Leptospirosis is a major cause of reproductive system disease among cattle.1–12 Leptospira borgpetersenii serovar Hardjo type hardjo-bovis is the most common cause of bovine leptospirosis in the United States13 and is common worldwide.14–23
Cattle are the maintenance host for LHB, and infections can cause abortions,11 stillbirths,4 weak calves,4,11 embryonic death,24 higher number of breedings required per conception,24 and mastitis.25–28 Congenital infection is a possible sequela for calves born to infected dams.4 Transmission primarily occurs through exposure to contaminated urine, reproductive fluids, and tissues. Infected cattle can become chronic renal carriers of LHB and serve as a reservoir of infection for the cattle herd.29
Vaccination of cattle against viral and bacterial causes of reproductive failure such as IBRV, BVDV, Campylobacter fetus, and Leptospira spp is an important means of disease control for US cattle owners. Multiple commercial vaccines exist that contain different combinations of viral and bacterial antigens aimed at immunization of cattle against respiratory and reproductive disease. In the United States, vaccines that are currently available to immunize cattle against leptospirosis contain either LHB or LHP, which is primarily isolated from cattle in the United Kingdom,14 with no known reports of LHP isolation from cattle in the United States.
Recommendations on providing protection for young calves (< 2 months old) against LHB infection and colonization are limited. A study in Australia in 1991 that included an experimental alum bivalent Leptospira vaccine containing L interrogans serovar Pomona and LHB (at that time referred to as L interrogans serovar Hardjo type hardjo-bovis strain 181) confirmed immune protection following challenge 28 weeks after calves were vaccinated at 4 weeks of age and booster vaccinated at 8 weeks after the first vaccination.30 Leptospira vaccines that contain LHP lack the ability to provide a long duration of immunity, prevent infection, prevent urinary shedding, or prevent reproductive losses when challenged with US strains of LHB.4,31 Monovalent vaccines have had variable responses.32–34
The purpose of the study reported here was to determine the duration of immunity of an inactivated commercially available viral-bacterial combination vaccine containing IBRV, BVDV, parainfluenza 3 virus, BRSV fractions, and C fetus; L interrogans serovars Canicola, Icterohemorrhagiae, and Pomona; Leptospira kirschneri serovar Grippotyphosa; and LHB bacterial fractions to prevent urinary shedding and urogenital tract colonization.
Materials and Methods
Cattle—All procedures involving cattle were approved by the Institutional Animal Care and Use Committees at Novartis Animal Health and Rural Technologies. Thirty-six 4-week-old Holstein heifers with no history of vaccination or exposure to LHB from a single source dairy were selected for use in this study. The calves had been fed colostrum from cows vaccinated with a commercially available multivalent respiratory-reproductive vaccine that contained L interrogans serovars Pomona, Hardjo, Canicola, and Icterohemorrhagiae, and L kirschneri serotypye Grippotyphosa. All animals were ear-notch negative to BVDV via immunohistochemical testing.
Animal housing and feeding—Upon arrival at the research facility, the heifers were housed individually in calf huts and fed an age-appropriate diet. Heifers were comingled in a single pen following the second vaccination at approximately 2 months of age and had a common source of feed and water. Housing conditions, feeder space, and water sources were all in accordance with the Guide for the Care and Use of Agricultural Animals in Agricultural Research and Teaching as set forth by both institutional animal care and use committees.
Group assignments and vaccination—Heifers were randomly assigned to 1 of 2 groups (treatment or control). All research personnel were unaware of the treatment allocations. Treatment group heifers (n = 18) were vaccinated with a commercial vaccinea that contained IBRV, BVDV, parainfluenza 3 virus, BRSV fractions, and C fetus; L interrogans serovars Canicola, Icterohemorrhagiae, Pomona; L kirschneri serovar Grippotyphosa; and LHB bacterial fractions with a unique adjuvant system.b Control group heifers (n = 18) received a commercially available vaccinec that contained the same adjuvant and IBRV, BVDV, parainfluenza 3 virus, and BRSV fractions, without the bacterial fractions that treatment group heifers received. Both groups were vaccinated with a 5-mL dose at approximately 1 month of age and booster vaccinated 4 weeks later (at approx 2 months of age). On day 358 following the second vaccination, all heifers were relocated to a challenge facility.
