Temporal Distribution and Parasite Load Kinetics in Blood and Tissues during Neospora caninum Infection in Mice
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感染与免疫杂志 2006年第4期
Departamento de Sanidad Animal, Facultad de Veterinaria, Universidad Complutense de Madrid, 28040 Madrid, Spain
ABSTRACT
The kinetics of Neospora caninum loads in mice inoculated with NC-Liv or NC-1 isolates were studied. The acute phase was characterized by parasitemia and the detection of parasite DNA in several organs, whereas during the chronic phase, the parasite was detected mainly in the brain. Mice infected with NC-Liv developed clinical signs, showing higher brain parasite burdens than NC-1-infected mice.
TEXT
Neospora caninum is considered one of the main agents implicated in bovine abortion (8). Mouse inoculation is the model most widely used to study aspects of N. caninum infection (3, 6, 10, 11, 12, 13, 14, 15) and offers a tool with which to characterize and compare the virulence of N. caninum isolates (1, 18). However, studies of parasite loads in blood and tissues during the acute and chronic phases of infection are lacking. Here we determined the organs that support parasite replication and persistence in a mouse model and the relationship between the parasite isolate and virulence. This knowledge will aid in evaluating the efficiency of drugs and vaccines.
Fifty-five 6-week-old female BALB/c mice (Harlan, Barcelona, Spain) were infected with 106 tachyzoites from NC-Liv (2) (group 1) or NC-1 (9) (group 2) isolates by intraperitoneal (i.p.) injection as described previously (6). A control group (group 3) was inoculated i.p. with phosphate-buffered saline. Three random animals from each group were sacrificed on days 1, 2, and 3 postinfection (p.i.), then every 2 days until day 21, and finally weekly up to day 63 p.i. All animal-handling procedures complied with EU legislation. Mice that appeared moribund were killed by CO2 inhalation, and samples were collected. Animals were monitored daily for the presence of clinical signs of neosporosis, and the time of the event (presence of clinical signs) was recorded, for 63 days. Evaluation of clinical signs was based on previous N. caninum infection studies, in which mice with clinical neosporosis can develop rough hair coats, inactivity, anorexia, and neurological signs consisting of head tilting, walking in circles, ataxia, pelvic limb weakness, and paralysis (1, 10, 13, 14). Blood samples (300 to 500 μl) were collected in EDTA tubes and centrifuged, and plasma was cryopreserved for enzyme-linked immunosorbent assays. Mesenteric, iliac, mediastinal, and cervical lymph nodes, spleens, livers, lungs, and brains were recovered and frozen. DNA was extracted from 107 NC-1 tachyzoites as a positive control and from 10 to 20 mg of murine tissues by using a GenomicPrep cell and tissue DNA isolation kit (Amersham Biosciences, Uppsala, Sweden). DNA from blood samples was extracted with a GenomicPrep blood DNA isolation kit (Amersham Pharmacia) by following the manufacturer's instructions. Parasite DNA in murine tissues was measured by real-time PCR using SYBR Green I. We used primer pairs from the N. caninum Nc-5 sequence to quantify parasites, and we used primers from the 28S rRNA gene to quantify host DNA (7). N. caninum-specific serum isotypes immunoglobulin G2a (IgG2a) and IgG1 were determined by an enzyme-linked immunosorbent assay using soluble N. caninum tachyzoite antigen (10, 15) (0.5 μg in 100 μl/well). We used diluted murine serum samples (1:100) and an anti-mouse IgG2a or IgG1 antibody (1:5,000; Southern Biotechnology, Birmingham, AL).
