C3/CD11b-Mediated Leishmania major Internalization by Neutrophils Induces Intraphagosomal NOX2-Mediated Respiratory Burst but Fails to Eliminate Parasites and Induces a State of Stalled Apoptosis

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Trypanosomatid protozoan parasites of the genus Leishmania are the causative agent of a cluster of related diseases in mammalian hosts called leishmaniasis (1). These parasites are transmitted through the bite of a female hematophagic sand fly. During blood feeding, a dermal wound and blood pool forms, into which metacyclic promastigotes, the infectious life cycle stage of the parasite, are deposited (2, 3). This wound initiates robust recruitment of neutrophils, and Leishmania–neutrophil interactions are a defining feature of the first few hours of infection (3, 4). In the “Trojan horse” model of infection proposed by Laskay et al. (5), Leishmania parasites survive traditional neutrophil defense mechanisms such as NETosis (6, 7) and NADPH oxidase isoform 2 (NOX2) (Phox)–mediated production of reactive oxygen species (ROS) and establish an intracellular infection in a parasite-containing phagolysosome, often referred to as the parasitophorous vacuole (8, 9). Leishmania is then thought to induce and/or pause a form of noninflammatory programmed cell death (PCD) called apoptosis (10–12) and subsequently parasitize neutrophil efferocytosis, the anti-inflammatory process by which phagocytic mononuclear cells clear dead and dying apoptotic neutrophils or neutrophil-derived apoptotic bodies (5, 13). This strategy would facilitate infection of long-lived phagocytes without inducing an antiparasitic response initiated by direct parasite uptake. In vivo observations in which the presence or absence of neutrophils is manipulated experimentally at the time of infection support this proparasitic role for neutrophils (3, 14, 15).

An important consideration in initial Leishmania–neutrophil interactions is the role of the complement system of serum proteins in the blood pool into which the parasite is deposited. Spontaneous C3 hydrolysis and deposition of C3b on the parasite surface is the predominant complement pathway triggered by Leishmania (16–21). However, C3b deposition does not lead to the efficient formation of the membrane attack complex (22–24). Rather, the Leishmania virulence factors metalloproteinase gp63 and the plasma serine protease factor 1 (SPF-1) convert C3b to inactive C3b (iC3b) on the parasite surface, which is highly opsonic, and, in classical studies employing macrophages, iC3b opsonization facilitates internalization of parasites via the phagocyte C3b receptor, CD11b (also referred to as complement receptor 3 [CR3] or macrophage-1 Ag [MAC-1]) (25–38). In macrophages, the mechanism of Leishmania internalization has been proposed to have significant consequences for parasite survival, with iC3b/CD11b facilitating “silent entry” into macrophages, decoupled from the production of microbicidal NOX2-linked ROS in the phagolysosome (34–37) versus uptake by non–iC3b/CD11b-mediated mechanisms, resulting in “activated entry,” triggering macrophage immune effector functions and contributing to ROS-dependent parasite killing (39). Although the role of C3/CD11b during Leishmania infection has been investigated previously, largely in susceptible BALB/c mice (40–42), the mechanisms by which neutrophils internalize parasites have not been characterized apart from observations that uptake happens rapidly in vitro in the presence of serum (43–45).