Challenge preparation—On day 280 after vaccination, 2 untreated heifers (passage heifers) were used for production of challenge inoculum. These heifers were housed in a separate building at the same facility, maintained on the same ration as the other 2 groups, and allowed free access to grass hay. Water was provided ad libitum.
The LHB strain 203 designated for the challenge was originally obtained from the National Animal Disease Center, Ames, Iowa. This strain has been described.4–6 The 2 passage heifers were challenged on days 280, 281, and 282 after vaccination. All heifers were challenged by periocular administration of LHB strain 203. This strain has been used in previous studies.4,31–33 Blood and urine samples were obtained from the passage heifers on DAC 0, 15, 23, 28, 36, 42, and 51 to follow progress of the infection. Urine samples collected from both passage heifers were tested for the presence of the organism. Bacteriologic culture of urine yielded positive results beginning the third week following challenge and continuing until necropsy. On DAC 51, the passage heifers were euthanized by IV administration of a barbiturated overdose. The kidneys were immediately removed and transported to the laboratory for processing and Leptospira isolation. From each kidney, tissues affected with lesions were removed, weighed, and homogenized before being added to the culture medium. The kidney homogenates were diluted in growth medium and submitted for bacteriologic culture under the same conditions as was the urine. Those with positive culture results were passaged into fresh growth medium to produce the challenge inoculum. The new cultures were checked weekly to determine organism growth and motility. The cultures were considered appropriate for challenge when sufficient motility was observed and a sufficient amount of culture inoculum was available for all heifers to be challenged with 1 mL of inoculum containing approximately 1 × 106 organisms/mL on 3 consecutive days. The identity of the organisms as spirochetes was confirmed by dark-field microscopy, and these were considered to be leptospires.
Challenge—Each heifer was challenged on 3 consecutive days by periocular administration of 1 mL (0.5 mL in each eye) of live LHB strain 203 bacterial culture containing 1 × 106 cells/mL of the challenge inoculum. The challenge was initiated 369 days after the second vaccination. The heifers were observed once daily for general clinical signs from the day of challenge through the day of necropsy.
Blood and urine sample collection—Blood samples were collected via jugular venipuncture from all animals prior to vaccination and prior to challenge (day −1) and at DAC 28 and 56. The samples were processed, and an aliquot was submitted to South Dakota State University Animal Disease Research and Diagnostic Laboratory for Leptospira MAT. Urine samples were collected from each heifer weekly, starting 1 day before the challenge and continuing until euthanasia, for a total of 9 samples/heifer (8 samples/heifer were collected from DAC 7 through 55). Urine was collected following furosemide administration and submitted for bacteriologic culture and IFA.
Animal disposition and tissue sample collection—All heifers were euthanized on DAC 56, and reproductive system and kidney tissues were collected at necropsy and submitted for bacteriologic culture to detect Leptospira. Reproductive system tissues sampled included the uterus, ovary, and oviducts. Kidney tissue samples obtained directly from observed lesions were submitted for bacteriologic culture. If no lesions were found, then tissues from 3 random sites including the cortex were harvested, pooled, and submitted for bacteriologic culture.
Bacteriologic culture of urine—Urine samples were diluted in transport medium (1× PBS solution with 1% bovine serum albumin), and dilutions of 10−0, 10−1, and 10−2 were inoculated in Ellis (80/40) culture medium and incubated at 29°C for up to 2 months. Culture samples with bacterial growth were confirmed to have leptospires by use of dark-field microscopy. Cultures with no growth after 2 months of observation were considered to have yielded negative results.