Differences between groups were analyzed using the Kruskal-Wallis test, the Mann-Whitney U test, and the Student t test. When statistically significant differences were found by the Kruskal-Wallis test, a nonparametric multiple-comparison test was applied to examine all possible pairwise comparisons. A P value of less than 0.05/[k x (k – 1)/2] was considered statistically significant, with k corresponding to the number of animals in the group. Frequencies of clinical signs of neosporosis in infected groups were compared by Fisher's exact test. In addition, the Kaplan-Meier survival method (4) was applied to estimate the portion of all individuals surviving (not showing clinical signs) at each time point (days p.i.). Data from subjects removed from the study (either because they had to be killed or because the study ended) were censored. Subsequently, the log rank statistical test for comparison of survival curves between infected groups was applied (5). Statistical analysis was carried out using SAS 8.02 (SAS Institute, Cary, NC) and GraphPad Prism 4, v. 4.03 (GraphPad, San Diego, CA), software.
None of the mice died spontaneously during the follow-up period. In infected groups, clinical signs of neosporosis were more often observed in group 1 (25.4% versus 1.8%; P < 0.01 by the Fisher F test). In the group infected with NC-Liv isolates, the numbers of mice showing clinical signs of neosporosis were four, six, three, and one on days 11, 21, 35, and 42 p.i., respectively, whereas only one mouse infected with NC-1 exhibited clinical neosporosis on day 21 p.i. (Fig. 1). Rough hair coats and apathy were the first signs observed, followed by anorexia, inactivity, and nervous signs (pelvic limb weakness, head tilting, and walking in circles). As shown in Fig. 1, the percentage of survival among NC-1-infected mice was significantly greater than that among NC-Liv-infected mice (P < 0.001 by the log rank test). Control mice remained clinically normal until the end of the experiment. Parasite DNA was detected in blood, spleens, and livers of infected mice from day 1 to days 5 to 7 p.i. The parasite load peaked on days 1 to 3 p.i. and decreased thereafter (P < 0.0001 by the Kruskal-Wallis H test; P < 0.01 by the nonparametric multiple-comparison test for days 1 to 3 versus 5 to 7 p.i. and for 5 to 7 versus >7 days p.i.) (Table 1; Fig. 2). Parasitemia was subsequently transient and very low, with sporadic detection on days 11, 13, 19, and 56 p.i. (Table 1). In the upper and lower lymph nodes, the parasite was detected intermittently at very low levels throughout the experiment (Table 1). N. caninum DNA was detected in lungs from day 1 to days 11 to 13 p.i. There was a higher parasite burden on days 1 to 3 that gradually declined until extinction around days 11 to 13 p.i. (P < 0.0001 by the Kruskal-Wallis H test; P < 0.008 by the nonparametric multiple-comparison test for days 1 to 3, 5 to 7, and 9 to 13 versus >13 days p.i.). In brains, parasites were detected from days 5 to 7 p.i. until the end of the experiment (day 63). The parasite burden reached a peak on day 13 p.i. (Fig. 2) and decreased gradually thereafter (P < 0.0001 by the Kruskal-Wallis H test; P < 0.003 by the nonparametric multiple-comparison test for days 9 to 13, 15 to 21, 28 to 35, and 42 to 49 versus days 1 to 7 p.i.). Comparison of parasite loads in organs of mice inoculated with the two isolates showed similar numbers (P > 0.05 by the Mann-Whitney U test) except in the liver and the brain. In the liver, NC-1-inoculated mice had significantly higher parasite loads on day 1 p.i. than NC-Liv-inoculated mice (P < 0.05), whereas in the brain, NC-Liv-inoculated mice had significantly higher parasite burdens than NC-1-inoculated mice on days 13, 19, and 56 p.i. (P < 0.05). Subsequently, data were also compared per isolate and group 1 produced a higher parasite burden in the brain (P < 0.05).
We compared brain parasite numbers in infected mice with and without clinical signs. The data showed significantly higher brain parasite burdens in samples from mice with clinical signs (14 mice inoculated with NC-Liv and 1 with NC-1) than in those with no clinical signs of disease (95 infected mice) (P < 0.05 by the Mann-Whitney U test).
