Abstract Novel treatments for cutaneous leishmaniasis (CL) are needed, due to current lack of effective universal treatments, increasing resistance among the parasite, and toxic effects or impracticality of the current therapeutics. Models of direct infection with high number of Leishmania parasites in the current research of CL involving the BALB/c mouse or Golden Syrian Hamster are considered not suitable for the assessment of antileishmanial drug efficacy because of the lack of disease similarities with humans. The saliva of the sand fly vector is known to affect the host response to infection by the Leishmania parasite. Here, we build upon a previous BALB/c model infected with luciferase-expressing Leishmania major parasites. In the present study, we infect the ear dermis instead of the foot pad or base of the tail, and compare multiple methods of infection, using parasites alone or mixed with either bites from uninfected sand flies (Phlebotomus duboscqi Diptera Psychodidae:Neveu-Lemaire) or salivary gland sonicate from sand flies. Our data show a dose–response of bioluminescent signal (which represents the parasite load at the infection site), dermal lesion development, and Leishmania Donovan Units in liver and spleen. This in vivo L. major ear infection model, once optimized, can be used for assessing the efficacy of drug compounds that have been determined as very effective in the other, high inoculum CL models. Sand fly, cutaneous leishmaniasis, bioluminescent signal, BALB/c mouse, lesion cure Cutaneous leishmaniasis (CL) is a disease with the greatest burden among people in South America, Africa, and the Middle East (Karimkhani et al. 2016). The World Health Organization (WHO) estimates 2 million new cases of CL emerge each year (World Health Organization 2010); however, the incidence is difficult to determine due to inconsistent and often unavailable diagnoses and reporting. This disease is also a risk for travelers and military personnel in these areas (World Health Organization 2010). There are currently no effective vaccines, and because of the complex parasite–host interaction and many different species of Leishmania that cause disease, there are also no effective universal treatments for CL (Aronson et al. 2016). Many of the current treatments of choice are highly toxic, require lengthy administration, or require direct lesion topical therapies (Kunzler 2013). Also, as resistance to therapeutics emerges within the Leishmania parasites (Croft et al. 2006a), there is a need to develop effective, safe, and fast-acting therapeutics that can be used for multiple lesions or for the Leishmania species that produce mucocutaneous or visceral leishmaniasis. New antileishmanial therapies for CL have been developed and tested using Leishmania spp. infections in the footpad or dorsal tail of the BALB/c mouse or Golden Syrian Hamster (Milon et al. 1995, Croft et al. 2006b, Gamboa et al. 2008, Robledo et al. 2012, Gomes-Silva et al. 2013, Ribeiro-Romao et al. 2014, Mears et al. 2015). Currently, the vast majority of leishmaniasis in vivo models use a needle challenge of parasites alone and in large quantities. However, Leishmania is transmitted to animals and humans by a sand fly vector, and the current use of the BALB/c mouse model has been questioned in that it does not represent the clinical disease progression or immune response seen in humans (Mears et al. 2015). In a natural sand fly infection, 100 to 100,000 metacyclic Leishmania parasites together with approximately 40 salivary proteins may be injected intradermally (Kimblin et al. 2008, Abdeladhim et al. 2014). The salivary proteins encourage macrophage recruitment to the bite site, and then the Leishmania parasites invade and modulate the macrophages to effectively evade the host immune system and replicate. Extracted salivary glands, when injected along with Leishmania, were shown to increase infection and duration of the ulcer compared to Leishmania alone (Belkaid et al. 1998). Leishmaniasis in vivo models were improved by the integration of reporter genes into the Leishmania species genome to monitor intracellular proliferation of the parasites over time without the need to euthanize animals. The firefly luciferase gene (LUC) is an example of a reporter gene that has been stably integrated within the parasite genome, and has shown a direct correlation between parasite numbers and luciferase activity (Roy et al. 2000). In a recent study (Schuster et al. 2014), it was also determined that use of bioluminescent parasites to quantify Leishmania spp. infection in the ear is a more sophisticated and accurate approach compared with the more traditional measurements of lesion diameter, volume, and thickness. This is an ideal refinement for conducting longitudinal studies in a more noninvasive in vivo drug screening model (Thalhofer et al. 2010). Infected sand fly challenge introduces variable parasite loads, and thus is a confounding variable for drug discovery models. It is logistically challenging to incorporate sand fly saliva into a leishmaniasis in vivo model, but we hypothesize that allowing sand flies to feed on a mouse ear and then immediately injecting a set number of parasites results in a more repeatable, natural host–parasite response that will be conducive to a CL drug discovery program. To our knowledge, this is the first attempt to use the in vivo imaging technology in developing a sand fly saliva and