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*
Laboratory of Parasitic Diseases and
Laboratory of Clinical Immunology, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD 20892;
Corixa Corp., Seattle, WA 98104; and
Infectious Disease Research Institute, Seattle, WA 98104
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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production by CD8+ T cells in lymph nodes draining
the challenge site. These data suggest that under conditions of natural
challenge, DNA vaccination has the capacity to confer complete
protection against cutaneous leishmaniasis and to prevent the
establishment of infection reservoirs. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Leishmania major, the etiologic agent of zoonotic cutaneous
leishmaniasis in the Old World, has been extensively used in mouse
models to understand the requirements for effective vaccination against
healing and nonhealing forms of leishmaniasis. Depending on the
genotype of the mouse, L. major infection leads to the
development of polarized Th1 or Th2 responses that control resistance
or susceptibility, respectively, to this intracellular parasite
(8, 9). There have been a number of studies involving
murine vaccination with DNA encoding parasite Ags, including gp63
(10, 11), LACK (12, 13) and PSA-2
(14), with varying results. Although most of the studies
involving DNA as well as protein-based vaccines have been conducted in
susceptible BALB/c mice, the healing lesions produced in C57BL/6 mice
provide a more relevant model of L. major infection in
natural reservoirs and in human hosts. Furthermore, in almost every
case the efficacy of Leishmania vaccines has been evaluated
using a high dose of parasites
(105–107) inoculated into
the footpad or other s.c. sites. Recently, a natural infection model in
resistant mice has been developed that takes into account two main
features of natural transmission: low-dose (100 metacyclic
promastigotes) and intradermal inoculation (the ear dermis)
(15). In this model, the evolution of small, healing
dermal lesions occurs in three distinct phases: 1) an initial
"silent" phase, lasting 4–5 wk, favoring the establishment of the
peak load of parasites in the dermis in the absence of lesion
formation; 2) an acute phase, lasting 5–10 wk, corresponding to the
development and resolution of a lesion that is associated with an acute
infiltration of neutrophils, macrophages, and eosinophils into the
dermis, and is coincident with the onset of immunity and the killing of
parasites in the site; and 3) a chronic phase, lasting for the life of
the animal, during which a low number of parasites persists in the skin
in the absence of overt pathology. Adaptive immunity in this model
confirmed a role for Th1 cells, and in addition revealed a requirement
for CD8+ T cells, based on the results obtained
in 
2-microglobulin-deficient mice,
CD8-deficient mice, and CD8-depleted mice, which in each case failed to
control infection in the
skin.2
In the present work, the natural challenge model was used to evaluate
the potency and durability of two vaccines, components of which are
currently being tested in preclinical and clinical trials: 1) a mixture
of plasmid DNAs encoding the Leishmania Ags LACK (12, 13, 16), LmSTI1 (17), and TSA (18);
and 2) heat-killed promastigotes plus recombinant IL-12
(19, 20, 21, 22). Vaccine efficacy was evaluated in the
context of all three phases of infection. Although both vaccines
conferred complete protection against the development of dermal
lesions, this complete protection lasted longer in the DNA-vaccinated
mice. Furthermore, only the DNA vaccine reduced the parasitic burden in
the skin during the acute and chronic stages to the low levels achieved
in healed mice, and only the DNA vaccine eliminated the capacity of
healed mice to serve as infection reservoirs for vector sand flies. The
powerful, long-lasting protection conferred by AgDNA was associated
with the trafficking of CD8+ T cells to the site
of challenge in the skin and with the production of IFN- by
CD8+ T cells in draining lymph nodes.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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C57BL/6 (B6) and BALB/c mice were purchased from the Division of Cancer Treatment, National Cancer Institute (Frederick, MD). All mice were maintained in the National Institute of Allergy and Infectious Diseases Animal Care Facility under pathogen-free conditions.