In addition to the role of complement, the efficacy of NOX2-dependent ROS in neutrophil-mediated parasite elimination following phagocytosis is also unclear. This may be because studies on the role of ROS are largely conducted employing macrophages and predominantly employ parasites derived from stationary phase cultures versus purified metacyclic promastigotes (46–49). Stationary phase populations are likely more susceptible to ROS-mediated elimination versus metacyclic promastigotes given their lower surface expression of lipophosphoglycan (LPG) and gp63 at the population level, both of which are associated with increased resistance to ROS-mediated killing (8, 9, 22, 37). Previously, we found that NOX2-dependent ROS production had no impact on Leishmania amazonensis parasite numbers in vivo (50). Rather, NOX2 deficiency resulted in a shift from apoptosis to necrosis during the late stages (≥10 h postinfection [p.i.]) of neutrophil infection as defined by decreased cell surface expression of PS, a process that occurs when the inner PS-containing leaflet of the plasma membrane becomes exposed on the outer cell membrane during apoptosis, and increased uptake of the cell death dye 7-aminoactinomycin D (7-AAD). Increased neutrophil necrosis was associated with increased inflammation and dermal lesions despite equivalent parasite loads (50). Several studies have suggested that Leishmania-infected neutrophils have an altered cell death program (10–13, 51–53) including promoting apoptosis as indicated by PS externalization (10, 13) but also suppressing apoptosis as suggested by decreased enzymatic activity of caspase 3 (11, 12). Apoptosis can be triggered via engagement of both extracellular and intracellular receptor pathways (54). In each case, the process of apoptosis engages caspases, proteases that regulate PCD (55, 56). Activation of either pathway converges on the activation of shared executioner caspases existing within the cytoplasm as dimeric inactive zymogens (55, 57, 58). The most well-studied executioner caspase is caspase 3, which becomes active through proteolytic cleavage of procaspase 3 to expose the mature caspase 3 active site, which is capable of catalytically degrading cell maintenance proteins involved in regulating the cell cycle and DNA synthesis (58, 59), resulting in apoptotic PCD, including cell shrinkage, cytoplasm condensation, chromatin condensation, nuclear fragmentation, flipping of inner leaflet–associated “eat me” signals such as PS to the cell surface, and budding of apoptotic bodies (58, 60).

The factors involved in initial parasite uptake, induction and localization of ROS, and the subsequent impact on neutrophil viability are likely important to understanding how the parasite establishes infection. This information is relevant to vaccination, as neutrophils compromise the efficacy of Leishmania vaccine-induced CD4⁺ Th1 cell–mediated immunity, and this adaptive response must operate in the context of sand fly–mediated neutrophil recruitment (3, 15, 61–64). In the present work, we propose that Leishmania infection induces a state of stalled apoptosis that may aid parasite survival by facilitating the subsequent transition of the parasite from neutrophils to long-lived mononuclear phagocytic host cells via efferocytosis.

C57BL/6J, gp91^PHOX−/− (B6.129S-Cybb^tm1Din/J), C3^−/− (B6.129S4-C3^tm1Crr/J), and CD11b^−/− (B6.129S4-Itgam^tm1Myd/J) mice were acquired from The Jackson Laboratory and maintained under specific pathogen-free conditions in the Animal Resource Center at the University of Calgary Cumming School of Medicine. All animal procedures were performed in accordance with Canadian Council for Animal Care guidelines and were approved by the Health Sciences Animal Care Committee at the University of Calgary (protocol no. AC19-007).

The FV1 Leishmania major (MHOM/IL/80)–red fluorescence protein (RFP)–expressing parasite (via a Geneticin resistance gene/RFP gene–containing plasmid) was generated as previously described (65). Parasites were grown at 26°C in medium 199 supplemented with 20% heat-inactivated (HI) FCS (Gemini Bio-Products, San Francisco, CA), 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). L. major-RFP⁺ parasite cultures were supplemented with 10 μl of 50 mg/ml Geneticin (Life Technologies, 10131035) antibiotic as a precaution to maintain the plasmid.

Cultures were incubated at 26°C until a high proportion of parasites were active, fast-swimming, infective-stage metacyclic promastigotes. Metacyclic promastigotes were isolated from stationary cultures (4–5 d old) in Ficoll gradient as described previously (66).

Purified metacyclic promastigotes were resuspended in 1 ml of HBSS (Mg⁻/Ca⁻), then washed with HBSS (Mg⁻/Ca⁻) and centrifuged at 4000 rpm for 15 min (20°C). After centrifugation, the supernatant was aspirated and the pellet was resuspended in 1.5 ml of HBSS (Mg⁺/Ca⁺, i.e., sufficient for magnesium and calcium ions). Then, 75 μl of serum was added to the solution and mixed gently by finger flicking. The mixed parasite/serum suspension was placed into a preheated 37°C water bath for 30 min. Excess complete M199 medium was added, and the suspension was spun at 500 rpm for 5 min (20°C) to remove potential aggregates. The supernatant was then collected in a 50-ml Falcon tube and spun at 4000 rpm for 15 min (20°C) to pellet the opsonized metacyclic promastigotes. Finally, the pellet was resuspended in complete RPMI 1640 and parasites were quantified employing a hemocytometer.