Urine IFA—Urine samples were concentrated by centrifugation. The 3 mL of urine from the bottom of each tube was removed and divided into aliquots in new tubes. The concentrated urine samples were spotted onto a glass slide, allowed to dry, and fixed in acetone. A fluorescein isothiocyanate–conjugated anti-Leptospira antisera conjugatee and counterstainf were used to visualize any leptospires present in the samples. Leptospires were identified by their typical shape and fluorescence through the use of incident-light fluorescence microscopy.
Leptospira culture of urogenital tissues—Kidney and reproductive system tissues were harvested immediately following euthanasia. Each tissue sample was homogenized and diluted in transport medium, and dilutions of 10−1, 10−2, and 10−3 were inoculated in Ellis (80/40) culture medium. Samples were then incubated at 29°C for up to 2 months. Culture samples with bacterial growth were confirmed to have leptospires by use of dark-field microscopy. Cultures with no growth after 2 months were considered to have yielded negative results.
Statistical analysis—A statistical software programg was used to perform the Fisher exact test and compare treatment groups for dichotomous data. Serologic titers were transformed into logarithms, and Student t tests were used to compare differences in transformed titers. Values of P < 0.05 were considered significant.
Results
MAT results—Blood samples were obtained from heifers in the treatment and control groups on the day of vaccination for MAT to determine maternal antibody titers against LHB; L interrogans serovars Canicola, Icterohemorrhagiae, and Pomona; and L kirschneri serovar Grippotyphosa. All calves had titers of < 1:100 for all 5 serovars.
Blood samples were taken from all heifers in treatment and control groups on days −1,28, and 56 after challenge for MAT to determine antibody titers against LHB; L interrogans serovars Canicola, Icterohemorrhagiae, and Pomona; and L kirschneri serovar Grippotyphosa. Antibodies against L interrogans serovars Canicola, Icterohemorrhagiae, and Pomona as well as L kirschneri serovar Grippotyphosa were not detected on postchallenge MAT in any animals through the end of the study. Postchallenge MAT results for LHB remained negative in 8 of 18 vaccinated heifers at DAC 28 (mean log2 titer, 3.45), and all were negative at DAC 56. The log2 MAT titers from the 10 vaccinated heifers that were seropositive on MAT ranged from 7.64 to 8.64 (9 heifers with titers of 200 [log2, 7.64] and 1 with a titer of 400 [log2, 8.64]). In contrast the postchallenge MAT antibody response was evident in 17 of 18 control heifers by DAC 28 (mean log2 titer, 8.33 [range, 7.64 to 9.64]). Three of these 17 control heifers had a titer of 200 (log2, 7.64), 8 had a titer of 400 (log2, 8.64), and 6 had a titer of 800 (log2, 9.64). Results of MAT were still positive in the controls at DAC 56, with 12 of 18 having a mean log2 titer of 5.82 (range of positive animals, 7.64 to 9.64; 4/12 heifers had titers of 200; 3/12 had titers of 400, and 5/12 had titers of 800). Control heifers had significantly (P < 0.05) higher titers against LHB than did vaccinated heifers at DAC 28 and DAC 56.
Urogenital tract colonization—All vaccinated heifers (18/18 heifers) that received the LHB vaccine were protected from kidney colonization following challenge, whereas all 18 control heifers had positive results of bacteriologic culture of kidney tissue. All heifers that received the LHB vaccine were protected from colonization of the reproductive system following challenge, whereas 5 of 18 control heifers had positive results of bacteriologic culture of reproductive system tissue; this difference was significant.