Serum antibody response kinetics followed similar patterns in groups 1 and 2 (P > 0.05 by t test) (Fig. 3). When infected groups were compared with noninfected controls, significant increases in IgG2a and IgG1 levels were detected on days 7 and 9 p.i., respectively (P < 0.05 by t test). Mice had a mixed IgG2a/IgG1 response, with a slight predominance of IgG2a levels during the first 2 to 3 weeks p.i., followed by a tendency toward a IgG1 bias thereafter (P < 0.0001 by paired t tests).
In conclusion, we have characterized the temporal tissue distribution and parasite loads of N. caninum in a murine model. The course of N. caninum infection in mice could be separated into three phases. During the first phase (day 1 to days 5 to 7 p.i.), parasitemia was present and the peak of parasite burden in blood coincided with the maximum parasite load in the spleen, liver, and lungs. The second period (days 5 to 7 to days 11 to 13 p.i.) coincided with the development of host immunity; tachyzoites were cleared from host tissues and were detected only in the lungs and brain. The number of tachyzoites decreased in the lungs but increased in the brain. Infection became chronic in the third phase (day 13 onward); the parasite disappeared from the lungs, and the parasite load decreased in the brain and remained at low levels until the end of the experiment (day 63 p.i.). The parasite was detected intermittently at very low levels in lymph nodes throughout the experiment. In this study, we provide evidence that the major target organ in acute N. caninum infection was the lung, whereas in the chronic phase it was the brain, corroborating that this is the major site of parasite persistence. These results agree with the findings of previous N. caninum and Toxoplasma gondii mouse model experiments (10, 12, 16, 17, 19), in which the most frequently parasitized tissues were the lungs and brain.
The parasite distribution kinetics observed strongly suggested immune regulation of infection. During the chronic infection phase, N. caninum induced high levels of both isotypes, but IgG1 production predominated and the organism established a long-term chronic infection in the brain. As in other studies, disease susceptibility during chronic infection was associated with a mixed response characterized by a low gamma interferon/interleukin-4 ratio (15).
In N. caninum infection, the parasite isolate appears to have a significant role in the progression of infection (1, 18). Here, a larger number of NC-Liv-inoculated mice than of NC-1-inoculated mice showed clinical signs, with higher brain parasite burdens, and survival curve analysis showed a marked difference between isolates. Our results suggest that the NC-Liv isolate might actually have detrimental effects on morbidity, as reported previously (1). This study also demonstrates an association between brain parasite load and the presence of clinical signs. On the other hand, NC-1-inoculated mice had higher parasite loads in the liver on day 1 p.i. than mice inoculated with NC-Liv. Our observations suggest that N. caninum isolates may have different tissue tropisms depending on the infection phase. We underline the importance of quantifying the parasite burden in order to follow the course of infection and to determine differences in virulence among N. caninum isolates.
ACKNOWLEDGMENTS
We thank Ricardo García de la Mata for statistical analyses and Vanesa Navarro for excellent technical assistance. We gratefully acknowledge D. J. L. Williams (Liverpool School of Tropical Medicine, Liverpool, United Kingdom) for generously providing the NC-Liverpool isolate and Veronica Risco for valuable help. We express our gratitude to two anonymous referees for helpful comments on the manuscript.
E.C.-F. was financed by the Spanish Ministry of Science and Technology (M.I.T. fellowship, Spain). This work was supported by a grant from the Spanish Government (AGL2001-1362) and was part of the EU research collaboration COST-854.
REFERENCES
1. Atkinson, R., P. A. Harper, C. Ryce, D. A. Morrison, and J. T. Ellis. 1999. Comparison of the biological characteristics of two isolates of Neospora caninum. Parasitology 118:363-370.
2. Barber, J., A. J. Trees, M. Owen, and B. Tennant. 1993. Isolation of Neospora caninum from a British dog. Vet. Rec. 133:531-532.