Plasmid construction and purification
A cDNA encoding a truncated LACK protein (aa 143–312) was
cloned in-frame downstream to a Kozak consensus sequence and an
initiation codon into a pECE vector. The insert was excised using
HindIII and ligated into expression vector PcDNA-3
downstream to the CMV promoter (Invitrogen, San Diego, CA). The
full-length sequences of LmSTI1 and TSA were PCR amplified (
1.64 and
0.6 kb. respectively) from L. major genomic DNA using
sequence-specific primers and subcloned into pcDNA3.1 (BamHI
and EcoRI sites). Plasmid DNAs were purified by
double-banding cesium chloride gradient ultracentrifugation. The
260:280 UV absorption ratios ranged from 1.8 to 2.0.
Immunization
Mice were injected in the left hind footpad with a mixture of 100 µg of each plasmid DNA encoding either LACK, LmSTI1, or TSA (300 µg of total DNA) or 300 µg of control DNA (empty vector) suspended in 50 µl of sterile PBS. The immunization with protein was conducted by the injection of 50 µg of heat-killed L. major promastigotes (ALM)3 with or without 1.5 µg of rIL-12 (Genetics Institute, Cambridge, MA). The protein-based vaccine was prepared from whole-cell, heat-killed L. major (ALM) and is identical to that being used with BCG as adjuvant in phase III clinical trials in Iran and Sudan (21, 22). Each group was boosted 2 wk later using the same regimen.
Infectious challenge
L. major clone V1 (MHOM/IL/80/Friedlin) promastigotes were grown at 26°C in medium 199 supplemented with 20% Hi-FCE (HyClone, Logan, UT), 100 U/ml penicillin, 100 µg/ml streptomycin, 2 mM L-glutamine, 40 mM HEPES, 0.1 mM adenine (in 50 mM HEPES), 5 mg/ml hemin (in 50% triethanolamine), and 1 mg/ml 6-biotin (M199/S). Infective-stage promastigotes (metacyclics) of L. major were isolated from stationary cultures (4- to 5-day old) by negative selection using peanut agglutinin (Vector Laboratories, Burlingame, CA). Mice were challenged 2 or 12 wk postboost using 100 metacyclic promastigotes. Parasites were inoculated intradermally into the ear dermis using a 27.5-gauge needle in a volume of 10 µl. The evolution of the lesion was monitored by measuring the diameter of the induration of the ear lesion with a direct reading Vernier caliper (Thomas, Swedesboro, NJ).
Parasite quantitation
Parasite loads in the ears were determined as previously described (15). Briefly, the ventral and dorsal sheets of the infected ears were separated, deposited dermal side down in DMEM containing 100 U/ml penicillin, 100 µg/ml streptomycin, and 1 mg/ml collagenase A (Sigma, St. Louis, MO), and incubated for 2 h at 37°C. The sheets were cut into small pieces and homogenized using a Teflon-coated microtissue grinder in a microfuge tube containing 100 µl of M199/S. The tissue homogenates were filtered using a 70-µm cell strainer (Falcon Products, St. Louis, MO) and serially diluted in a 96-well flat-bottom microtiter plate containing biphasic medium, prepared using 50 µl of NNN medium containing 30% of defibrinated rabbit blood and overlaid with 50 µl of M199/S. The number of viable parasites in each ear was determined from the highest dilution at which promastigotes could be grown out after 7 days of incubation at 26°C. The number of parasites was also determined in the local draining lymph nodes (retromaxilar). The lymph nodes were recovered and mechanically dissociated using a pellet pestle and then serially diluted as above.
In vivo recall response
Mice were injected in both ears with a combination of living and killed Ags comprised of 106 metacyclic L. major promastigotes and 12.5 µg of soluble leishmanial Ag (SLA) prepared from 3x freeze-thawed stationary phase L. major promastigotes. The increase of the ear thickness was measured 72 h later with a direct reading Vernier caliper. At this time, mice were sacrificed and cells from the ear dermis and local draining lymph nodes (three to four mice) were obtained. Briefly, the retromaxillar draining lymph nodes were recovered, mechanically dissociated using a pellet pestle, and pooled. The ears were collected and the ventral and dorsal dermal sheets were separated and incubated, dermal side down on RPMI 1640, NaHCO3, penicillin/streptomycin/gentamicin containing 1 mg/ml collagenase A (Sigma) for 2 h or a mixture of collagenase A (1 mg/ml) and liberase CI enzyme blend (0.28 Wünsch units/ml; Boehringer Mannheim, Indianapolis, IN) for 1 h. The ears were pooled, cut in small pieces, and filtered through a 70-µm nylon cell strainer (Becton Dickinson, Mountain View, CA) before being washed twice in RPMI 1640, NaHCO3, penicillin/streptomycin/gentamicin, 10% FCS, and 0.05% DNase (Sigma).