Urine shedding—Urine shedding occurred in only 4 of 18 vaccinated heifers, whereas all 18 control heifers had positive results of bacteriologic culture of urine (P < 0.05), with the first positive culture results starting at DAC 21 and continuing through DAC 56 in the controls and on DAC 28 only in 1 of the 4 vaccinates. The other 3 vaccinates had positive culture results at DAC 42, 49, or 56 (1 each day). Leptospires were detected by culture in the urine at DAC 28, 42, 49, or 56 (1 different animal on each day), whereas the control heifers shed leptospires continuously in the urine through DAC 56. Following challenge, leptospires were detected in the urine of 1 vaccinated heifer via IFA at 3 time points, compared with 12 control heifers that were IFA positive for up to 5 time points (P < 0.05).
Discussion
This study revealed that calves < 2 months of age vaccinated with a commercially available multivalent reproductive-respiratory vaccine containing a US-origin LHB isolate were protected 12 months later against colonization following LHB strain 203 challenge. Because cattle are the reservoir host, developing active protection against colonization at an early age is important. This vaccine prevented renal and reproductive system colonization while significantly reducing shedding. Furthermore, following direct LHB challenge, the vaccinated heifers were exposed to the leptospiuric control heifers for an additional 8 weeks; only 4 vaccinated heifers had positive results of a bacteriologic culture (each on a single day) and no evidence of urogenital tract infection at the time of necropsy. In contrast, all the control heifers shed leptospires in their urine at 4 or more collection time points. The control heifers also had a significantly higher rate of urogenital tract colonization, with isolation of leptospires from the kidneys in all heifers (n = 18) at necropsy.
The prevention of congenital infections in apparently healthy calves4 along with the chronic renal shedding of LHB by cattle29 necessitates the protection of calves at an early age for both individual-animal protection as well as herd-level biosecurity. The respiratory-reproductive vaccine containing LHB evaluated in the present study provided that protection. The results determined in this study, to the authors' knowledge, represent the earliest documented protection against leptospirosis provided to calves by a commercial vaccine. A previous experimental vaccine induced immune protection in calves at 28 weeks of age when they were challenged with LHB strain 033.30 Calves in that study were similarly vaccinated at 4 weeks and booster vaccinated at 8 weeks of age.
The immune response in the present study likely occurred in the presence of maternal antibody. Although the calves were fed colostrum from cows vaccinated with a commercially available multivalent respiratory-reproductive vaccine that contained LHB, L kirschneri serovar Grippotyphosa and L interrogans serovars Pomona, Canicola, and Icterohemorrhagiae, the LHB titers of all calves at the time of initial vaccination were < 100, and therefore calves were considered seronegative.35 However, in an Australian study,30 1- to 2-month-old calves had MAT LHB titers that ranged from 5 to 32, so it is likely that the calves of the present study had MAT titers less than the detection limit of the MAT (100).
Duration of immunity is also an important aspect in the prevention of a reproductive system disease in cattle where frequent exposure to the organism is a possibility. In the present study, the duration of immunity was 12 months following the second vaccination. Another study34 that used a monovalent LHB vaccine found a 1-year duration of immunity, but the calves were immunized at 10 months of age. Other studies have revealed protection at 4 months following the second vaccination in calves at 8 to 12 months of age.33 Additionally, an experimental bivalent Leptospira vaccine induced protection in calves 7 months following vaccination. In that study,30 calves were vaccinated at 4 weeks, booster vaccinated at 8 weeks of age, and challenged 28 weeks after the first vaccination with LHB strain 033. To our knowledge, no multivalent vaccine has achieved 1-year duration of immunity.
Other multivalent Leptospira vaccines have been evaluated for protection against LHB. Two pentavalent Leptospira experimental vaccines prepared by a commercial company, one containing an LHB isolate and another containing an LHP isolate, were tested against an LHB challenge.31 Both vaccines failed to protect calves vaccinated at 4 to 8 months of age and subsequently challenged with LHB 6.5 months following vaccination.