3. Baszler, T. V., M. T. Long, T. F. McElwain, and B. A. Mathison. 1999. Interferon-gamma and interleukin-12 mediate protection to acute Neospora caninum infection in BALB/c mice. Int. J. Parasitol. 29:1635-1646.
4. Bland, J. M., and D. G. Altman. 1998. Survival probabilities. The Kaplan-Meier method. BMJ 317:1572.
5. Bland, J. M., and D. G. Altman. 2004. The logrank test. BMJ 328:1073.
6. Collantes-Fernandez, E., G. Alvarez-Garcia, V. Perez-Perez, J. Pereira-Bueno, and L. M. Ortega-Mora. 2004. Characterization of pathology and parasite load in outbred and inbred mouse models of chronic Neospora caninum infection. J. Parasitol. 90:579-583.
7. Collantes-Fernandez, E., A. Zaballos, G. Alvarez-Garcia, and L. M. Ortega-Mora. 2002. Quantitative detection of Neospora caninum in bovine aborted fetuses and experimentally infected mice by real-time PCR. J. Clin. Microbiol. 40:1194-1198.
8. Dubey, J. P. 2003. Neosporosis in cattle. J. Parasitol. 89(Suppl.):S42-S56.
9. Dubey, J. P., J. L. Carpenter, C. A. Speer, M. J. Topper, and A. Uggla. 1988. Newly recognized fatal protozoan disease of dogs. J. Am. Vet. Med. Assoc. 192:1269-1285.
10. Eperon, S., K. Bronnimann, A. Hemphill, and B. Gottstein. 1999. Susceptibility of B-cell deficient C57BL/6 (microMT) mice to Neospora caninum infection. Parasite Immunol. 21:225-236.
11. Khan, I. A., J. D. Schwartzman, S. Fonseka, and L. H. Kasper. 1997. Neospora caninum: role for immune cytokines in host immunity. Exp. Parasitol. 85:24-34.
12. Liddell, S., M. C. Jenkins, and J. P. Dubey. 1999. Vertical transmission of Neospora caninum in BALB/c mice determined by polymerase chain reaction detection. J. Parasitol. 85:550-555.
13. Lindsay, D. S., and J. P. Dubey. 1989. Neospora caninum (Protozoa: apicomplexa) infections in mice. J. Parasitol. 75:772-779.
14. Lindsay, D. S., S. D. Lenz, R. A. Cole, J. P. Dubey, and B. L. Blagburn. 1995. Mouse model for central nervous system Neospora caninum infections. J. Parasitol. 81:313-315.
15. Long, M. T., T. V. Baszler, and B. A. Mathison. 1998. Comparison of intracerebral parasite load, lesion development, and systemic cytokines in mouse strains infected with Neospora caninum. J. Parasitol. 84:316-320.
16. Luo, W., F. Aosai, M. Ueda, K. Yamashita, K. Shimizu, S. Sekiya, and A. Yano. 1997. Kinetics in parasite abundance in susceptible and resistant mice infected with an avirulent strain of Toxoplasma gondii by using quantitative competitive PCR. J. Parasitol. 83:1070-1074.
17. Nishikawa, Y., N. Inoue, X. Xuan, H. Nagasawa, I. Igarashi, K. Fujisaki, H. Otsuka, and T. Mikami. 2001. Protective efficacy of vaccination by recombinant vaccinia virus against Neospora caninum infection. Vaccine 19:1381-1390.
18. Quinn, H. E., C. M. Miller, C. Ryce, P. A. Windsor, and J. T. Ellis. 2002. Characterization of an outbred pregnant mouse model of Neospora caninum infection. J. Parasitol. 88:691-696.