L. major parasite model. Materials and Methods Animals Female BALB/c mice, weighing 20–25 g, were purchased from Charles River Laboratories (Wilmington, MA), and maintained in an AAALACi-accredited institution in accordance with the Guide for the Care and Use of Laboratory Animals (Institute for Laboratory Animal Research ). The study was approved by the Walter Reed Army Institute of Research Institutional Animal Care and Use Committee (IACUC, protocol number 16-VET-08), and conducted in compliance with the Animal Welfare Act, the Guide for the Care and Use of Laboratory Animals (2011), and other federal and Department of Defense statutes and regulations related to animals and experiments involving animals. Mice were housed in individually ventilated cage systems, in solid bottom polysulfone cages (Tecniplast, Buguggiate, Italy) with 1/8-inch corncob bedding (Bed-o-cobs, The Andersons, Maumee, OH) in ABSL-2 conditions. Animals were maintained on a 12:12 (L:D) h cycle with access to ad libitum food (LabDiet 5001, St. Louis, MO), automatic RO-filtered water (onsite), Nestlets (Animal Specialties and Provisions, Quakertown, PA) and shelter enrichment (Bio-Serve, Flemington, NJ). Sentinel animals exposed to dirty bedding and rack exhaust filters were tested quarterly, and found to be negative, for the following agents by PCR and serology: mouse hepatitis virus, epizootic diarrhea of infant mice, mouse norovirus, Mouse parvoviruses (minute virus of mice; Mouse Parvovirus, Theiler’s mouse encephalomyelitis virus, mousepox, lymphocytic choriomeningitis virus, mouse adenovirus 1 and 2, pneumonia virus of mice, reovirus (REO type 3), sendai virus, mouse thymic virus, murine cytomegalovirus, Helicobacter spp., Mycloplasma pulmonis, Pasteurella pneumotropica, Pneumocystis murina, Spironucleus muris, cilia-associated respiratory bacillus, and endo- and ecto-parasites. Mice were allowed to acclimate for 1 wk after arrival prior to experimental manipulations. Parasite Cultures L. major promastigotes (NIH173 (MHOM/IR/-/173) were cultured in Schneider’s medium (Lonza Life Sciences, Walkersville, MD) supplemented with 20% heat-inactivated fetal bovine serum, as described by Caridha et al. (2017). Cultures were maintained in T75 tissue cultures flasks (Corning Life Sciences, Manassas, VA) at 22°C. Promastigotes were harvested from culture for infection by spinning at 2000 rpm for 20 min. Media was removed and the resulting pellet was suspended in 1× PBS. Two additional spins at 2000 rpm each were conducted in 1× PBS before making the final solutions of parasites suspended in 1× PBS. Generation of the Bioluminescent L. major Parasite NIH173 (MHOM/IR/-/173) was a gift from Dr. Genevieve Milon, Department of Parasitology and Mycology, Institute Pasteur, Paris, France (Lecoeur et al. 2007, 2010). Sand Flies Phlebotomus duboscqi from Mali were obtained from the National Institutes of Health and reared in the insectary at the Walter Reed Army Institute of Research by the methods described by Modi and Rowton (1999). The sand flies were maintained at 26°C and 80% relative humidity. Salivary Gland Sonicate Inoculation Combined With Leishmania Newly emerged sand flies were aspirated into paper cups and held for 7 d. Sand flies were then aspirated into soapy water to immobilize the flies. The soapy water containing the flies was poured over a fine mesh screen and rinsed with deionized water to remove the soap. The flies were placed on a microscope slide containing 1× PBS. Using dissecting pins, the head of the flies was separated from each fly, and the salivary glands were teased out from the back of the head. Twenty pairs of sand fly salivary glands were placed in 400 µl of 1× PBS. The glands were then disrupted by ultrasonication (Sonics VibraCell) for 40 cycles, lasting approximately 2 min each cycle. The tube was immediately frozen at -80°C. Prior to the start of the inoculations, the tube was removed from the freezer and placed on ice. Four microliters of the salivary gland sonicate (SGS) (0.2 glands) and 6 µl of 1× PBS containing 5 × 103, 1 × 104, or 2 × 104 parasites were inoculated into the ear dermis of each mouse. Animal Infections With L. major Mice were anesthetized using isoflurane (4% induction, 2.5% maintenance) in an induction chamber (MWI Veterinary Supply, Harrisburg, PA). Once anesthetized, 10 µl of L. major luciferase-expressing (L. major-LUC) parasite solution was injected intradermally in the concave surface of the pinna, using a 28G needle. The concentration of the parasite solution was 1 × 103, 5 × 103, 1 × 104, or 2 × 104L. major-LUC stationary phase promastigotes per 10 µl 1× PBS, with or without SGS. Sand Fly Feeding Assays Twelve hours prior to the start of each sand fly feeding assay, 10 uninfected sand flies were aspirated into 5 dram vials with a mesh screen over the opening to allow sand flies an area to feed through. The sand flies were sugar starved for 12 hr prior to the experiments. A single mouse was anesthetized using a mixture of ketamine/xylazine (50mg/kg and 5 mg/kg, respectively) IP and placed inside an empty glove box with the temperature range of 20–25°C and 70–80% relative humidity. The vial containing the sand flies was then clamped to the mouse’s ear using a C-clamp. The flies were allowed to feed undisturbed on the ear for 30 min, or until 100% of the flies had taken a bloodmeal. After the flies fed, the clamp was removed. The mouse ear was then immediately