Bone marrow was flushed from the tibias and femurs of euthanized mice onto a 70-μm cell strainer (VWR, 10199657) inserted into the top of a 50-ml Falcon tube using cold, sterile PBS loaded into a 10-ml syringe (BD Biosciences, 302995) tipped with a 27.5G needle (BD Biosciences, 305109). Bone marrow was strained through the 70-μm mesh using mechanical force applied by the plunger of a sterile syringe and flushed with cold, sterile PBS.

Bone marrow–derived cells were pelleted and resuspended in 1 ml of ACK (ammonium-chloride-potassium) lysis buffer (Life Technologies, A1049201) and incubated for 5 min at room temperature to lyse the RBCs. The solution was washed with 9 ml of cold HBSS (Life Technologies, 14175095) and resuspended in 2.5 ml of cold DMEM (Life Technologies, 11885084) and laid over 3.0 ml of Ficoll-Paque Plus (1.077 g/ml) (GE Healthcare, 17-1440-02) in a 15-ml Falcon tube. The Ficoll gradient column was centrifuged at 400 rcf for 30 min (20°C) with the acceleration speed set to 4/10 and the deceleration speed set to 2/10. Purified neutrophils were isolated by aspirating as much supernatant as possible while leaving the pellet completely undisturbed and then washing immediately with HBSS (Ca⁻/Mg⁻) to remove Ficoll. Finally, the purified neutrophil pellet was resuspended in an appropriate volume of complete RPMI 1640.

Parasites and neutrophils were cocultured at the indicated ratios and times at 37°C and 5% CO₂. Samples for all experiments were constructed to maintain a constant concentration equal to 1 million neutrophils per 1000 μl of total sample volume. For time periods up to and including 2–3 h, neutrophils were analyzed directly without a parasite washout step. For time periods >2–3 h, an excess of PBS was added to cocultures and the sample was spun at 1500 rpm (526 rcf) for 10 min to pellet the neutrophils and remove uninternalized parasites. The neutrophil pellet was then resuspended in a volume of complete RPMI 1640 equivalent to the initial coincubation volume and the sample was incubated at 37°C (5% CO₂) for the remaining duration of the experiment. In all cases, prior to preparation for flow or Western blot analysis, parasites were washed to remove uninternalized parasites during the preparation process.

All flow cytometric data were collected employing unfixed, stained neutrophils suspended in 5 ml polystyrene FACS tubes (Falcon, 352008) using a FACSCanto flow cytometer (BD Biosciences) controlled by BD FACSDiva software (version 8.0) (BD Biosciences). In some experiments (Fig. 1A–D) data were collected using a Cytek Aurora (Cytek Biosciences). Briefly, samples were washed with an excess of cold PBS and spun at 1500 rpm for 10 min (4°C) to remove culture media. Samples were then resuspended in 50 μl of cold FACS buffer containing (1:200) Fc Block (BD Biosciences) and (1:500) viability dye (BD Biosciences) and then incubated for 30 min at 4°C. After 30 min of incubation, 50 μl of FACS buffer containing the neutrophil surface staining mixture (1:400 anti-CD11b-PerCP-Cy5.5 [BD Biosciences], 1:200 anti-Ly6C-allophycocyanin-Cy7 (BD Biosciences), 1:200 anti-Ly6G-AF700 [BD Biosciences]) was added, homogenized, and incubated at 4°C for 20 min. Next, the samples were washed with 1 ml of cold FACS buffer and spun at 1500 rpm for 10 min (4°C). Finally, the samples were resuspended in 200 μl of FACS buffer and transferred to polystyrene FACS tubes and run on the FACSCanto as described while taking care to avoid sample exposure to light and maintain a cold sample temperature. To visualize neutrophil events, the following thresholds were used: forward scatter of ∼110 and side scatter of ∼390. Compensation was performed using single-stained neutrophils. Flow cytometric analysis was performed using FlowJo analysis software (version 10.6) and consisted of verifying compensations followed by sequentially gating for singlets, live versus dead cells (depending on needs of analysis), CD11b⁺ cells, Ly6G⁺/Ly6C^int neutrophils, RFP⁺ versus RFP⁻ neutrophils, and then analyzing for population characteristics unique to specific experimental questions (e.g., phosphatidylserine [PS] expression or ROS production). Analyzed data were exported to Excel (Microsoft) and then entered in to GraphPad Prism (versions 8 and 9) for the generation of figures and statistics.

