine the IgG response at days 0, 5, 10, 30, and 45. In each case, negative and positive control sera (to C. parvum antigens) were included in each microtiter plate. All tests were done in duplicate. Negative and positive control sera yielded mean ODs of 0.143 and 0.262, respectively. For each subject, the mean absorbance at day 0 was subtracted from the peak postchallenge absorbance (day 30 or day 45) and expressed as the change in OD ( OD). The OD was plotted for each volunteer, and three groupings were apparent. Lowest ODs (0.0 0.054) were considered negative, mid-group ODs (0.079 0.113) were considered indeterminate, and highest ODs ( 0.147) were considered positive. Oocyst propagation and isolation. Cryptosporidium hominis oocysts (TU502) were originally isolated from a child with cryptosporidiosis and were propagated in the gnotobiotic piglet model. 27 Oocysts produced in this model were purified by the ether/nycodenz method with a final purification step using the micro-scale cesium chloride gradient technique described by Widmer and others, 20 Akiyoshi and others, 28 and Arrowood and Donaldson. 29 Purified oocysts were placed in 2.5% potassium dichromate for shipment to The University of Texas School of Public Health in Houston. Oocyst excystation and viability were determined within 48 hours of volunteer challenge. Excystation assays were carried out as previously described. 30 Viability of oocysts was determined with a Baclight Viability Kit (Molecular Probes Inc., Eugene, OR) as directed by the manufacturer. Oocyst preparation and delivery to volunteers has been described elsewhere. 24 Oocyst suspensions were subjected to serial dilution with phosphatebuffered saline, and replicate hemacytometer counts (n 6) were done to estimate the number of oocysts per unit volume. Once the desired concentration was reached, a 10- L aliquot of the suspended oocysts was removed and instilled into gelatin powder contained in a capsule. The capsules were delivered to volunteers and ingested within one hour of preparation. TU502 oocysts isolated from the original source as well as those passaged in the gnotobiotic pig were genotyped using a restriction fragment length polymorphism located in the oocyst wall protein gene (COWP) as previously described. 31,32 Subsequent passages in the gnotobiotic pig and stool samples from challenged volunteers were also tested in the same fashion. In addition, oocyst samples were genotyped with the species-specific PCR marker Lib13 and the Cp492 and Cp358 microsatellite markers. 7,33,34 Monitoring of volunteers. Volunteers were monitored as described. 26 Briefly, each volunteer was examined daily for the first 14 days after challenge. A personal diary, which documented the number and time of stool passage and any gastrointestinal symptoms that may have occurred, was kept by each participant and audited daily by the nursing staff. Also, stool samples collected during the previous 24 hours were delivered to the laboratory for analysis. After the first two weeks of the study, volunteers were examined as above three times per week for four additional weeks and asked to provide at least two 24-hour stool samples per week. Detection of oocyst shedding. Fecal specimens were collected from volunteers throughout the six-week study period and held at 4 C for no more than 24 hours prior to transfer to the laboratory. Specimens were then tested in duplicate for the presence of oocysts antigens using a commercially available enzyme immunoassay (EIA) kit as described by the manufacturer (ProspecT Cryptosporidium microplate assay; Alexon-Trend, Ramsey MN). All specimens that were positive by EIA were quantified by immunofluorescent assay (IFA) (Merifluor C/G; Meridian Bioscience, Inc., Cincinnati, OH) as previously described. 