inoculated with 10 µl of 5 × 103, 1 × 104, or 2 × 104 parasites in 1× PBS, injected intradermally in the concave surface of the pinna, using a 28G needle. All sand flies from the vial were immediately dissected to check for blood. In Vivo Bioluminescence Imaging of a Luciferase-Expressing L. major Parasite Immediately following infection and then once each week following infection, Luciferin (d-Luciferin potassium salt, Xenogen, CA and Goldbio, St. Louis, MO), the luciferase substrate, was administered IP into the mice at a concentration of 200 mg/kg, 18 min before bioluminescence analysis, in the method used by Caridha et al. (2017). Animals were anesthetized using isoflurane (4% induction, 2.5% maintenance) in an induction chamber (MWI Veterinary Supply, Harrisburg, PA) and maintained on isoflurane in the imaging chamber for analysis. Emitted photons were collected by auto acquisition with a charge couple device camera (Perkin Elmer Spectrum IVIS Spectrum) using the medium resolution (medium binning) mode. Analysis was performed after defining a region of interest that delimited the surface of the affected area. Total photon emission from the ear pinna infected area was quantified with Living Image software (Xenogen Corporation, Almeda, CA), and results were expressed in numbers of photons/s. Lesion Clinical Development Mice were checked daily for the development of cutaneous lesions on the inoculated ear pinna for up to 150 d, and lesions were measured over time using a caliper instrument (Fisher Scientific, USA) with 0.1 mm sensitivity. Length and width measurements were taken to account for asymmetrical lesions. Lesion size area was then calculated using the π × r1 × r2 formula (where r1 = d1/2 and r2 = d2/2). Animals were euthanized when lesions averaged approximately 10–15 mm2 or at 150 d, in accordance with the AVMA Guidelines on Euthanasia (Leary et al. 2013), and tissues were collected for further characterization of organ parasite load. Tissue Analysis A segment of liver and spleen were cut lengthwise, and the cut surface was gently placed against a glass slide several times in sequence to make impression smears. These smears were fixed with methanol and then stained with 20% Geimsa for 5 min. Parasite burden was determined by Leishman Donovan Units (LDU), which is calculated as organ mass (g) multiplied by the number of amastigotes per 500 macrophages’ nuclei (Stauber et al. 1958). AmBisome Treatment Response Four mice with average lesion sizes with no statistical differences were treated with amphotericin B (AmBisome, Astellas Pharma, US Inc., Northbrook, IL) at 25 mg/kg IP each day for 12 treatments, and compared with an untreated control group, injected IP with sterile water vehicle daily for 12 treatments. IVIS signal was taken for both groups 24 h after the 5th, 10th, and 12th treatments. Statistics Five mice were randomly assigned to each Leishmania infection group. For the AmBisome treatment response, four mice were assigned in treatment and control groups in which the average lesion sizes were not statistically different. One-way ANOVA was used to determine statistical significance and provide confidence intervals for comparisons of multiple groups. A Student’s t-test was used when directly comparing two experimental groups. Drug efficacy in the treatment response screen was evaluated by calculating, respectively, the bioluminescence signal suppression and lesion size reduction in treated groups compared with the vehicle control treated group. A probability (P) level of <0.05 was accepted for the purpose of declaring that the treatments had statistically significant effects. Results Dermal Lesion Development Five mice were each inoculated with 5 × 103, 1 × 104, or 2 × 104L. major-LUC parasites intradermally into the ear (see Materials and Methods). Additional groups of five mice were treated with the same doses of parasites mixed with 0.2 pairs of sand fly SGS, or were exposed to sand fly bites (10 flies) immediately before the doses of parasites were injected (see Materials and Methods). For all inoculation combinations containing 2 × 104 parasites, papules developed on the ear by Day 35, and opened into ulcers by Day 40 post inoculation. For all inoculation combinations containing 1 × 104 parasites, papules developed by Day 38, and opened into ulcers by Day 41 post inoculation. These ulcers developed steadily in size over the observation period (Fig. 1). For all inoculation combinations containing 5 × 103 parasites, papules developed by Day 42, but did not open to ulcers or change in size within a 90-d observation period after inoculation. There was a statistically significant dose–response for uninfected fly bites plus parasites on Day 55 post infection (Fig. 1B; F = 4.22, df = 2, P = 0.04), for parasites plus SGS on Day 55 post infection (Fig. 1C; F = 7.25, df = 2, P = 0.009), and for parasites alone (Fig. 1D; F = 10.72, df = 2, P = 0.01). However, when comparing treatment groups for 2 × 104 parasites (Fig. 1A; parasites alone, parasites + SGS, parasites + uninfected fly bite), there was no statistical difference (F = 0.017, df = 2, P = 0.98). Fig. 1. View largeDownload slide Dermal lesion development in BALB/c mouse infected ID in the ear pinna with (A) 2 × 104 stationary-phase L. major-LUC promastigotes alone, with sand fly SGS, or with uninfected sand fly bites immediately prior to injection of parasites; (B) uninfected sand fly bites