Samples were then resuspended in 200 μl of complete RPMI 1640 containing 20 ng/ml dihydrorhodamine 123 (DHR123)-FITC (Invitrogen) fluorescent ROS probe and incubated for 5 min at 37°C and then washed with 2 ml of cold FACS buffer to remove excess probe. After being spun at 1500 rpm for 10 min (4°C), samples were resuspended in 50 μl of cold FACS buffer and stained as described above. Fluorescence minus one (FMO) was employed as a gating control.

Samples were first stained for surface markers. Next, the samples were washed with 1 ml of cold PBS and spun at 1500 rpm for 10 min (4°C) prior to being resuspended in 50 μl of cold 1× annexin V binding buffer (BD Biosciences, 556454; diluted 1:10 in cold double-distilled H₂O) containing 2 μl/sample annexin V-allophycocyanin (BD Biosciences) and incubated in the dark for 20 min at room temperature. Finally, the samples were washed with 1 ml of 1× annexin V binding buffer, centrifuged at 1500 rpm for 10 min (4°C) prior to being resuspended in 200 μl of 1× annexin V binding buffer, and transferred to polystyrene FACS tubes for data acquisition. FMO employing media without Ca/Mg was used as a gating control.

Cytospins were performed employing a Shandon Cytospin 4 cytocentrifuge (Thermo Fisher Scientific) using a single cytology funnel filter card (Simport, M965FW) between the cytocentrifuge cytology funnel and Superfrost Plus microscope slides (VWR, CA45311703) within the supplied stainless steel funnel clip. Samples were washed with cold PBS and resuspended in 100 μl of cold HBSS (Ca⁻/Mg⁻) containing 10% HI FBS. Prior to loading the experimental samples, the cytospin funnels were prewetted with 100 μl of PBS and the machine was run for 1 min at 1500 rpm (fast acceleration setting) to wet the filter paper. Samples were loaded into the cytospin funnels and centrifuged at 1500 rpm for 5 min (fast acceleration setting). Slides were left to sit at room temperature in the biological safety cabinet until completely dry (∼30 min).

Dried slides were fixed and stained using a three-step Wright–Giemsa staining kit for hematology (Quick III, VWR, 10143-226) by successively bathing the slides for 30 s in the methanol-based fixative solution, followed by 30 s of bathing in solution 1, and finally 30 s in solution 2 (which contains an equilibrium of methylene blue trihydrate and demethylated azure B to counterstain the eosin Y and allows for the visualization of nuclei). Excess fixative/stain was removed from the slide after each bathing step by gently dabbing the bottom corner of the microscope slide on to a dry paper towel. After completion of the staining procedure, the slides were washed with double-distilled H₂O. The fixed and stained slides were air dried and microscope slide covers were adhered over the dried sample using Entellan new rapid mounting medium (MilliporeSigma, 107961).

Samples were washed with an excess of cold PBS, and lysis of the pellet was performed on ice for 10 min using 80 μl of RIPA lysis buffer + cOmplete mini protease inhibitor mixture (Roche, 11836153001) per 1 million cells. After lysis, the lysate was transferred to a 1.5-ml Eppendorf tube. Twenty-five microliters of 5× sample buffer containing 0.5 M DTT (Roche, 10197777001) was then added and the lysate was placed in a 95°C heating block for 10 min. Halfway through the heating step, the samples were quickly homogenized by finger flicking. Precipitation generated during the heating step was spun back into the lysate suspension by room temperature centrifugation at 4000 rpm for 10 min. Typically, the prepared samples were then frozen at −20°C overnight

Samples were then separated using a 12% SDS-PAGE resolving gel employing the Bio-Rad Mini-PROTEAN Tetra cell system (Bio-Rad) along with PageRuler prestained protein ladder (10–180 kDa) (Thermo Fisher Scientific, 26616). Protein samples were run through the stacking gel at 100 V for ∼12 min and then the voltage was increased to 180 V and the proteins were resolved for ∼45 min. Once resolved, the proteins were wet transferred from the resolving gel to a nitrocellulose membrane (Amersham Protran premium 0.2 µm NC, GE Healthcare, 10600004) employing a mini trans-blot central core electrode assembly (Bio-Rad, 1703812). Gel-to-membrane protein transfer was run at 100 V for 45 min in ice-cooled, SDS⁻ transfer buffer.