19. Zenner, L., A. Foulet, Y. Caudrelier, F. Darcy, B. Gosselin, A. Capron, and M. F. Cesbron-Delauw. 1999. Infection with Toxoplasma gondii RH and Prugniaud strains in mice, rats and nude rats: kinetics of infection in blood and tissues related to pathology in acute and chronic infection. Pathol. Res. Pract. 195:475-485.(Esther Collantes-Fernánde)
ABSTRACT
The kinetics of Neospora caninum loads in mice inoculated with NC-Liv or NC-1 isolates were studied. The acute phase was characterized by parasitemia and the detection of parasite DNA in several organs, whereas during the chronic phase, the parasite was detected mainly in the brain. Mice infected with NC-Liv developed clinical signs, showing higher brain parasite burdens than NC-1-infected mice.
TEXT
Neospora caninum is considered one of the main agents implicated in bovine abortion (8). Mouse inoculation is the model most widely used to study aspects of N. caninum infection (3, 6, 10, 11, 12, 13, 14, 15) and offers a tool with which to characterize and compare the virulence of N. caninum isolates (1, 18). However, studies of parasite loads in blood and tissues during the acute and chronic phases of infection are lacking. Here we determined the organs that support parasite replication and persistence in a mouse model and the relationship between the parasite isolate and virulence. This knowledge will aid in evaluating the efficiency of drugs and vaccines.
Fifty-five 6-week-old female BALB/c mice (Harlan, Barcelona, Spain) were infected with 106 tachyzoites from NC-Liv (2) (group 1) or NC-1 (9) (group 2) isolates by intraperitoneal (i.p.) injection as described previously (6). A control group (group 3) was inoculated i.p. with phosphate-buffered saline. Three random animals from each group were sacrificed on days 1, 2, and 3 postinfection (p.i.), then every 2 days until day 21, and finally weekly up to day 63 p.i. All animal-handling procedures complied with EU legislation. Mice that appeared moribund were killed by CO2 inhalation, and samples were collected. Animals were monitored daily for the presence of clinical signs of neosporosis, and the time of the event (presence of clinical signs) was recorded, for 63 days. Evaluation of clinical signs was based on previous N. caninum infection studies, in which mice with clinical neosporosis can develop rough hair coats, inactivity, anorexia, and neurological signs consisting of head tilting, walking in circles, ataxia, pelvic limb weakness, and paralysis (1, 10, 13, 14). Blood samples (300 to 500 μl) were collected in EDTA tubes and centrifuged, and plasma was cryopreserved for enzyme-linked immunosorbent assays. Mesenteric, iliac, mediastinal, and cervical lymph nodes, spleens, livers, lungs, and brains were recovered and frozen. DNA was extracted from 107 NC-1 tachyzoites as a positive control and from 10 to 20 mg of murine tissues by using a GenomicPrep cell and tissue DNA isolation kit (Amersham Biosciences, Uppsala, Sweden). DNA from blood samples was extracted with a GenomicPrep blood DNA isolation kit (Amersham Pharmacia) by following the manufacturer's instructions. Parasite DNA in murine tissues was measured by real-time PCR using SYBR Green I. We used primer pairs from the N. caninum Nc-5 sequence to quantify parasites, and we used primers from the 28S rRNA gene to quantify host DNA (7). N. caninum-specific serum isotypes immunoglobulin G2a (IgG2a) and IgG1 were determined by an enzyme-linked immunosorbent assay using soluble N. caninum tachyzoite antigen (10, 15) (0.5 μg in 100 μl/well). We used diluted murine serum samples (1:100) and an anti-mouse IgG2a or IgG1 antibody (1:5,000; Southern Biotechnology, Birmingham, AL).