Pooled cells from draining lymph nodes were resuspended in RPMI 1640
containing 10% FBS, 100 U/ml penicillin, and 100 µg/ml streptomycin
at 2.5 x 106 cells/ml, and 0.2 ml was
plated in U-bottom 96-well plates. Cells were incubated at 37°C in
5% CO2 for 24 h with or without addition of
soluble L. major Ag (SLA, 25 µg/ml) or Con A (10 mg/ml).
IFN- in 24-h culture supernatants was quantitated by ELISA. For the
analysis of surface markers and intracytoplasmic staining for IFN-,
cells were stimulated with 25 µg/ml SLA in the presence of
anti-CD28 and recombinant mouse IL-2 for 6 h, at which time
brefeldin A was added (10 µg/ml). The cells were cultured for an
additional 18 h and then fixed in 4% paraformaldehyde. Before
staining, cells were incubated with an anti-FcIII/II
(PharMingen, San Diego, CA) receptor and 10% normal mouse serum in PBS
containing 0.1% BSA and 0.01% NaN3. The
staining of surface and cytoplasmic markers was performed sequentially:
the cells were stained for the surface marker CD3 (145-2 C11, FITC
labeled; PharMingen), CD4 or CD8 (RM4-5 and 53-6.7, cychrome
conjugated; PharMingen) followed by a permeabilization step and
staining with anti-IFN- conjugated to R-PE (JE56-5H4;
PharMingen). Each incubation was conducted for 30 min on ice. The
isotype controls used were rat IgG2b (A95-1; PharMingen) and rat IgG2a
(R35-95; PharMingen). The frequency of CD4+ and
CD8+ T cells was determined by gating on
CD3+ cells. For each sample, at least 50,000
cells were analyzed. The data were collected and analyzed using
CellQuest software and a FACSCalibur flow cytometer (Becton
Dickinson).
Transmissibility of parasites from infected ears to sand flies
At 12 wk after challenge, the ability of the infected ears to provide a source of parasites to sand flies was investigated. Two- to 4-day-old Phlebotomus papatasi females were obtained from a colony initiated by field-caught specimens from the Jordan Valley and were reared at the Department of Entomology, Walter Reed Army Institute of Research (Silver Spring, MD). Fifteen sand flies were placed in individual vials with meshed surfaces. Mice were anesthetized i.p. with 200 µl of 20 mg/ml ketamine HCl (Phoenix Pharmaceuticals, St. Joseph, MO). Individual ears of anesthetized mice were pressed flat against the meshed surface of the vials using clamps that were specially designed for this purpose. The flies were allowed to feed in the dark for a period of 30 min to 1 h. Blood-fed females from each vial were separated and maintained in individual pots lined with plaster of Paris, provided a 50% sucrose solution and water, and their midguts were dissected 48 h later and examined microscopically for the presence of promastigotes.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Mice vaccinated with either AgDNA or ALM/rIL-12 and challenged 2
wk later with 100 metacyclic promastigotes in the ear dermis were
almost completely protected against the development of dermal lesions,
comparable to the resistance displayed by healed mice (Fig. 1
). The mice vaccinated with ALM alone or
control DNA (empty plasmid) developed lesions similar in size and
duration to the unvaccinated mice: the lesions appeared at week 4,
reached a peak at 6–7 wk, and were completely healed at 10–12 wk.