Blots were then blocked for 1 h at room temperature under gentle agitation on a Belly Button orbital shaker platform using a solution of 1× TBS containing 0.1% Tween 20 (TBST) and 5% (w/v) powdered skim milk (Carnation, instant skim milk powder). Diluted primary Abs were diluted in 5% TBST-milk and were incubated at 4°C under gentle agitation for 1–3 d. Blots were washed five to six times for 5–10 min each with 0.1% TBST under vigorous agitation at room temperature. Secondary HRP-conjugated Ab suspensions in 5% TBST-milk were poured over the appropriate blots and incubated at room temperature for 1 h under gentle agitation. Blots were then washed five to six times for 5–10 min each with 0.1% TBST under vigorous agitation at room temperature. Primary and secondary Abs used to conduct Western blotting analyses are summarized as follows: anti-Caspase 3 (CST 9662, polyclonal rabbit 1:1000), anti-caspase 8 (CST 4927, polyclonal rabbit 1:1000) anti-cleaved. Caspase 8 (CST 8592, clone D5B2 rabbit 1:1000), anti-caspase 9 (CST 9508, clone C9 mouse 1:1000), anti-β-actin (CST 3700, clone 8H10D10 mouse 1:1000), anti-GAPDH (CST 5174, clone D16H11 rabbit 1:1000), anti-mouse IgG-HRP (Sigma-Aldrich A4416, polyclonal goat 1:4000), and anti-rabbit IgG-HRP (Sigma-Aldrich A0545, polyclonal goat 1:5000).

Imaging of Ab-labeled proteins was achieved using ECL Prime (Amersham, 45-002-401) and detected using various chemiluminescent signal accumulation mode settings (e.g., signal accumulation, high-sensitivity signal accumulation, or high-resolution signal accumulation) of a ChemiDoc MP imaging system (Bio-Rad) controlled by Image Lab software (version 5.1) (Bio-Rad) until signal saturation (signal intensity beyond a measurable range) was detected for any one of the target proteins. An image of the blot’s protein ladder was captured using the machine’s colorimetric mode to superimpose the ladder onto an image of the fully accumulated protein signal during post hoc analysis.

Analysis of Western blots was conducted using Image Lab version 6 (Bio-Rad). Detection of the bands allowed for semiquantitative analysis of the Western blot results by exporting the adjusted volume (i.e., the sum of all intensities recorded within the band boundaries subtracted by the background intensity) value for each protein-of-interest band and housekeeping protein band into Excel (Microsoft) for signal normalization. Normalized signal intensity values were plotted using GraphPad Prism (versions 8 and 9) (GraphPad, San Diego, CA).

Fresh bone marrow–isolated neutrophils, that is, noninfected or infected neutrophils from C57BL/6 WT mice, were lysed, after 1 and 3 h p.i., for 10 min using guanidine isothiocyanate–containing RLT lysis buffer (Qiagen, 79216). The lysate samples were then passed through QIAshredder columns, and RNA was purified using the RNeasy mini kit according to the manufacturer’s protocol (Qiagen). Reverse transcription was performed using a high-capacity cDNA reverse transcriptional kit (Thermo Fisher Scientific). Real-time PCR was performed on an ABI Prism 7900 sequence detection system (Applied Biosystems). The results were analyzed by the comparative threshold cycle method using 2^−ΔΔCt to determine the fold increase. Each gene was normalized to the 18S rRNA endogenous control and to “fresh” neutrophils direct isolated from the bone marrow. Transformed ΔΔCt values were then plotted using GraphPad Prism (versions 8 and 9) (GraphPad, San Diego, CA). Samples for each condition were run at least as triplicates. The TaqMan probes used were as follows: casp3 (Mm01195085_m1), casp8 (Mm01255716_m1), casp9 (Mm00516563_m1), bcl2 (Mm00477631_m1), mcl1 (Mm01257351_g1), and 18s rRNA (Mm03928990_g1).