26 Definitions of infection and illness. Illness attack rate was defined as the number of cases of diarrhea divided by the number of volunteers who were exposed. Infection was confirmed when fecal oocysts were detected by EIA, IFA, or both at 36 hours post-challenge. Criteria for diarrhea included passage of 200 g of unformed stool per day, 3 unformed stools in eight hours, or 4 unformed stool in 24 hours. Symptoms included 2 concurrent gastrointestinal complaints (such as abdominal pain/cramps, tenesmus, gas, nausea, vomiting, fecal urgency, or fecal incontinence) in the context of at least one unformed stool. Duration of diarrhea was measured as previously described. 23 Cryptosporidiosis was defined as diarrhea in addition to 1 gastrointestinal symptoms with or without demonstrated oocysts within 30 days post-challenge. Analysis of data. The oocyst dose sufficient to infect 50% of susceptible persons (ID 50 ) was estimated using the cumulative endpoint method. 35 Kruskal-Wallis analysis of variance (ANOVA) with Dunn s multiple comparison tests were used to compare the IgG response ( OD) among clinical outcome groups. Analysis of variance with Welch s correction was used to compare outcome values for onset, duration, and severity of illness between C. hominis and C. parvum isolates. A P value < 0.05 was considered statistically significant. Data were analyzed using Instat software (GraphPad Software Inc., San Diego, CA). RESULTS Description of study population. Fifteen women (71.4%) and six (28.6%) men ranging in age from 19 to 50 years (mean 32.9 years, median 34 years) were enrolled in the study. Ethnicity of the volunteers was 12 Black (57.1%), 8 White (38.1%), and 1 (4.8%) Hispanic. Stability and delivery of C. hominis oocysts. Five batches of TU502 were produced in gnotobiotic pigs and used in volunteer challenge studies. All challenge doses were given within six weeks of oocyst production. At the time of volunteer challenge, oocyst excystation rates and viability ranged from 67 83% and 66 87%, respectively. Intended (actual ± SD) doses were 10 (10.3 ± 5.1), 30 (32.6 ± 6.5), 100 (105.3 ± 13.5), and 500 oocysts (500.8 ± 34.8). The coefficient of variations in the doses were 49.5%, 19.6%, 12.8%, and 6.9%, respectively. Genotypic analysis. TU502 oocysts passaged in pigs and oocysts excreted by four volunteers were genotyped using one restriction fragment length polymorphism marker (COWP), one species-specific PCR assay (Lib13), and two microsatellite length polymorphisms (Cp492 and Cp358) (Figure 1). These analyses confirmed that volunteers were excreting C.

C. HOMINIS IN HEALTHY ADULTS 853 FIGURE 1. Genotypic analysis of Cryptosporidium hominis (TU502). Lib13 (C. hominis specific, top) and Cp492 (bottom) amplicons were amplified from oocysts used for volunteer challenge (E numbers) and oocysts excreted by volunteers (T numbers) and visualized on native polyacrylamide gels. Each number represents a separate batch of inoculum or volunteer. Lane M shows molecular mass markers. Values on the right are in basepairs. Amplicons from C. parvum isolate MD 22 and TU502 were obtained in parallel as controls. neg. negative control polymerase chain reaction with no DNA added. hominis oocysts and showed no changes in the parasite population after pig-to-human passage. The Lib13 amplicons were obtained using the C. hominis-specific primers for three of four volunteer samples (Figure 1, top). Control C. parvum and C. hominis samples were tested in parallel and the expected negative and positive amplification, respectively, was observed. The same Cp358 amplicon was amplified from four pigs and three human samples, but this marker was less informative because the same allele was also found in two C. parvum controls. The Cp492 amplicon was amplified from three volunteer and four pig samples and displayed the same allele diagnostic for C. hominis, which was different from that obtained from the C. parvum control isolate Moredun (MD) (Figure 1, bottom). One volunteer sample failed to amplify with Lib13, Cp358, and Cp492. Infectivity of C. hominis oocysts. Only those persons who were IgG negative for antibodies to Cryptosporidium were enrolled in the study. Of the 21 volunteers who received a challenge dose, 13 developed diarrhea, and 9 had oocysts detected in fecal samples (Table 1). All nine oocyst-positive