An interesting finding of the present study was the lack of a strong anamnestic response in the vaccinated heifers. The LHB challenge elicited a significantly lower mean MAT serologic response to LHB (log2 titer, 3.45) at DAC 28 in 8 of 18 vaccinated heifers, compared with a mean log2 MAT titer of 8.33 in 17 of 18 controls. All vaccinates (18/18) were seronegative (titers < 100) at DAC 56, compared with only 6 of the 18 controls. It is likely that the control heifers' mean MAT response was significantly higher as a result of the active infection. This was in sharp contrast to the limited MAT serologic response after challenge in the vaccinated heifers. The lack of a strong MAT serologic response in the vaccinated heifers could simply indicate that replication of the challenge organism did not occur, at least to an extent sufficient to elicit an anamnestic serologic response.33,36 However, other studies31,32 that challenged animals with high titers of anti–LHB antibody found that they were not protected against LHB. Studies37,38 involving use of vaccines that provided protection against challenge with LHB strain 203 induced a potent postvaccination Th1 type immune response. This Th1 type protective vaccine response involved both CD4 and Λδ T lymphocytes.
Pentavalent Leptospira vaccines containing LHP have failed to protect against LHB challenge.31,32 The use of a nonadjuvanted pentavalent Leptospira vaccine containing LHP also did not elicit a strong Th1 response, compared with 2 monovalent LHB vaccines.39 The protection provided by a vaccine depends not only on the specifics of the serovar strain of the organism contained in the vaccine, but also on the adjuvant vaccine formulation. Serovar differences combined with adjuvant formulation may affect the ability of the vaccine to induce a protective immune response. The pentavalent vaccine used in the study reported here contained a US LHB strain along with a unique adjuvant system.
Vaccination with a commercial vaccine containing IBRV, BVDV, parainfluenza 3 virus, BRSV, C fetus vaccine fractions, and the LHB vaccine fraction tested here was effective in disease protection. This vaccine was effective in young animals and could be used to prevent early infections in young animals as well as prevent colonization and therefore reduce chronic LHB infections. This protection occurred in the absence of detectable antibodies on MAT, which may indicate that the vaccine induced a protective Th1 cell–mediated response. Future studies will need to be performed to directly determine the protective Th1 cell–mediated response.
ABBREVIATIONS
| BRSV | Bovine respiratory syncytial virus |
| BVDV | Bovine viral diarrhea virus |
| DAC | Day after challenge |
| IBRV | Infectious bovine rhinotracheitis virus |
| IFA | Immunofluorescence assay |
| LHB | Leptospira borgpetersenii serovar Hardjo type hardjo bovis |
| LHP | Leptospira interrogans serovar Hardjo type hardjo prajitino |
| MAT | Microscopic agglutination test |
| Th1 | T-helper 1 cell |
Vira Shield 6+VL5HB, Novartis Animal Health, Larchwood, Iowa.
Xtend SP, Novartis Animal Health, Larchwood, Iowa.
Vira Shield 6, Novartis Animal Health, Larchwood, Iowa.
Euthasol, Virbac Animal Health Inc, Fort Worth, Tex.
Internal antibody conjugate NVSL920ARS8901/NADC11397ARSR206 fluorescein isothiocyanate-conjugated anti-Leptospira antisera, National Animal Disease Center, Ames, Iowa.
Flazo Orange, Sigma-Aldrich, St Louis, Mo.
SAS, version 9.3, SAS Institute Inc, Cary, NC.
References
1. Pearson JK, Mackie DP, Ellis WA. Milk drop syndrome resulting from Leptospira hardjo. Vet Rec 1980; 106: 135–136.
2. Sullivan ND, Callan DP. Isolation of Leptospira hardjo from cows with mastitis. Aust Vet J 1970; 46: 537–539.
3. Dhaliwal GS, Murray RD, Dobson H, et al. Effect of vaccination against Leptospira interrogans serovar hardjo on milk production and fertility in dairy cattle. Vet Rec 1996; 138: 334–335.