Since in the natural challenge model, the peak number of parasites in
the site occurs just before the development of the lesion, the parasite
load in the ear was monitored at week 4. The absence of dermal
pathology in the vaccinated mice correlated with an approximate
100-fold reduction in the number of parasites in the skin (4.4 x
104 parasites in naive mice vs 3.5 x
102 in the AgDNA group and 8 x
102 in ALM/rIL12 group) (Fig. 2). These groups also had an approximate
10-fold reduction in the parasite burden in the lymph nodes draining
the inoculation site. Vaccination with ALM alone or with the empty
vector did not reduce the number of parasites found in the skin or in
the draining nodes. After healing (10 wk post infection), 90% of
the organisms had been killed or cleared from the inoculation site in
all of the groups, although the number of parasites in the ears of
AgDNA-vaccinated mice (1.2 x 102) or
ALM/rIL-12 vaccinated mice (1.7 x 102)
remained 100-fold lower compared with the unvaccinated group
(4.2 x 104). Interestingly, no parasites
were found in the local draining nodes of the AgDNA-vaccinated animals
at 10 wk postinfection, a level of control that was otherwise achieved
only in the mice that had healed their primary infections.
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To evaluate the durability of the immunity induced by the DNA and
killed vaccines, the animals were challenged 12 wk after boosting.
BALB/c mice were included for comparison. As shown in Fig. 3, the protection against dermal
pathology that was achieved using ALM/rIL-12 was mouse strain
dependent: BALB/c mice showed no evidence of protection, confirming
earlier observations (13), whereas B6 mice maintained
significant, though partial protection. In contrast, both BALB/c and B6
mice vaccinated with AgDNA were almost completely protected against the
development of dermal lesions for at least 10 wk postchallenge. Only
one AgDNA-vaccinated animal of either strain showed any lesion, which
in each case remained small and rapidly resolved. Mice vaccinated with
ALM alone or control DNA again showed no significant reduction in
dermal pathology compared with unvaccinated animals.
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). The tissue
parasite burdens in each of these sites was actually increased when
re-examined at week 10, suggesting that the vaccine may have delayed
but not significantly reduced the peak parasite loads established in
the inoculation site. In contrast, the parasite loads in the mice
vaccinated with AgDNA were reduced to barely detectable levels in the
skin (<10) and undetectable levels in the draining nodes, even when
examined at week 4, when >10,000 parasites were found in the skin of
unvaccinated and control DNA-vaccinated mice. In terms of tissue
parasite burdens, the immunity conferred by AgDNA was as effective as
the naturally acquired immunity operating in the healed mice, and its
potency was actually increased at 12 wk compared with 2 wk
postvaccination.
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To determine whether the number of parasites in the skin during
the posthealing, chronic stage of infection (12 wk postchallenge) was
sufficiently high to be picked up by vector sand flies, the ears of
unvaccinated and vaccinated mice, challenged 12 wk after vaccination,
were exposed to the bites of a natural vector, P. papatasi
(15 flies per ear). At least 70% of the flies in each group
successfully obtained a blood meal. Blood-engorged flies were dissected
48 h later and scored for the presence or absence of parasites in
their midguts. In results pooled from two separate studies, 50% of the
ears of the unvaccinated mice transmitted parasites to the sand flies
(Fig. 5). Such efficient transmission was
also found in the mice vaccinated with ALM/rIL-12 (42%), ALM alone
(50%), or control plasmid (50%). In contrast, the transmission in
mice vaccinated with AgDNA was completely abrogated (0 of 12 ears), and
no transmission was observed in the rechallenged site of the healed
mice. Of the ears that were capable of transmitting L.
major, the efficiency of transmission, as determined by the
percentage of blood-fed flies that were positive for parasites, ranged
between 10 and 50%, with no significant difference between the groups
(data not shown).