subjects had diarrhea and/or additional gastrointestinal symptoms. In contrast, 7 of 13 subjects with diarrhea and/or GI symptoms had no detectable oocysts. Five other volunteers had no evidence of infection. None of the volunteers who experienced a diarrhea had evidence of other enteric pathogens, despite having a complete microbiologic workup of all diarrheic stools. Dose-response curves were calculated using two different outcome variables: oocyst shedding with or without diarrhea (microbiologic definition) or diarrhea with or without detectable oocysts (clinical definition) (Figure 2). For oocystpositive persons (n 9), the estimated ID 50 was 83 oocysts compared with 10 oocysts using the clinical definition. Clinical outcomes of C. hominis infections. Diarrhea developed in 13 volunteers who ingested TU502 oocysts, yielding an illness attack rate of 61.9%. Three volunteers who did not meet criteria for diarrhea reported at least one unformed stool along with two or more additional gastrointestinal symptoms (Table 1). Six (66.7%) of the nine volunteers who had oocysts detected in their stools developed diarrhea. All volunteers received oral rehydration therapy when they developed diarrhea. One volunteer (no. 134) presented to the Clinical Research Center with mild dehydration, was rehydrated with intravenous fluids, and released after 15 hours of overnight observation. A dose-response relationship was seen in subjects that developed diarrhea. Those receiving lower doses were less likely to experience a diarrheal illness. The doses and percent of TABLE 1 Clinical, microbiologic, and serologic outcomes after healthy volunteers were challenged with a single dose of Cryptosporidium hominis (TU502) oocysts* Subject no. Intended dose Oocysts detected Enteric symptoms Diarrhea IgG response 147 10 + + 175 10 + + 176 10 + + 177 10 + + + 179 10 + 136 30 + + + 137 30 I 144 30 + + + 145 30 146 30 + + + + 132 100 + + + + 133 100 + + I 134 100 + + I 135 100 + + + + 150 100 I 151 100 + + 152 100 I 153 500 + + + 154 500 + + + 156 500 178 500 + + + + Total 21 9 14 13 8 * Positive (+), negative ( ), and indeterminant (I) outcomes are indicated. See Materials and Methods for definitions of each category.

854 CHAPPELL AND OTHERS FIGURE 2. Dose response for healthy adult volunteers experimentally challenged with a single dose of Cryptosporidium hominis oocytes. Data are presented using the cumulative endpoint method. 35 Dose response is shown for subjects with diarrhea in the presence or absence of detectable fecal oocytes (solid line) or subjects with confirmed infections (detected fecal oocytes, dashed line). volunteers with diarrhea were as follows:10, 40%; 30, 60%; 100, 71.4%; and 500, 75%. Interestingly and in contrast to previous studies done with C. parvum, asymptomatic shedding was not seen in any of the volunteers receiving C. hominis oocysts. 20 22 The incubation period for diarrhea ranged from 2 to 10 days after oocyst challenge with a mean ± SD and median of 5.4 ± 2.7 days and 4 days post-challenge, respectively (Table 2). The duration of diarrhea also varied approximately 10- fold with a range of 49 hours (2 days) to 518 hours (21.6 days). Mean ± SD and median duration were 137.3 ± 142.3 (5.7 days) and 75 hours (3.1 days), respectively. Of note, however, three subjects experienced a symptomatic episode of 9, 13, or 21 days. Severity of illness was evaluated by the number of unformed stools and the total unformed stool weight per diarrheal episode. The mean ± SD number of unformed stools was 8.9 ± 5.0 (median 9), and the total stool weight was 1.08 ± 0.72 (median 0.86) kilograms. The mean ± SD total number of unformed stools passed per day was 3.2 ± 1.0 and did not exceed five stools on any day. Infection and illness parameters from volunteers challenged with C. hominis were compared with similar experiments using four C. parvum isolates (Iowa, UCP, TAMU, and MD, ANOVA with Welch s correction). Onset, duration, or severity of diarrheal illness were not statistically different (P > 0.05) among isolates. Post-challenge serum