Differences between groups were analyzed using the Kruskal-Wallis test, the Mann-Whitney U test, and the Student t test. When statistically significant differences were found by the Kruskal-Wallis test, a nonparametric multiple-comparison test was applied to examine all possible pairwise comparisons. A P value of less than 0.05/[k x (k – 1)/2] was considered statistically significant, with k corresponding to the number of animals in the group. Frequencies of clinical signs of neosporosis in infected groups were compared by Fisher's exact test. In addition, the Kaplan-Meier survival method (4) was applied to estimate the portion of all individuals surviving (not showing clinical signs) at each time point (days p.i.). Data from subjects removed from the study (either because they had to be killed or because the study ended) were censored. Subsequently, the log rank statistical test for comparison of survival curves between infected groups was applied (5). Statistical analysis was carried out using SAS 8.02 (SAS Institute, Cary, NC) and GraphPad Prism 4, v. 4.03 (GraphPad, San Diego, CA), software.
None of the mice died spontaneously during the follow-up period. In infected groups, clinical signs of neosporosis were more often observed in group 1 (25.4% versus 1.8%; P < 0.01 by the Fisher F test). In the group infected with NC-Liv isolates, the numbers of mice showing clinical signs of neosporosis were four, six, three, and one on days 11, 21, 35, and 42 p.i., respectively, whereas only one mouse infected with NC-1 exhibited clinical neosporosis on day 21 p.i. (Fig. 1). Rough hair coats and apathy were the first signs observed, followed by anorexia, inactivity, and nervous signs (pelvic limb weakness, head tilting, and walking in circles). As shown in Fig. 1, the percentage of survival among NC-1-infected mice was significantly greater than that among NC-Liv-infected mice (P < 0.001 by the log rank test). Control mice remained clinically normal until the end of the experiment. Parasite DNA was detected in blood, spleens, and livers of infected mice from day 1 to days 5 to 7 p.i. The parasite load peaked on days 1 to 3 p.i. and decreased thereafter (P < 0.0001 by the Kruskal-Wallis H test; P < 0.01 by the nonparametric multiple-comparison test for days 1 to 3 versus 5 to 7 p.i. and for 5 to 7 versus >7 days p.i.) (Table 1; Fig. 2). Parasitemia was subsequently transient and very low, with sporadic detection on days 11, 13, 19, and 56 p.i. (Table 1). In the upper and lower lymph nodes, the parasite was detected intermittently at very low levels throughout the experiment (Table 1). N. caninum DNA was detected in lungs from day 1 to days 11 to 13 p.i. There was a higher parasite burden on days 1 to 3 that gradually declined until extinction around days 11 to 13 p.i. (P < 0.0001 by the Kruskal-Wallis H test; P < 0.008 by the nonparametric multiple-comparison test for days 1 to 3, 5 to 7, and 9 to 13 versus >13 days p.i.). In brains, parasites were detected from days 5 to 7 p.i. until the end of the experiment (day 63). The parasite burden reached a peak on day 13 p.i. (Fig. 2) and decreased gradually thereafter (P < 0.0001 by the Kruskal-Wallis H test; P < 0.003 by the nonparametric multiple-comparison test for days 9 to 13, 15 to 21, 28 to 35, and 42 to 49 versus days 1 to 7 p.i.). Comparison of parasite loads in organs of mice inoculated with the two isolates showed similar numbers (P > 0.05 by the Mann-Whitney U test) except in the liver and the brain. In the liver, NC-1-inoculated mice had significantly higher parasite loads on day 1 p.i. than NC-Liv-inoculated mice (P < 0.05), whereas in the brain, NC-Liv-inoculated mice had significantly higher parasite burdens than NC-1-inoculated mice on days 13, 19, and 56 p.i. (P < 0.05). Subsequently, data were also compared per isolate and group 1 produced a higher parasite burden in the brain (P < 0.05).
We compared brain parasite numbers in infected mice with and without clinical signs. The data showed significantly higher brain parasite burdens in samples from mice with clinical signs (14 mice inoculated with NC-Liv and 1 with NC-1) than in those with no clinical signs of disease (95 infected mice) (P < 0.05 by the Mann-Whitney U test).