followed immediately by 5 × 103, 1 × 104, or 2 × 104 parasites; (C) parasites mixed with 0.2 pair salivary gland from sand flies; (D) parasites alone. Single-factor ANOVA (Dunnett’s test) was used to determine whether differences in group means existed across the experiment. Statistical significance (P < 0.05) is indicated by asterisk. Fig. 1. View largeDownload slide Dermal lesion development in BALB/c mouse infected ID in the ear pinna with (A) 2 × 104 stationary-phase L. major-LUC promastigotes alone, with sand fly SGS, or with uninfected sand fly bites immediately prior to injection of parasites; (B) uninfected sand fly bites followed immediately by 5 × 103, 1 × 104, or 2 × 104 parasites; (C) parasites mixed with 0.2 pair salivary gland from sand flies; (D) parasites alone. Single-factor ANOVA (Dunnett’s test) was used to determine whether differences in group means existed across the experiment. Statistical significance (P < 0.05) is indicated by asterisk. Luciferin Kinetics In the base-of-tail CL mouse model (Caridha et al. 2017), luciferin kinetics were performed in BALB/c mice to find the peak luminescence signal in minutes post luciferin administration on 18 d post infection. A similar kinetics study was performed on this ear infection model of L. major parasites on both Day 18 and Day 30 post infection. Luciferin was injected IP into each mouse at a dose of 200 mg/kg. Consecutive bioluminescence readings were taken every minute, starting at 8 min after luciferin administration, using the Perkin Elmer IVIS Spectrum. As shown in Fig. 2, at both 18 and 30 d post infection, luminescence signal sharply rose until approximately 18–20 min post injection. We measured all subsequent bioluminescent signals 18 min after luciferin was administered to keep consistent readings between models. Fig. 2. View largeDownload slide Luciferin kinetics in pinna with 2 × 104 luciferase-expressing stationary phase L. major promastigotes. Bioluminescence measurements were taken every 1 min starting at 8 min post luciferin administration. Data points represent the mean bioluminescent signal ± SEM for a total of six BALB/c mice. Two hundred mg/kg luciferin was administered IP in BALB/c mice infected in the ear dermis. Fig. 2. View largeDownload slide Luciferin kinetics in pinna with 2 × 104 luciferase-expressing stationary phase L. major promastigotes. Bioluminescence measurements were taken every 1 min starting at 8 min post luciferin administration. Data points represent the mean bioluminescent signal ± SEM for a total of six BALB/c mice. Two hundred mg/kg luciferin was administered IP in BALB/c mice infected in the ear dermis. Ambisome Response The activity of a known antileishmanial drug, amphotericin B (Aronson et al. 2016), was evaluated in this ear infection model for both lesion suppression and bioluminescent signal reduction. The bioluminescent signal in the infected ear pinna 24 h after the end of the treatment period had decreased by 96.6% compared with the untreated control group, and all lesions of the treatment group were clinically cured within 10 d after the end of the treatment period (Fig. 3). Fig. 3. View largeDownload slide The evolution of bioluminescent signal (A) and of lesion sizes (B) in BALB/c mice infected ID in the ear pinna with 2 × 104 stationary-phase L. major promastigotes and treated IP for 12 consecutive days with 25 mg/kg AmBisome is shown plotted against days post start of treatment. Bars represent the mean bioluminescent signal ± SEM for a total of four BALB/c mice. Single-factor ANOVA (Dunnett’s test) was used to determine whether differences in group means existed across the experiment. Statistical significance (P < 0.05) is indicated by asterisk. Fig. 3. View largeDownload slide The evolution of bioluminescent signal (A) and of lesion sizes (B) in BALB/c mice infected ID in the ear pinna with 2 × 104 stationary-phase L. major promastigotes and treated IP for 12 consecutive days with 25 mg/kg AmBisome is shown plotted against days post start of treatment. Bars represent the mean bioluminescent signal ± SEM for a total of four BALB/c mice. Single-factor ANOVA (Dunnett’s test) was used to determine whether differences in group means existed across the experiment. Statistical significance (P < 0.05) is indicated by asterisk. In Vivo Bioluminescence Signal Development and Limit of Detection In previous studies, we have determined that there is a very strong correlation between the bioluminescence signal and the parasite load at the infection site in the early stages post infection of BALB/c mice with luciferase-expressing L. major promastigotes (R2 = 0.96) (Caridha et al. 2017). Bioluminescent signal was measured as previously described (Caridha et al. 2017) in mice infected ID in the ear pinna with 2 × 104, 1 × 104, 5 × 103, and 1 × 103 luciferase-expressing L. major parasites. Immediately after infections, a measurable bioluminescent signal at the ear pinna was visible in 2/3 mice belonging to the group infected with 2 × 104 parasites and was still present in both mice 10 h post infections. As described in the literature, a several-week-long silent phase follows infections with low doses of L. major parasites (Belkaid et al. 2000). This period is characterized by a low but increasing number of parasites at the infection site (Belkaid et al. 2000). In our study, the parasite load stayed below the limit of detection in all infected groups until Day 19 post