4. Bolin CA, Thiermann AB, Handsaker AL, et al. Effect of vaccination with a pentavalent leptospiral vaccine on Leptospira interrogans serovar hardjo type hardjo-bovis infection of pregnant cattle. Am J Vet Res 1989; 50: 161–165.
5. Ellis WA, O'Brien JJ, Neill SD, et al. Bovine leptospirosis: experimental serovar hardjo infection. Vet Microbiol 1986; 11: 293–299.
6. Ellis WA, O'Brien JJ, Bryson DG, et al. Bovine leptospirosis: some clinical features of serovar hardjo infection. Vet Rec 1985; 117: 101–104.
7. Higgins RJ, Harbourne JF, Little TW, et al. Mastitis and abortion in dairy cattle associated with Leptospira of the serotype hardjo. Vet Rec 1980; 107: 307–310.
8. Kirkbride CA, Johnson MW. Serologic examination of aborted ovine and bovine fetal fluids for the diagnosis of border disease, bluetongue, bovine viral diarrhea, and leptospiral infections. J Vet Diagn Invest 1989; 1: 132–138.
9. Ellis WA. Leptospirosis as a cause of reproductive failure. Vet Clin North Am Food Anim Pract 1994; 10: 463–478.
10. Heath SE, Johnson R. Leptospirosis. J Am Vet Med Assoc 1994; 205: 1518–1523.
11. Thiermann AB. Experimental leptospiral infections in pregnant cattle with organisms of the Hebdomadis serogroup. Am J Vet Res 1982; 43: 780–784.
12. Giles N, Hathaway SC, Stevens AE. Isolation of Leptospira interrogans serovar hardjo from a viable premature calf. Vet Rec 1983; 113: 174–176.
13. Zuerner RL, Bolin CA. Repetitive sequence element cloned from Leptospira interrogans serovar hardjo type hardjo-bovis provides a sensitive diagnostic probe for bovine leptospirosis. J Clin Microbiol 1988; 26: 2495–2500.
14. Ellis WA, Thiermann AB, Marshall RB. Genotypes of Leptospira hardjo and their role in clinical disease, in Proceedings. 14th World Congr Dis Cattle 1986; 966–970.
15. Djordjevic S, Hornitzky M, Ross AD, et al. Restriction-endonuclease analysis of Australian isolates of Leptospira interrogans serovar hardjo from cattle with agalactia and abortion. Aust Vet J 1993; 70: 98–100.
16. Thiermann AB, Handsaker AL, Foley JW, et al. Reclassification of North American leptospiral isolates belonging to serogroups Mini and Sejroe by restriction endonuclease analysis. Am J Vet Res 1986; 47: 61–66.
17. Silbreck NW, Davies WD. Restriction endonuclease analysis of Australian isolates of Leptospira interrogans serovar hardjo. Aust Vet J 1989; 66: 183–184.
18. Talpada MD, Garvey N, Sprowls R, et al. Prevalence of leptospiral infection in Texas cattle: implications for transmission to humans. Vector Borne Zoonotic Dis 2003; 3: 141–147.
19. White FH, Sulzer KR, Engel RW. Isolations of Leptospira interrogans serovars hardjo, balcanica, and pomona from cattle at slaughter. Am J Vet Res 1982; 43: 1172–1173.
20. Miller DA, Wilson MA, Beran GW. Survey to estimate prevalence of Leptospira interrogans infection in mature cattle in the United States. Am J Vet Res 1991; 52: 1761–1765.
21. Miller DA, Wilson MA, Beran GW. Relationships between prevalence of Leptospira interrogans in cattle, and regional, climatic, and seasonal factors. Am J Vet Res 1991; 52: 1766–1768.
22. Bolin CA. Bovine leptospirosis prevalent in US dairy herds. Bovine Veterinarian 2003;Feb:14–15.
23. Wikse SE, Rodgers GM, Ramachandean S, et al. Herd prevalence and risk factors of Leptospira infection in beef cow/calf operations in the United States: Leptospira borgpetersenii serovar Hardjo. Bovine Pract 2007; 41(1):15–23.