Serum antibody response kinetics followed similar patterns in groups 1 and 2 (P > 0.05 by t test) (Fig. 3). When infected groups were compared with noninfected controls, significant increases in IgG2a and IgG1 levels were detected on days 7 and 9 p.i., respectively (P < 0.05 by t test). Mice had a mixed IgG2a/IgG1 response, with a slight predominance of IgG2a levels during the first 2 to 3 weeks p.i., followed by a tendency toward a IgG1 bias thereafter (P < 0.0001 by paired t tests).
In conclusion, we have characterized the temporal tissue distribution and parasite loads of N. caninum in a murine model. The course of N. caninum infection in mice could be separated into three phases. During the first phase (day 1 to days 5 to 7 p.i.), parasitemia was present and the peak of parasite burden in blood coincided with the maximum parasite load in the spleen, liver, and lungs. The second period (days 5 to 7 to days 11 to 13 p.i.) coincided with the development of host immunity; tachyzoites were cleared from host tissues and were detected only in the lungs and brain. The number of tachyzoites decreased in the lungs but increased in the brain. Infection became chronic in the third phase (day 13 onward); the parasite disappeared from the lungs, and the parasite load decreased in the brain and remained at low levels until the end of the experiment (day 63 p.i.). The parasite was detected intermittently at very low levels in lymph nodes throughout the experiment. In this study, we provide evidence that the major target organ in acute N. caninum infection was the lung, whereas in the chronic phase it was the brain, corroborating that this is the major site of parasite persistence. These results agree with the findings of previous N. caninum and Toxoplasma gondii mouse model experiments (10, 12, 16, 17, 19), in which the most frequently parasitized tissues were the lungs and brain.
The parasite distribution kinetics observed strongly suggested immune regulation of infection. During the chronic infection phase, N. caninum induced high levels of both isotypes, but IgG1 production predominated and the organism established a long-term chronic infection in the brain. As in other studies, disease susceptibility during chronic infection was associated with a mixed response characterized by a low gamma interferon/interleukin-4 ratio (15).
In N. caninum infection, the parasite isolate appears to have a significant role in the progression of infection (1, 18). Here, a larger number of NC-Liv-inoculated mice than of NC-1-inoculated mice showed clinical signs, with higher brain parasite burdens, and survival curve analysis showed a marked difference between isolates. Our results suggest that the NC-Liv isolate might actually have detrimental effects on morbidity, as reported previously (1). This study also demonstrates an association between brain parasite load and the presence of clinical signs. On the other hand, NC-1-inoculated mice had higher parasite loads in the liver on day 1 p.i. than mice inoculated with NC-Liv. Our observations suggest that N. caninum isolates may have different tissue tropisms depending on the infection phase. We underline the importance of quantifying the parasite burden in order to follow the course of infection and to determine differences in virulence among N. caninum isolates.
ACKNOWLEDGMENTS
We thank Ricardo García de la Mata for statistical analyses and Vanesa Navarro for excellent technical assistance. We gratefully acknowledge D. J. L. Williams (Liverpool School of Tropical Medicine, Liverpool, United Kingdom) for generously providing the NC-Liverpool isolate and Veronica Risco for valuable help. We express our gratitude to two anonymous referees for helpful comments on the manuscript.
E.C.-F. was financed by the Spanish Ministry of Science and Technology (M.I.T. fellowship, Spain). This work was supported by a grant from the Spanish Government (AGL2001-1362) and was part of the EU research collaboration COST-854.
REFERENCES
1. Atkinson, R., P. A. Harper, C. Ryce, D. A. Morrison, and J. T. Ellis. 1999. Comparison of the biological characteristics of two isolates of Neospora caninum. Parasitology 118:363-370.
2. Barber, J., A. J. Trees, M. Owen, and B. Tennant. 1993. Isolation of Neospora caninum from a British dog. Vet. Rec. 133:531-532.
3. Baszler, T. V., M. T. Long, T. F. McElwain, and B. A. Mathison. 1999. Interferon-gamma and interleukin-12 mediate protection to acute Neospora caninum infection in BALB/c mice. Int. J. Parasitol. 29:1635-1646.