infections, when 3/3 mice belonging to the 2 × 104 parasite-infected group started showing a weak but measurable signal. Furthermore, the bioluminescent signal increased in all groups in a dose-dependent manner. A measurable bioluminescent signal was present on Day 37 in 3/3 and on Day 42 in 2/3 BALB/c mice infected, respectively, with 1 × 104 and 5 × 103L. major parasites. No bioluminescent signal was ever present at the infected site in the ear pinna of BALB/c mice infected with 1 × 103 stationary phase L. major parasites until Day 60 post infections which was the last day of the study (Fig. 4). Fig. 4. View largeDownload slide Evolution of bioluminescent signal in BALB/c mice infected with different concentrations of luciferase-expressing L. major parasites. Fig. 4. View largeDownload slide Evolution of bioluminescent signal in BALB/c mice infected with different concentrations of luciferase-expressing L. major parasites. In a consecutive experiment, groups of three mice were each infected with 2 × 104, 1 × 104, 5 × 103, or 1 × 103L. major parasites. Papules were present in the ear pinna in 3/3, 3/3, and 2/3 mice, respectively, on Days 35, 39, and 43 post infections. Lesion sizes grew in a dose-dependent manner. On Day 60 post infections, the average lesion sizes for the three groups mentioned earlier were, respectively, 15.09, 12.88, and 7.64 mm2 (data not shown). There were no lesions present in the BALB/c mice infected with 1 × 103L. major parasites, which suggests that the intensity of the bioluminescent signal (or lack thereof) in the early stages post infection can predict lesion sizes in the later stages of disease progression. In order to shorten the length of studies in our new model, we chose to infect the mice with 2 × 104 luciferase-expressing L. major promastigotes, where we detect a measurable bioluminescent signal as soon as Day 19 post infections and there are measurable lesions as soon as 30–35 d post infection. Effect of SGS and Fly Bite in the Evolution of Bioluminescent Signal with Different Doses Of Luciferase-Expressing L. major Promastigotes Three groups of five BALB/c mice were infected with 2 × 104L. major-LUC promastigotes in the presence of SGS or fly bite or neither as described in the Materials and Methods. One-way ANOVA and t-test show that there was no statistical significance in lesion sizes in all time points in all three groups in this study (data not shown). Organ Analysis Groups of five mice were infected ID in the ear pinna with 5 × 103, 1 × 104, or 2 × 104 stationary-phase L. major-LUC promastigotes alone, with sand fly salivary gland extract, or with uninfected sand fly bites immediately prior to injection of parasites. Liver and spleen were collected and analyzed for LDU, when the ear lesions within an infection group averaged 10–15 mm2. Single-factor ANOVA (Dunnett’s test) revealed statistically significant differences between the 5 × 103, 1 × 104, and 2 × 104 dose groups, but no significant difference with the addition SGS or fly feeding compared to promastigote injection alone in each dose group (Fig. 5). Fig. 5. View largeDownload slide LDU parasite burden in liver (A) and spleen (B). L. major parasite burden in liver and spleen were measured in BALB/c mice infected in the ear pinna with 5 × 103, 1 × 104, or 2 × 104 stationary-phase L. major-LUC promastigotes alone, with sand fly salivary gland extract, or with uninfected sand fly bites immediately prior to injection of parasites. Bars represent the mean LDU (organ mass (g) × number of amastigotes per 500 macrophage nuclei) ± SEM for a total of five BALB/c mice per group. Single-factor ANOVA (Dunnett’s test) was used to determine whether differences in group means existed across the experiment. Statistical significance (P < 0.05) is indicated by asterisk. Fig. 5. View largeDownload slide LDU parasite burden in liver (A) and spleen (B). L. major parasite burden in liver and spleen were measured in BALB/c mice infected in the ear pinna with 5 × 103, 1 × 104, or 2 × 104 stationary-phase L. major-LUC promastigotes alone, with sand fly salivary gland extract, or with uninfected sand fly bites immediately prior to injection of parasites. Bars represent the mean LDU (organ mass (g) × number of amastigotes per 500 macrophage nuclei) ± SEM for a total of five BALB/c mice per group. Single-factor ANOVA (Dunnett’s test) was used to determine whether differences in group means existed across the experiment. Statistical significance (P < 0.05) is indicated by asterisk. Discussion Leishmaniasis is a disease caused by parasites transmitted by sand flies, and there is a need for a natural infection model to assess antileishmanial drug efficacy. In a previous study (Belkaid et al. 1998), the addition of Phlebotomus papatasi SGS to L. major parasites was shown to increase the clinical dermal lesion development (compared with the vehicle control) of a CL mouse model. Studies (Belkaid et al. 1998, Abdeladhim et al. 2014) have shown that the salivary proteins from sand flies create an immune response that mimics a natural infection immune response to the L. major parasites compared with injection of parasites alone, and produce a faster and more destructive cutaneous lesion. In evaluating dermal lesions alone, there is extensive necrosis and dermal erosion in the BALB/c mouse’s ear by 2–3 mo post infection (Belkaid et al. 1998), which increases the need for shorter studies for drug screening. In