24. Guitian J, Thurmond MC, Hietala SK. Infertility and abortion among first-lactation dairy cows seropositive or seronegative for Leptospira interrogans serovar hardjo. J Am Vet Med Assoc 1999; 215: 515–518.
25. Hairgrove TB. Leptospirosis in cattle, in Proceedings. 37th Annu Conv Am Assoc Bovine Pract 2004; 36–39.
26. Kingscote BF. The diagnosis of Leptospira interrogans serovar hardjo in cattle in Canada. Can Vet J 1985; 26: 270–274.
27. Robertson A, Boulanger P, Mitchell D. Isolation and identification of a leptospire of the Hebdomadis serogroup (L. Hardjo) from cattle in Canada. Can J Comp Med Vet Sci 1964; 28: 13–18.
28. Sulzer CR, Shotts EB Jr, Olsen CD, et al. Leptospirosis due to serotype hardjo in cattle. J Am Vet Med Assoc 1964; 144: 888–890.
29. Leonard FC, Quinn PJ, Ellis WA, et al. Duration of urinary excretion of leptospires by cattle naturally or experimentally infected with Leptospira interrogans serovar hardjo. Vet Rec 1992; 131: 435–439.
30. Palit A, Middleton H, Sheers J, et al. The influence of maternal antibody and age of calves on effective vaccination against Leptospira interrogans serovar hardjo. Aust Vet J 1991; 68: 299–303.
31. Bolin CA, Zuerner RL, Trueba G. Effect of vaccination with a pentavalent leptospiral vaccine containing Leptospira interrogans serovar hardjo type hardjo-bovis on type hardjo-bovis infection of cattle. Am J Vet Res 1989; 50: 2004–2008.
32. Bolin CA, Cassells JA, Zuerner RL, et al. Effect of vaccination with a monovalent Leptospira interrogans serovar hardjo type hardjo-bovis vaccine on type hardjo-bovis infection of cattle. Am J Vet Res 1991; 52: 1639–1643.
33. Bolin CA, Alt DP. Use of a monovalent leptospiral vaccine to prevent renal colonization and urinary shedding in cattle exposed to Leptospira borgpetersenii serovar hardjo. Am J Vet Res 2001; 62: 995–1000.
34. Zuerner RL, Alt DP, Palmer MV, et al. A Leptospira borgpetersenii serovar hardjo vaccine induces a Th1 response, activates NK cells, and reduces renal colonization. Clin Vaccine Immunol 2011; 18: 684–691.
35. Hotka ML, Wilson MA, Anderson TM, et al. Comparative serological study of Leptospira serovar hardjo genotypes for use in the microscopic agglutination test. J Vet Diagn Invest 2007; 19: 84–87.
36. Trueba GA, Bolin CA, Thoen CO. Evaluation of an enzyme immunoassay for diagnosis of bovine leptospirosis caused by Leptospira interrogans serovar hardjo type hardjo-bovis. J Vet Diagn Invest 1990; 2: 323–329.
37. Naiman BM, Alt D, Bolin CA, et al. Protective killed Leptospira borgpetersenii vaccine induces potent Th1 immunity comprising responses by CD4 and gammadelta T lymphocytes. Infect Immun 2001; 69: 7550–7558.
38. Naiman BM, Blumerman S, Alt D, et al. Evaluation of type 1 immune response in naïve and vaccinated animals following challenge with Leptospira borgpetersenii serovar hardjo: involvement of WC1(+) gammadelta and CD4 T cells. Infect Immun 2002; 70: 6147–6157.
39. Brown RA, Blumerman S, Gay C, et al. Comparison of three different leptospiral vaccines for induction of a type 1 immune response to Leptospira borgpetersenii serovar hardjo. Vaccine 2003; 21: 4448–4458.