IgG response. For each volunteer, the specific serum IgG response was assessed after challenge and compared with the pre-challenge value. Overall, baseline absorbance values had a mean ± SD of 0.183 ± 0.055. The kinetics of the response varied somewhat but all positive sera reached peak values by days 30 or 45. The response category (i.e., positive, indeterminant, or negative) for each volunteer was then grouped according to challenge dose (Table 3). None of the five volunteers receiving a challenge dose of 10 oocysts mounted a measurable serum IgG response despite the fact that all met the clinical definition of infection and four were microbiologically confirmed. Sixteen volunteers received higher oocyst doses: eight (50%) had a positive IgG response (all with diarrhea), five (45.5%) were indeterminant (three with no evidence of infection), and three (27.3%) were serum IgG negative (two with no evidence of infection). Furthermore, the OD for each volunteer was plotted against the clinical outcome (Figure 3). Since none of the five persons who were challenged with 10 oocysts showed an IgG response, these volunteers were not included in the analysis. Volunteers receiving 30 oocysts and who were asymptomatic with no detectable oocysts showed the lowest ODs (mean ± SD 0.056 ± 0.049). Those with diarrhea but no detectable oocysts were slightly more reactive with a mean ± SD OD of 0.148 ± 0.118. The highest values were seen in volunteers who had a diarrheal illness and detectable oocysts (0.220 ± 0.076). This latter category was significantly different in mean OD (P 0.014) than the no diarrhea, no oocysts category. DISCUSSION This study is the first report of experimental challenge with C. hominis (TU502) in healthy adult volunteers. The estimated ID 50 for TU502 was 10 83 oocysts, depending on whether the microbiologic or clinical definition of infection Subject no. TABLE 2 Clinical features of subjects with diarrhea following ingestion of Cryptosporidium hominis (TU502) oocysts Incubation period (day post-challenge) Duration of illness (hours) Maximal no. of unformed stools per day No. of unformed stools per illness Total unformed stool weight per episode of diarrhea (kg) 132 5 161 5 14 1.78 133 4 60 3 5 0.41 134 4 51 5 10 0.86 135 4 97 2 4 0.36 136 4 126 3 13 1.69 144 10 49 3 4 0.53 146 6 518 4 19 2.29 151 2 225 3 13 2.22 153 10 312 2 9 1.11 154 4 73 3 7 0.47 177 4 75 4 11 1.43 178 4 34 3 5 0.66 179 10 4 2 2 0.28 Mean ± SD 5.4 ± 2.7 137.3 ± 142.3 3.2 ± 1.0 8.9 ± 5.0 1.08 ± 0.72

C. HOMINIS IN HEALTHY ADULTS 855 TABLE 3 Serum IgG response to Cryptosporidium hominis following challenge with C. hominis oocytes* Oocyst dose N IgG positive Indeterminant IgG negative 10 5 0 0 5 30 5 3 1 1 100 7 2 4 1 500 4 3 0 1 * Responses were categorized as the following changes (day 0 versus peak) in optical density over the six-week study period: negative, 0.054; indeterminant, 0.079 0.113; positive, 0.147. Volunteer had no diarrhea, gastrointestinal (GI) symptoms, or detectable oocysts. Volunteer had diarrhea and GI symptoms, but no detectable oocysts. was used. In this study, consecutive stool samples from subjects with confirmed TU502 infections were not always positive, suggesting that the intensity of oocyst shedding was variable and sometimes below the detectable limit even on some days when diarrhea was present. Furthermore, although 13 of the volunteers developed a diarrheal illness characteristic of cryptosporidiosis, seven had no detectable fecal oocysts in any stool sample, suggesting that these persons had light infections that remained below the detectable limit. This phenomenon is comparable to several subjects in the C. parvum challenge studies. 