4. Bland, J. M., and D. G. Altman. 1998. Survival probabilities. The Kaplan-Meier method. BMJ 317:1572.
5. Bland, J. M., and D. G. Altman. 2004. The logrank test. BMJ 328:1073.
6. Collantes-Fernandez, E., G. Alvarez-Garcia, V. Perez-Perez, J. Pereira-Bueno, and L. M. Ortega-Mora. 2004. Characterization of pathology and parasite load in outbred and inbred mouse models of chronic Neospora caninum infection. J. Parasitol. 90:579-583.
7. Collantes-Fernandez, E., A. Zaballos, G. Alvarez-Garcia, and L. M. Ortega-Mora. 2002. Quantitative detection of Neospora caninum in bovine aborted fetuses and experimentally infected mice by real-time PCR. J. Clin. Microbiol. 40:1194-1198.
8. Dubey, J. P. 2003. Neosporosis in cattle. J. Parasitol. 89(Suppl.):S42-S56.
9. Dubey, J. P., J. L. Carpenter, C. A. Speer, M. J. Topper, and A. Uggla. 1988. Newly recognized fatal protozoan disease of dogs. J. Am. Vet. Med. Assoc. 192:1269-1285.
10. Eperon, S., K. Bronnimann, A. Hemphill, and B. Gottstein. 1999. Susceptibility of B-cell deficient C57BL/6 (microMT) mice to Neospora caninum infection. Parasite Immunol. 21:225-236.
11. Khan, I. A., J. D. Schwartzman, S. Fonseka, and L. H. Kasper. 1997. Neospora caninum: role for immune cytokines in host immunity. Exp. Parasitol. 85:24-34.
12. Liddell, S., M. C. Jenkins, and J. P. Dubey. 1999. Vertical transmission of Neospora caninum in BALB/c mice determined by polymerase chain reaction detection. J. Parasitol. 85:550-555.
13. Lindsay, D. S., and J. P. Dubey. 1989. Neospora caninum (Protozoa: apicomplexa) infections in mice. J. Parasitol. 75:772-779.
14. Lindsay, D. S., S. D. Lenz, R. A. Cole, J. P. Dubey, and B. L. Blagburn. 1995. Mouse model for central nervous system Neospora caninum infections. J. Parasitol. 81:313-315.
15. Long, M. T., T. V. Baszler, and B. A. Mathison. 1998. Comparison of intracerebral parasite load, lesion development, and systemic cytokines in mouse strains infected with Neospora caninum. J. Parasitol. 84:316-320.
16. Luo, W., F. Aosai, M. Ueda, K. Yamashita, K. Shimizu, S. Sekiya, and A. Yano. 1997. Kinetics in parasite abundance in susceptible and resistant mice infected with an avirulent strain of Toxoplasma gondii by using quantitative competitive PCR. J. Parasitol. 83:1070-1074.
17. Nishikawa, Y., N. Inoue, X. Xuan, H. Nagasawa, I. Igarashi, K. Fujisaki, H. Otsuka, and T. Mikami. 2001. Protective efficacy of vaccination by recombinant vaccinia virus against Neospora caninum infection. Vaccine 19:1381-1390.
18. Quinn, H. E., C. M. Miller, C. Ryce, P. A. Windsor, and J. T. Ellis. 2002. Characterization of an outbred pregnant mouse model of Neospora caninum infection. J. Parasitol. 88:691-696.
19. Zenner, L., A. Foulet, Y. Caudrelier, F. Darcy, B. Gosselin, A. Capron, and M. F. Cesbron-Delauw. 1999. Infection with Toxoplasma gondii RH and Prugniaud strains in mice, rats and nude rats: kinetics of infection in blood and tissues related to pathology in acute and chronic infection. Pathol. Res. Pract. 195:475-485.(Esther Collantes-Fernánde)