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Surrogate markers of immunity to leishmaniasis are thought to
include Ag-induced delayed-type hypersensitivity (DTH) responses in
vivo and IFN- production by T cells following restimulation with Ag
in vitro. These recall responses were evaluated 12 wk after vaccination
by injecting into the ear dermis a mixture of L.
major-soluble Ag (SLA, 12.5 µg) and living metacyclic
promastigotes (106), and then measuring 1) the
change in ear thickness at 48 h and 72 h, 2) the number and
kinds of T cells in the inflammatory dermis, and 3) IFN- production
by lymph node cells draining the ear following restimulation in vitro
using SLA, anti-CD28, and IL-2. Table I shows that the mice receiving AgDNA
displayed a stronger DTH (0.51-mm increase in ear thickness) compared
with the other vaccinated or unvaccinated groups (0–0.25 mm). The
healed mice showed the strongest DTH response (1.20-mm increase). The
DTH response was associated with the recruitment of
CD4+ (3.2 x 104) and
CD8+ cells (1.2 x
104) in the ears of mice vaccinated with AgDNA
that in each case was 2- to 3-fold greater than the number of
CD4+ (0.6–1.5 x 104)
or CD8+ cells (0.3–0.7 x
104) recovered from the ears of each of the other
vaccinated or unvaccinated groups (Table II, experiment 1). T cell recruitment was
highest in the rechallenged site of the healed mice (23 x
104 and 7.2 x 104 for
CD4+ and CD8+,
respectively). In a subsequent experiment involving DNA-vaccinated mice
and more efficient extraction of cells from the inflammatory ear
dermis, there were 2.5 x 105
CD4+ and 1.2 x 105
CD8+ T cells present in the inoculation site of
mice immunized with AgDNA, representing a 4- and 6- fold increase in
the numbers of CD4+ cells and a 12- and 6- fold
increase in the number of CD8+ cells compared
with unvaccinated and control DNA-vaccinated groups, respectively
(Table II, experiment 2). Furthermore, examination of the frequency of
T cells in the inflammatory dermis revealed a 2-fold increase
compared with each of the other groups, including the healed
mice.
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by ELISA. Cells from
healed mice and from both ALM/rIL-12- and AgDNA-vaccinated mice,
secreted high levels of IFN- (Fig. 6).
Unvaccinated and control-vaccinated groups secreted little or no
IFN- in response to Ag. The frequency of lymph node cells staining
for IFN- following restimulation in vitro was high in the healed
mice for both CD4+ and CD8+
T cells (9.5 and 8.4%, respectively; Fig. 7). An increased frequency of
CD4+-IFN--producing cells was observed in the
ALM/rIL-12 group (6.2%). The only vaccine group with a substantial
increase in the number of CD8+ IFN--producing
cells was the AgDNA group (7.2%).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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by
lymph node cells from infected mice (17). The DNAs
encoding the Ags LACK (12, 13) and LmSTI1 and TSA (S.
Gurunathan, unpublished data) have each also conferred some
protection in BALB/c mice following high-dose footpad challenge. The rationale for applying a model of natural challenge in a resistant mouse strain to the evaluation of vaccines intended for use against cutaneous leishmaniasis is based on the contention that the model more accurately reproduces clinical-pathological findings associated with human disease. In particular, the model has revealed that the development of dermal lesions occurs only after a prolonged silent phase of parasite amplification in the skin, and is due to an inflammatory infiltrate that is dependent on and coincident with the onset of acquired immunity and the killing of parasites in the inoculation site. In contrast, high-dose s.c. inocula, particularly in BALB/c mice, produce rapidly evolving lesions that are formed as a consequence of large numbers of infected macrophages accumulating in the inoculation site. Vaccines that moderate the development of these sorts of lesions may not necessarily protect against, and could conceivably exacerbate, immune-mediated dermal pathology. In addition, the natural challenge model has revealed a role for CD8+ T cells in the resolution of primary infection in the skin,2 which again has not been appreciated as an important component of adaptive immunity to primary infection following conventional high-dose s.c. challenge (23, 24, 25, 26).