26 In those earlier studies where cryptosporidiosis could not be confirmed by IFA or EIA, additional testing using a more sensitive flow cytometric method (detection limit 10 3 /ml) showed that a low level of oocysts were present in IFA-negative, diarrheic stools (Chappell CL and others, unpublished data). To address the possibility of diarrhea from other causes, all diarrheic stools were subjected to a complete microbiologic work-up, and no pathogens other than Cryptosporidium were detected. Furthermore, symptoms exhibited by volunteers did not include nausea and vomiting, as may be expected with viral gastroenteritis. Finally, the high degree of infectivity (i.e., low ID 50 ) associated with TU502 is comparable to the most infectious C. parvum isolates (TAMU and Iowa) used in other volunteer FIGURE 3. Serum IgG response to the homologous antigen in volunteers challenged with 30 500 Cryptosporidium hominis oocytes. Each points represents the change in optical density (OD) between day 0 and the peak day of response during a six-week period. P 0.014, by Kruskal-Wallis analysis of variance. studies. Since only one C. hominis isolate was tested, the potential variability in infectivity among isolates remains known. However, if C. hominis mimics C. parvum, significant phenotypic variability may be expected. This would be consistent with a similar level of genetic heterogenity observed in a geographically diverse collection of C. parvum and C. hominis isolates. 36 However, the occurrence and extent of this variation requires additional study. A recently described rodent model for C. hominis will enable a comparative study of genetically distinct C. hominis isolates. 37 To our knowledge, the C. hominis isolate used in the volunteer studies, is the only laboratory-maintained C. hominis isolate. This isolate was also selected for a recently completed genome sequencing project. 38 A dose-response relationship for diarrhea was evident in volunteers who were challenged with C. hominis oocysts. This was not observed in previous studies with C. parvum oocysts. 22 24 Of note, however, an earlier study showed that enteric symptoms (including but not limited to diarrhea) were significantly (P 0.018) more common in volunteers receiving higher oocyst doses ( 500) of the Iowa isolate. 22 Cryptospordium hominis circulates in human populations and has not been associated with zoonotic transmission. However, the gnotobiotic pig is susceptible to C. hominis infection and was especially useful for these studies given the relative ease of oocyst purification in the absence of bacterial flora in the animal s gut. Although fewer oocysts were produced in comparison to C. parvum in calves, the number of purified oocysts derived from the gnotobiotic pig was sufficient for the described studies. We did, however, note important differences in the stability of the purified oocysts in storage. In past studies, C. parvum oocysts were kept in 2.5% potassium dichromate for at least three months without exhibiting significant loss of viability. In those studies, it was a simple matter to maintain a viability 80% prior to volunteer challenge. In contrast, C. hominis oocysts were less stable at room temperature, showing an accelerated decrease in oocyst survival compared with C. parvum oocysts, an observation consistent with earlier findings. 32 Therefore, excystation rates of oocysts delivered to volunteers were between 67% and 80% at the time of volunteer challenge depending on the oocyst batch. The infectivity estimates reported herein do not include any corrections for the lower excystation rate, and thus the reported ID 50 may be slightly overestimated. Little is known regarding the initiation of the serologic response to Cryptosporidium and whether the predominant antigenic stimulus is delivered with the challenge dose, during the replication process, or both. Previous studies with a C. parvum isolate (Iowa) indicated that subjects failed to mount a serum IgG response after primary challenge, but 33% did after rechallenge one year later. 39 Furthermore, oocyst challenge in volunteers with pre-existing specific serum IgG resulted in an anamnestic response. 25 In contrast to C. parvum (Iowa isolate), C. hominis resulted in a serologic response in 8 (38.1%) of 21 challenged volunteers and in 8 (53.3%) of 15 who had evidence of infection. Interestingly, only volunteers receiving 30 oocysts had a serum IgG response, even though all had diarrhea or fecal oocyst shedding. Furthermore, the degree of response was influenced by postchallenge outcome. Volunteers who had a diarrheal illness and who shed detectable levels of oocysts yielded the highest responses. Thus, data from this study suggest that the Cryp-