addition, these studies are long and expensive. Schuster et al. (2014) reported that the use of bioluminescent parasites was an accurate approach to quantify Leishmania spp. parasites in the ear over time in a noninvasive model. In BALB/c mice, there is also a direct correlation between parasite load and the size of the ear lesion (Baldwin et al. 2003). This study and our previous study (Caridha et al. 2017) have also shown a very strong correlation between both the lesion size and bioluminescence signal and the parasite load at the infection site in the early stages post infection with luciferase-expressing L. major stationary-phase promastigotes in BALB/c mice. In the AmBisome lesion cure study, reduction of bioluminescent signal at the infection site as well as lesion size can be used as an experimental endpoint to measure drug efficacy. In our study with P. duboscqi sand flies and 5 × 103, 1 × 104, or 2 × 104L. major-LUC stationary phase promastigote parasites, we saw no statistical differences in dermal lesion development between L. major-LUC parasites alone, L. major-LUC parasites combined with salivary components, or with uninfected sand fly bites prior to injection. A prior study (Belkaid et al. 1998) used P. papatasi sand flies, and 1,000 L. major metacyclic promastigote parasites, which are the infective life stage of the parasite in vertebrates (World Health Organization 2010). When comparing the current study to that of Belkaid et al. (1998), the authors demonstrated that coinoculation of 1,000 L. major metacyclic parasites and 0.2 pair of P. papatasi SGS significantly exacerbated disease progression in the ear dermis of both BALB/c and B/6 mice. In contrast, our study failed to demonstrate any exacerbation when P. duboscqi SGS was coinoculated with 5 × 103, 1x × 04, or 2 × 104 stationary-phase L. major-LUC promastigotes in the mouse ear. In addition to using SGS, our study also used 10 uninfected sand flies to expose the mice to sand fly saliva prior to inoculation. It should be noted that the use of sand flies to expose mice to salivary compounds is more natural than using SGS. The amount of saliva injected into the ear by the flies, however, is not accurately quantifiable and is undoubtedly highly variable. It is very important to note when comparing our current study to that of Belkaid et al. (1998) that a different sand fly vector of L. major was selected. Even though both of species of sand fly (P. duboscqi and P. papatasi) can transmit the L. major parasite, different sand fly species have different components in their saliva (Abdeladhim et al. 2014). The different species of sand fly used may account for the difference in lesion development in our study versus the study by Belkaid et al. (1998). Due to the low mature infection rate achieved in P. papatasi in our laboratory and several other laboratories, we chose to use P. duboscqi since we can achieve high numbers of naturally infected P. duboscqi that can be used for further vector transmission studies. During this study, we did attempt to transmit L.major-LUC by the bite of infected P. duboscqi; however, transmission was not successful (data not shown). In previous studies in our laboratory, we have determined that our cultures of L. major-LUC parasites contain 7–9 % metacyclic promastigotes. Thus, when infecting with 2 × 104L. major-LUC stationary phase promastigote parasites, there were approximately 1,600 metacyclic phase parasites. This dose of parasites was the only infection group in which we consistently achieved 100% infection rates of the mice and could read bioluminescent signal in all the infected mice. In the lower concentration groups of parasites, 5 × 103 and 1 × 104 stationary-phase parasites contained approximately 800 and 400 metacyclic-phase parasites, respectively. These infection levels gave a more delayed response and sometimes not all the animals had measurable signals or developed lesions. The animals infected with 1 × 103 stationary phase parasites (approximately 80 metacyclic parasites) showed no response at all. This also follows the dose–response of natural infections (Kimblin et al. 2008). Kimblin et al. (2008) demonstrated that the number of parasites transmitted to a mouse ear by a single infected sand fly widely varies from <10 to nearly 100,000 parasites, but the majority of the individual sand flies transmitted <600 parasites to the mouse ear. Our study achieved 100% transmission only when using 1,600 metacyclic promastigote parasites, even when coinoculating with P. duboscqi SGS or exposing the mice to uninfected flies prior to inoculation. Ultimately, the use of infected sand flies would provide the best assay used to test treatments for Leishmania spp. infections. The difficulty in consistently and reliably obtaining infected sand flies that successfully and predictably transmit by bite has proved to be a major constraint. In our study, however, the use of 1,600 metacyclic parasites is within the range of the reported number of Leishmania spp. parasites transmitted during a natural sand fly transmission by Kimblin et al. (2008). In our study, the bioluminescent signal dropped below the limit of detection until approximately 20 d post infection in the mice infected with 2 × 104L. major-LUC parasites, indicating this silent phase of infection. A drug screen could be started by Day 35 post infections when all these mice had small ear dermal lesions. For a 10–12-d