When challenged 2 wk postvaccination, both ALM/rIL-12 and AgDNA conferred striking immunity in C57BL/6 mice that in most cases resulted in the complete absence of dermal lesions. Such complete protection against clinical disease, which to our knowledge has not been observed in other laboratory-based trials of Leishmania vaccines, seems especially relevant to the field evaluation of vaccine efficacy, for which the primary measure is a decrease in disease incidence. Both vaccines maintained significant protection in C57BL/6 mice up to 12 wk after vaccination. In contrast, when challenged at 12 wk, only the DNA vaccine was able to protect BALB/c mice, confirming previous results established using high-dose s.c. challenge (13). The relative durability of ALM/rIL-12 vaccination in C57BL/6 mice, in which immune responses to L. major infection appear to more accurately reproduce those associated with human disease, suggests that BALB/c mice might in some cases undervalue the potential of vaccines intended for use against cutaneous leishmaniasis. The durability of Leishmania-specific effector and/or memory T cells in BALB/c mice may be compromised by the same conditions that establish their strong Th2 bias to L. major infection that is observed in naive mice.
Even in the C57BL/6 mice, however, the AgDNA demonstrated greater potency and durability than ALM/rIL-12. Whereas the immunity elicited by ALM/rIL-12 began to wane by 12 wk, both in terms of its ability to prevent lesion development and to reduce tissue parasite burden, the immunity elicited by AgDNA was if anything more potent at 12 wk postvaccination than at 2 wk. Furthermore, only the AgDNA conferred protection that extended to the chronic phase; at 10 wk postinfection, viable organisms were reduced to extremely low numbers in the skin (<10), and they were undetectable in draining nodes. The effect of vaccination on the chronic stage of infection has not, to our knowledge, been evaluated before. To establish the significance of this result in the context of reservoir potential, the chronic infection sites were exposed to the bites of P. papatasi, a natural vector of L. major. Whereas the ears of ALM/rIL-12-vaccinated mice transmitted parasites to sand flies as efficiently as the unvaccinated and control-vaccinated mice, the AgDNA-vaccinated mice failed to provide a source of parasites for pick up by flies. Applied to a field setting, these outcomes predict that DNA vaccination would have the capacity to reduce both the incidence of disease and, in foci involving anthroponotic species such as Leishmania tropica, the generation of infection reservoirs.
The more powerful, long-lasting immunity conferred by AgDNA may be
related to its ability to efficiently prime CD8+
as wells as CD4+ T cells (6, 27, 28, 29, 30), since both appear to be required for immunity in the
natural challenge model.2 Long-lived
LACK-responsive CD8+ and
CD4+ T cells are induced by LACK DNA in BALB/c
mice (13, 31). In the present analyses, only the AgDNA
induced a DTH response that could be elicited up to 12 wk
postvaccination, and only the AgDNA generated and maintained a high
frequency of CD8+ cells that could traffic to the
site of Ag challenge in the skin and produce IFN- in response to
reactivation with Ag in vitro.
In summary, the natural challenge model has revealed that both protein- and DNA-based vaccines have the capacity to completely protect mice against dermal leishmaniasis, but that the DNA vaccine can better maintain this immunity as well as reduce the intensity of acute and chronic phase infections to levels that prevent the generation of reservoirs of disease transmission.
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Acknowledgments |
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Footnotes |
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2 Y. Belkaid, E. von Stebut, S. Mendez, R. Lira, M. C. Udey, and D. L. Sacks. CD8+ T cells are required for pathogenesis and immunity in C57BL/6 mice following low-dose, intradermal challenge with Leishmania major. Submitted for publication.
3 Abreviations used in this paper: ALM, autoclaved Leishmania major promastigotes; SLA, soluble leishmanial Ag; DTH, delayed-type hypersensitivity.
Received for publication December 1, 2000. Accepted for publication February 8, 2001.