856 CHAPPELL AND OTHERS tosporidium species used in the challenge as well as the challenge dose and post-challenge events are important contributors to the overall serum IgG response. These observations, however, need to be confirmed with a larger population of subjects and with other C. hominis isolates before broad generalizations can be made. In summary, C. hominis oocysts are capable of causing infection and illness in healthy adults similar to that seen with C. parvum. The ID 50 of TU502 is in the low range compared with that of C. parvum isolates. However, it is unclear whether the TU502 isolate is representative of C. hominis isolates circulating in human populations. The data generated from the C. hominis dose response studies adds to the growing body of data regarding Cryptosporidium infectivity in immunocompetent humans and provides important and valuable infectivity estimates for use in risk assessment and the setting of water quality standards. 40 Received May 23, 2006. Accepted for publication July 1, 2006. Acknowledgments: We express our gratitude to the many individuals who contributed to these studies: the participating volunteers; Nai- Hui Chiu, Madeline Ottosen, and the research nursing staff at the University Clinical Research Center for their expertise and attention to the many details of the study; and to Philip Lupo, Audrey Wanger, and Zhi Dong Jiang for excellent technical assistance. Data from this study, in part, have been presented at the annual meeting of the American Society of Parasitologists. August 1 5, 2003, Halifax, Nova Scotia and the American Society of Tropical Medicine and Hygiene, November 7 11, 2004, Miami, Florida, Abstract no. 35. Financial support: This study was supported, in part, by the National Center for Environmental Research STAR Program of the Environmental Protection Agency (grant no. GR828035-01-0 to Cynthia L. Chappell), the National Institutes of Health General Clinical Research Centers (grant no. RR-02558), and the National Institute of Allergy and Infectious Diseases, (grant no. AI52781 to Giovanni Widmer and grant no. NO1-AI-25466 to Saul Tzipori). Authors addresses: Cynthia L. Chappell, University of Texas- Houston School of Public Health, 1200 Herman Pressler Street, Suite 118A Houston, TX 77030, Telephone: 713-500-9026, Fax: 713-500- 9020, E-mail: Cynthia.L.Chappell@uth.tmc.edu. Pablo C. Okhuysen, Division of Infectious Diseases, University of Texas Medical School, 6431 Fannin, Room 2.112, Houston, TX 77030, E-mail: Pablo.C.Okhuysen@uth.tmc.edu. Rebecca Langer-Curry, Office of Environmental Safety, Baylor College of Medicine, Room K104, 1 Baylor Plaza, Houston, TX 77030, E-mail: langercu@bcm.tmc.edu. Giovanni Widmer, Donna E. Akiyoshi, Sultan Tanriverdi, and Saul Tzipori, Division of Infectious Diseases, Tufts Cummings School of Veterinary Medicine, North Grafton, MA 01536, E-mails: giovanni.widmer@tufts.edu, donna.akiyoshi@tufts.edu, sultan.tanriverdi@tufts.edu, and saul.tzipori@tufts.edu. REFERENCES 1. Morgan UM, Constantine CC, Forbes DA, Thompson RCA, 1997. Differentiation between human and animal isolates of Cryptosporidium parvum using rdna sequencing and direct PCR analysis. J Parasitol 83: 825 830. 2. Morgan-Ryan UM, Fall A, Ward LA, Hijjawi N, Sulaiman I, Fayer R, Tompson RC, Olson M, Lal A, Xiao L, 2002. Cryptosporidium hominis n. sp. (Apicomplexa: Cryptosporidiidae) from Homo sapiens. J Euk Microbiol 49: 433 440. 3. Morgan UM, Xiao L, Hill BD, O Donoghue P, Limor J, Lal A, Thompson RCA, 2000. Detection of the Cryptosporidium parvum human genotype in a dugong (Dugong dugon). J Parasitol 86: 1352 1354. 4. Smith RA, Nichols B, Mallon M, Macleod A, Tait A, Reilly WJ, Browning LM, Gray D, Reid SW, Wastling JM, 2005. Natural Cryptosporidium hominis infections in Scottish cattle. Vet Rec 156: 710 711. 