treatment, as noted in our AmBisome response, a successful treatment shows the greatest reduction in bioluminescent signal 1 d after treatment is complete. Prior lesion screens (Caridha et al. 2017) in both BALB/c mice and Golden Syrian Hamsters validated that approximately 10 additional days were required for the lesions to heal post-treatment at the base of the tail. Our ear infection model showed the same pattern of timing of bioluminescent signal reduction and lesion healing. For the parasite infection alone, this drug screen process would thus require 55 d post infection until the final lesion measurement and endpoints for both parasite load reduction and drug efficacy. In accordance with the three R’s of animal testing (Replacement, Reduction, and Refinement) (Russell and Burch 1959), the use of luciferase-expressing L. major parasites enables the use of longitudinal studies, thus ensuring a significant reduction in the number of animals used in antileishmanial drug discovery. In addition, it increases the accuracy of assessing infection progression and drug response, refines the drug screen process by being a noninvasive assessment method, and improves animal welfare by allowing earlier endpoints before dermal necrosis develops. The use of the mouse ear to infect and reliably monitor disease progression in vivo makes this model more effective for Leishmania drug treatment studies and it can be used to test compounds that have been determined as very effective in the other, high inoculum CL models. Material has been reviewed by the Walter Reed Army Institute of Research. There is no objection to its presentation and/or publication. The opinions or assertions contained herein are the private views of the author, and are not to be construed as official, or as reflecting true views of the Department of the Army or the Department of Defense. Acknowledgments Special thanks to Connie Schmitt, DVM, DACVPM for assistance with mouse dissections; and to Carrie Benton, DVM, DACLAM and Chad Black, DVM, PhD for their mentorship during this residency project. We also like to thank Amanda Schenk, Ivan Sanchez, John Goulart, Taylor Bowman, Sophia Kish, and Margarita Vidal for excellent technical assistance. References Abdeladhim, M., Kamhawi S., and Valenzuela J. G.. 2014. What’s behind a sand fly bite? The profound effect of sand fly saliva on host hemostasis, inflammation and immunity. Infect. Genet. Evol . 28: 691– 703. Google Scholar CrossRef Search ADS PubMed Aronson, N., Herwaldt B. L., Libman M., Pearson R., Lopez-Velez R., Weina P., Carvalho E. M., Ephros M., Jeronimo S., and Magill A.. 2016. Diagnosis and treatment of leishmaniasis: clinical practice guidelines by the Infectious Diseases Society of America (IDSA) and the American Society of Tropical Medicine and Hygiene (ASTMH). Clin. Infect. Dis . 63: e202– e264. Google Scholar CrossRef Search ADS PubMed Baldwin, T. M., Elso C., Curtis J., Buckingham L., and Handman E.. 2003. The site of Leishmania major infection determines disease severity and immune responses. Infect. Immun . 71: 6830– 6834. Google Scholar CrossRef Search ADS PubMed Belkaid, Y., Kamhawi S., Modi G., Valenzuela J., Noben-Trauth N., Rowton E. D., Ribeiro J., and Sacks D.. 1998. Development of a natural model of cutaneous leishmaniasis: powerful effects of vector saliva and saliva preexposure on the long-term outcome of Leishmania major infection in the mouse ear dermis. J. Exp. Med . 188: 1941– 1953. Google Scholar CrossRef Search ADS PubMed Belkaid, Y., Mendez S., Lira R., Kadambi N., Milon G., and Sacks D.. 2000. A natural model of Leishmania major infection reveals a prolonged “silent” phase of parasite amplification in the skin before the onset of lesion formation and immunity. J. Immunol . 165: 969– 977. Google Scholar CrossRef Search ADS PubMed Caridha, D., Parriot S., Hudson T. H., Lang T., Ngundam F., Leed S., Sena J., Harris M., O’Neil M., Sciotti R.,et al. 2017. Use of optical imaging technology in the validation of a new, rapid, cost-effective drug screen as part of a tiered in vivo screening paradigm for development of drugs to treat cutaneous leishmaniasis. Antimicrob. Agents Chemother . 61. doi: 10.1128/AAC.02048-16. Croft, S. L., Sundar S., and Fairlamb A. H.. 2006a. Drug resistance in leishmaniasis. Clin. Microbiol. Rev . 19: 111– 126. Google Scholar CrossRef Search ADS Croft, S. L., Seifert K., and Yardley V.. 2006b. Current scenario of drug development for leishmaniasis. Indian J. Med. Res . 123: 399– 410. Gamboa, D., Torres K., De Doncker S., Zimic M., Arevalo J., and Dujardin J. C.. 2008. Evaluation of an in vitro and in vivo model for experimental infection with Leishmania (Viannia) braziliensis and L. (V.) peruviana. Parasitology 135: 319– 326. Google Scholar CrossRef Search ADS PubMed Gomes-Silva, A., Valverde J. G., Ribeiro-Romao R. P., Placido-Pereira R. M., and Da-Cruz A. M.. 2013. Golden hamster (Mesocricetus auratus) as an experimental model for Leishmania (Viannia) braziliensis infection. Parasitology 140: 771– 779. Google Scholar CrossRef Search ADS PubMed Institute for Laboratory Animal Research. 2011. Guide for the Care and Use of Laboratory Animals , 8th ed. National Academies Press, Washington D.C. Karimkhani, C., Wanga V., Coffeng L. E., Naghavi P., Dellavalle R. P., and Naghavi M.. 2016. Global burden of cutaneous leishmaniasis: a cross-sectional analysis from the Global Burden of Disease Study 2013. Lancet Infect. Dis . 