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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derived from CD4+ T cells is sufficient to mediate T helper cell type 1 development. J. Exp. Med. 188:1651.2-microglobulin-deficient mice fail to confirm a primary protective role for CD8 cells in experimental leishmaniasis. J. Immunol. 151:2077.[Abstract]
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S. Mendez, K. Tabbara, Y. Belkaid, S. Bertholet, D. Verthelyi, D. Klinman, R. A. Seder, and D. L. Sacks Coinjection with CpG-Containing Immunostimulatory Oligodeoxynucleotides Reduces the Pathogenicity of a Live Vaccine against Cutaneous Leishmaniasis but Maintains Its Potency and Durability Infect. Immun., September 1, 2003; 71(9): 5121 - 5129. [Abstract] [Full Text] [PDF]
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A. Iwasaki The Importance of CD11b+ Dendritic Cells in CD4+ T Cell Activation In Vivo: With Help from Interleukin 1 J. Exp. Med., July 21, 2003; 198(2): 185 - 190. [Full Text] [PDF]
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M. Colmenares, P. E. Kima, E. Samoff, L. Soong, and D. McMahon-Pratt Perforin and Gamma Interferon Are Critical CD8+ T-Cell-Mediated Responses in Vaccine-Induced Immunity against Leishmania amazonensis Infection Infect. Immun., June 1, 2003; 71(6): 3172 - 3182. [Abstract] [Full Text] [PDF]
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R. N. Coler, Y. A. W. Skeiky, K. Bernards, K. Greeson, D. Carter, C. D. Cornellison, F. Modabber, A. Campos-Neto, and S. G. Reed Immunization with a Polyprotein Vaccine Consisting of the T-Cell Antigens Thiol-Specific Antioxidant, Leishmania major Stress-Inducible Protein 1, and Leishmania Elongation Initiation Factor Protects against Leishmaniasis Infect. Immun., August 1, 2002; 70(8): 4215 - 4225. [Abstract] [Full Text] [PDF]
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E. G. Rhee, S. Mendez, J. A. Shah, C.-y. Wu, J. R. Kirman, T. N. Turon, D. F. Davey, H. Davis, D. M. Klinman, R. N. Coler, et al. Vaccination with Heat-killed Leishmania Antigen or Recombinant Leishmanial Protein and CpG Oligodeoxynucleotides Induces Long-Term Memory CD4+and CD8+T Cell Responses and Protection Against Leishmania major Infection J. Exp. Med., June 17, 2002; 195(12): 1565 - 1573. [Abstract] [Full Text] [PDF]
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A. Campos-Neto, J. R. Webb, K. Greeson, R. N. Coler, Y. A. W. Skeiky, and S. G. Reed Vaccination with Plasmid DNA Encoding TSA/LmSTI1 Leishmanial Fusion Proteins Confers Protection against Leishmania major Infection in Susceptible BALB/c Mice Infect. Immun., June 1, 2002; 70(6): 2828 - 2836. [Abstract] [Full Text] [PDF]
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Y. Belkaid, E. Von Stebut, S. Mendez, R. Lira, E. Caler, S. Bertholet, M. C. Udey, and D. Sacks CD8+ T Cells Are Required for Primary Immunity in C57BL/6 Mice Following Low-Dose, Intradermal Challenge with Leishmania major J. Immunol., April 15, 2002; 168(8): 3992 - 4000. [Abstract] [Full Text] [PDF]
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S. G. Reed Leishmaniasis Vaccination: Targeting the Source of Infection J. Exp. Med., August 6, 2001; 194(3): f7 - f10. [Full Text] [PDF]
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J. G. Valenzuela, Y. Belkaid, M. K. Garfield, S. Mendez, S. Kamhawi, E. D. Rowton, D. L. Sacks, and J. M.C. Ribeiro Toward a Defined Anti-Leishmania Vaccine Targeting Vector Antigens: Characterization of a Protective Salivary Protein J. Exp. Med., August 6, 2001; 194(3): 331 - 342. [Abstract] [Full Text] [PDF]
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)
or vaccinated twice (2-wk interval) in the footpad with either 100 µg
of each AgDNA (
), 300 µg of control DNA (
), 50 µg of ALM +
1.5 µg of rIL-12 (•), or 50 µg of ALM alone (
). Mice with
previously healed dermal lesions were included as immune controls
(
). Values represent the mean lesion diameter ± SEM of 4–10
mice, 8–20 ears/group. *, p < 0.001 for ALM +
rIL-12 (•) and AgDNA (
). Results are expressed as geometric mean ±
SEM of six to eight ears per group. Parasite loads in draining nodes
were determined from the pool of six to eight lymph nodes. *,
p < 0.05, significant decrease compared with
unvaccinated, ALM alone, and control DNA.