5. Ryan UM, Bath C, Robertson I, Read C, Elliot A, McInnes L, Traub R, Besier B, 2005. Sheep may not be an important zoonotic reservoir for Cryptosporidium and Giardia parasites. Appl Environ Microbiol 71: 4992 4997. 6. Giles M, Webster KA, Marshall JA, Catchpole J, Goddard TM, 2001. Experimental infection of a lamb with Cryptosporidium parvum genotype 1. Vet Rec 149: 523 525. 7. Tanriverdi S, Arslan MO, Akiyoshi DE, Tzipori S, Widmer G, 2003. Identification of genotypically mixed Cryptosporidium parvum populations in humans and calves. Mol Biochem Parasitol 130: 13 22. 8. O Donoghue P, 1995. Cryptosporidium and cryptosporidiosis in man and animals. Int J Parasitol 25: 139 195. 9. Ungar BLP, 1990. Cryptosporidiosis in humans (Homo sapiens). R. Fayer, ed. Cryptosporidium and Cryptosporidiosis. Boca Raton, FL: CRC Press, 59 82. 10. Frost FJ, Muller TB, Calderon RL, Craun GF, 2004. Analysis of serological responses to Cryptosporidium antigen among NHANES III participants. Ann Epidemiol 14: 473 478. 11. Casemore DP, Wright SE, Coop RL, 1990. Cryptosporidiosis: human and animal epidemiology. R. Fayer, ed. Cryptosporidium and Cryptosporidiosis. Boca Raton, FL: CRC Press, 66 92. 12. Leach CT, Koo FC, Kuhls TL, Hilsenbeck SG, Jenson HB, 2000. Prevalence of Cryptosporidium parvum infection in children along the Texas-Mexico border and associated risk factors. Am J Trop Med Hyg 62: 656 661. 13. Frost FJ, Fea E, Gilli G, Biorci F, Muller TM, Craun GF, Calderon RL, 2000. Serological evidence of Cryptosporidium infections in southern Europe. Eur J Epidemiol 16: 385 390. 14. Miron D, Colodner R, Kenes Y, 2000. Age-related seroprevalence of Cryptosporidium in northern Israel. Isr Med Assoc J 2: 343 345. 15. Tumwine JK, Kekitiinwa A, Nabukeera N, Akiyoshi D, Rich S, Widmer G, Feng X, Tzipori S, 2003. Cryptosporidium parvum among children with diarrhea in Mulago Hospital, Uganda. Am J Trop Med Hyg 68: 710 715. 16. Hlavsa MC, Watson JC, Beach MJ, 2005. Cryptosporidiosis surveillance United States, 1999 2002. MMWR Morb Mortal Wkly Rep 54: 1 8. 17. Ong CSL, Eisler DL, Goh SH, Tomblin J, Awad-El-Kariem FM, Beard CB, Xiao L, Sulaiman I, Lal A, Fyfe M, King A, Bowie WR, Isaac-Renton J, 1999. Molecular epidemiology of cryptosporidiosis outbreaks and transmission in British Colombia, Canada. Am J Trop Med Hyg 61: 63 69. 18. McLauchlin J, Amar C, Pedraza-Diaz S, Nichols GL, 2000. Molecular epidemiological analysis of Cryptosporidium spp. in the United Kingdom: results of genotyping Cryptosporidium spp. in 1705 fecal samples from humans and 105 fecal samples from livestock animals. J Clin Microbiol 38: 3984 3990. 19. Learmonth JL, Ionas G, Ebett KA, Kwan ES, 2004. Genetic characterization and transmission cycles of Cryptosporidium species isolated from human in New Zealand. Appl Environ Microbiol 70: 3973 3978. 20. Widmer G, Tzipori S, Fichtenbaum CJ, Griffiths JK, 1998. Genotypic and phenotypic characterization of Cryptosporidium parvum isolates from people with AIDS. J Infect Dis 178: 834 840. 21. Cama VA, Bern C, Sulaiman IM, Gilman RH, Ticona E, Vivar A, Kawai V, Vargas D, Zhou L, Xiao L, 2003. Cryptosporidium species and genotypes in HIV-positive patients in Lima, Peru. J Eukaryot Microbiol 50 (Suppl): 531 533. 22. DuPont HL, Chappell CL, Sterling CR, Okhuysen PC, Rose JB, Jakubowski W, 1995. The infectivity of Cryptosporidium parvum in healthy volunteers. N Engl J Med 30: 855 859. 23. Okhuysen PC, Chappell CL, Crabb JH, Sterling CR, DuPont HL, 1999. Virulence of three distinct Cryptosporidium parvum isolates for healthy adults. J Infect Dis 180: 1275 1281. 24. Okhuysen PC, Rich SM, Chappell CL, Grimes K, Widmer G, Feng X, Tzipori S, 2002. Infectivity of a Cryptosporidium parvum isolate of cervine origin for healthy adults and gamma interferon- knockout mice. J Infect Dis 185: 1320 1325. 25. Chappell CL, Okhuysen PC, Sterling CR, Wang C, Jakubowski W, DuPont HL, 1999. Infectivity of Cryptosporidium parvum in healthy adults with pre-existing anti-c. parvum serum immunoglobulin G. Am J Trop Med Hyg 60: 157 164.