16: 584– 591. Google Scholar CrossRef Search ADS PubMed Kimblin, N., Peters N., Debrabant A., Secundino N., Egen J., Lawyer P., Fay M. P., Kamhawi S., and Sacks D.. 2008. Quantification of the infectious dose of Leishmania major transmitted to the skin by single sand flies. Proc. Natl Acad. Sci. USA 105: 10125– 10130. Google Scholar CrossRef Search ADS Kunzler, B. 2013. Cutaneous leishmaniasis: the efficacy of nonantimony treatment in the austere environment. Using cryotherapy, thermotherapy, and photodynamic therapy as an alternative method of treatment. J. Spec. Oper. Med . 13: 40– 45. Google Scholar PubMed Leary, S., Underwood W., Anthony R., Cartner S., Corey D., Grandin T., Greenacre C., Gwaltney-Brant S., McCrackin M., Meyer R.,et al. 2013. AVMA Guidelines for the Euthanasia of Animals . Schaumburg, IL. Lecoeur, H., Buffet P., Morizot G., Goyard S., Guigon G., Milon G., and Lang T.. 2007. Optimization of topical therapy for Leishmania major localized cutaneous leishmaniasis using a reliable C57BL/6 Model. PLOS Negl. Trop. Dis . 1: e34. Google Scholar CrossRef Search ADS PubMed Lecoeur, H., Buffet P. A., Milon G., and Lang T.. 2010. Early curative applications of the aminoglycoside WR279396 on an experimental Leishmania major-loaded cutaneous site do not impair the acquisition of immunity. Antimicrob. Agents Chemother . 54: 984– 990. Google Scholar CrossRef Search ADS PubMed Mears, E. R., Modabber F., Don R., and Johnson G. E.. 2015. A review: the current in vivo models for the discovery and utility of new anti-leishmanial drugs targeting cutaneous leishmaniasis. PLOS Negl. Trop. Dis . 9: e0003889. Google Scholar CrossRef Search ADS PubMed Milon, G., Del Giudice G., and Louis J. A.. 1995. Immunobiology of experimental cutaneous leishmaniasis. Parasitol. Today 11: 244– 247. Google Scholar CrossRef Search ADS PubMed Modi, G., and Rowton E. D.. 1999. Laboratory maintenance of phlebotomine sand flies . Science Publishers Inc., New Hampshire. Ribeiro-Romao, R. P., Moreira O. C., Osorio E. Y., Cysne-Finkelstein L., Gomes-Silva A., Valverde J. G., Pirmez C., Da-Cruz A. M., and Pinto E. F.. 2014. Comparative evaluation of lesion development, tissue damage, and cytokine expression in golden hamsters (Mesocricetus auratus) infected by inocula with different Leishmania (Viannia) braziliensis concentrations. Infect. Immun . 82: 5203– 5213. Google Scholar CrossRef Search ADS PubMed Robledo, S. M., Carrillo L. M., Daza A., Restrepo A. M., Munoz D. L., Tobon J., Murillo J. D., Lopez A., Rios C., Mesa C. V.,et al. 2012. Cutaneous leishmaniasis in the dorsal skin of hamsters: a useful model for the screening of antileishmanial drugs. J. Vis. Exp . doi: 10.3791/3533. Roy, G., Dumas C., Sereno D., Wu Y., Singh A. K., Tremblay M. J., Ouellette M., Olivier M., and Papadopoulou B.. 2000. Episomal and stable expression of the luciferase reporter gene for quantifying Leishmania spp. infections in macrophages and in animal models. Mol. Biochem. Parasitol . 110: 195– 206. Google Scholar CrossRef Search ADS PubMed Russell, W. M. S., and Burch R. L.. 1959. The principles of humane experimental technique . Methuen, London, United Kingdom. Schuster, S., Hartley M. A., Tacchini-Cottier F., and Ronet C.. 2014. A scoring method to standardize lesion monitoring following intra-dermal infection of Leishmania parasites in the murine ear. Front. Cell. Infect. Microbiol . 4: 67. Google Scholar CrossRef Search ADS PubMed Stauber, L. A., Franchino E. M., and Grun J.. 1958. An eight-day method for screening compounds against leishmania donovani in the golden hamster. J. Protozool . 5: 269– 273. Google Scholar CrossRef Search ADS Thalhofer, C. J., Graff J. W., Love-Homan L., Hickerson S. M., Craft N., Beverley S. M., and Wilson M. E.. 2010. In vivo imaging of transgenic leishmania parasites in a live host. J. Vis. Exp . doi: 10.3791/1980. World Health Organization. 2010. Control of the leishmaniases, WHO Technical Report Series . World Health Organization, Geneva, Switzerland. Published by Oxford University Press on behalf of Entomological Society of America 2017. This work is written by (a) US Government employee(s) and is in the public domain in the US.
Journal of Medical Entomology – Oxford University Press
Published: Mar 1, 2018
It’s your single place to instantly
discover and read the research
that matters to you.
Enjoy affordable access to
over 18 million articles from more than
15,000 peer-reviewed journals.
All for just $49/month
Query the DeepDyve database, plus search all of PubMed and Google Scholar seamlessly
Save any article or search result from DeepDyve, PubMed, and Google Scholar... all in one place.
Get unlimited, online access to over 18 million full-text articles from more than 15,000 scientific journals.
Read from thousands of the leading scholarly journals from SpringerNature, Elsevier, Wiley-Blackwell, Oxford University Press and more.
All the latest content is available, no embargo periods.
“Hi guys, I cannot tell you how much I love this resource. Incredible. I really believe you've hit the nail on the head with this site in regards to solving the research-purchase issue.”Daniel C.
“Whoa! It’s like Spotify but for academic articles.”@Phil_Robichaud
“I must say, @deepdyve is a fabulous solution to the independent researcher's problem of #access to #information.”@deepthiw
“My last article couldn't be possible without the platform @deepdyve that makes journal papers cheaper.”@JoseServera