C. HOMINIS IN HEALTHY ADULTS 857 26. Chappell CL, Okhuysen PC, White AC Jr, 2003. Cryptosporidium parvum: Infectivity, pathogenesis, and the host-parasite relationship. Thompson RCA, Armson A, Morgan-Ryan U, eds. Cryptosporidium: From Molecules to Disease. Amsterdam, The Netherlands, Elsevier B.V. 19 49. 27. Akiyoshi DE, Feng X, Buckholt MA, Widmer G, Tzipori S, 2002. Genetic analysis of a Cryptosporidium parvum human genotype 1 isolate passaged through different host species. Infect Immun 70: 5670 5675. 28. Akiyoshi DE, Dilo J, Pearson C, Chapman S, Tumwine J, Tzipori S, 2003. Characterization of Cryptosporidium meleagridis of human origin passaged through different host species. Infect Immun 71: 1828 1832. 29. Arrowood MJ, Donaldson K, 1996. Improved purification methods for calf-derived Cryptosporidium parvum oocysts using discontinuous sucrose and cesium chloride gradients. J Eukaryot Microbiol 43: 89S. 30. Woodmansee DB, 1987. Studies of in vitro excystation of Cryptosporidium parvum from calves. J Protozool 34: 398 402. 31. Spano F, Putignani L, McLauchlin J, Casemore DP, Crisanti A, 1997. PCR-RFLP analysis of the Cryptosporidium oocyst wall protein (COWP) gene discriminates between C. wrairi and C. parvum, and between C. parvum isolates of human and animal origin. FEMS Microbiol Lett 150: 209 217. 32. Widmer G, Akiyoshi D, Buckholt MA, Feng X, Rich SM, Deary KM, Bowman CA, Xu P, Wang Y, Wang X, Buck GA, Tzipori S, 2000. Animal propagation and genomic survey of a genotype 1 isolate of Cryptosporidium parvum. Mol Biochem Parasitol 108: 187 197. 33. Feng X, Rich SM, Tzipori S, Widmer G, 2002. Experimental evidence for genetic recombination in the opportunistic pathogen Cryptosporidium parvum. Mol Biochem Parasitol 119: 55 62. 34. Caccio S, Homan W, Camilli R, Traldi G, Kortbeek T, Pozio E, 2000. A microsatellite marker reveals population heterogeneity within human and animal genotypes of Cryptosporidium parvum. Parasitol 120: 237 244. 35. Reed LJ, Muench H, 1938. A simple method of estimating fifty per cent endpoints. Am J Hyg 27: 493 497. 36. Tanriverdi S, Widmer G, 2006. Differential evolution of repetitive sequences in Cryptosporidium parvum and Cryptosporidium hominis. Infect Genet Evol 6: 113 122. 37. Baishanbo A, Gargala G, Delaunay A, François A, Ballet J, Favennec L, 2005. Infectivity of Cryptosporidium hominis and Cryptosporidium parvum genotype 2 isolates in immunosuppressed mongolian gerbils. Infect Immun 73: 5252 5255. 38. Xu P, Widmer G, Wang Y, Ozaki LS, Alves JM, Serrano MG, Puiu D, Manque P, Akiyoshi D, Mackey AJ, Pearson WR, Dear PH, Bankier AT, Peterson DL, Abrahamsen MS, Kapur V, Tzipori S, Buck GA, 2004. The genome of Cryptosporidium hominis. Nature 431: 1107 1112. 39. Okhuysen PC, Chappell CL, Sterling CR, Jakubowski W, Du- Pont HL, 1998. Susceptibility and serologic response of healthy adults to reinfection with Cryptosporidium parvum. Infect Immun 66: 441 443. 40. Anonymous, 2002. EPA National Primary Drinking Water Regulations: Long Term 1 Enhanced Surface Water Treatment Rule Federal Register [CFR Parts 9, 141, and 142], 67 (940): 1811 1844.