Naive mice were challenged with 1.5 × 10⁵L. major-RFP metacyclic promastigotes intradermally in the ear in a volume of 10 μl. At 90 min following inoculation, ears were removed and placed in 70% ethanol at room temperature for 2–5 min and then allowed to dry. Separated dorsal and ventral sheets of ears were then incubated at 37°C for 90 min in 1 ml of DMEM containing 16 μg/ml Liberase. Following Liberase treatment tissue was homogenized for 3.5 min in a Medicon using a Medimachine (BD Biosciences). The tissue homogenate was then flushed from the Medicon with 10 ml of RPMI 1640 media containing 0.05% DNase I and filtered using a 50-μm pore size cell strainer to generate a single-cell suspension. Cells were then employed for further analysis. In some experiments, mice were exposed to the bites of infected sand flies, as described (67). Adult Phlebotomus duboscqi, Strain Mali, NR–50159 were provided by Walter Reed Army Institute of Research for distribution by BEI Resources, NIAID, NIH.

Two-photon intravital imaging of in vitro neutrophil (LysM-GFP)/L. major-RFP cocultures was performed using an inverted LSM 510 NLO multiphoton microscope (Carl Zeiss Microimaging) enclosed in an environmental chamber that was maintained at 30°C. This system had been custom fitted with three external non-descanned photomultiplier tube detectors in the reflected light path. Images were acquired using either a ×20/0.8 air objective or a ×25/0.8 numerical aperture water immersion objective. Fluorescence excitation was provided by a Chameleon XR Ti:Sapphire laser (Coherent) tuned to 920 nm for enhanced GFP and RFP excitation. Voxel dimensions were 0.64 × 0.64 × 2 µm using the ×20 objective and 0.36–0.51 × 0.36–0.51 × 2 µm using the ×25 objective. Raw imaging data were processed with Imaris (Biplane) using a Gaussian filter for noise reduction. All images are displayed as two-dimensional maximum intensity projections. For imaging experiments, L. major-RFP parasites (1 × 10⁵/ml in 200 μl) were prepared as described and allowed to adhere to a glass chamber slide for 30 min, excess parasites were removed by two washes with PBS, imaging was started, and purified neutrophils from LysM-GFP mice (1 × 10⁵/ml in 200 μl) were added.

Neutrophils were plated in complete RPMI 1640 in an eight-well ibidi imaging dish (catalog no. 80826) at a concentration of 10⁶ neutrophils per 1000 μl. Promastigotes were added directly to the dish at the indicated multiplicity of infection followed by 50 μl of a 10 mg/ml (in water) NBT solution. The plate was incubated at 37°C, 5% CO₂ for 1 h and then imaged live on a Leica SP5 confocal microscope.

Most of the figures presented depict representative data collected in vitro using populations of genetically identical neutrophils (collected and/or pooled from one or two mice) and split into three or more technical replicates unless otherwise indicated. A minimum of two fully independent experiments were conducted for each set of observations. Each dataset was first tested for normality using a Shapiro–Wilk test. Information regarding the specific parameters of each statistical analysis performed can be found within the figure legends. In brief, comparisons between two sets of nonparametric data were analyzed either by a two-tailed Mann–Whitney unequal variances t test or by a two-tailed Wilcoxon matched pairs rank test depending on whether the dataset had equal variances or were nonnormally distributed, respectively. For all tests, p ≤ 0.05 was considered significant. When modeling the relationship between a suspected explanatory variable and a suspected dependent variable, a linear regression was performed to assess the strength of the relationship between the two variables using the R² goodness-of-fit test where R² = 1 represents a perfect linear relationship between the two variables. Error bars depict SD unless otherwise noted. Statistical analyses were performed using GraphPad Prism (versions 8 and 9) (GraphPad, San Diego, CA). Statistical significance is indicated as follows: ns, p > 0.05; *p ≤ 0.05, **p ≤ 0.005, ***p ≤ 0.0005, and ****p ≤ 0.00005.

The primary rationale for a close study of neutrophil–Leishmania interactions is evidence that neutrophils phagocytose most L. major parasites immediately following infected sand fly bite (3, 68). This conclusion is based on two-photon intravital microscopy (2P-IVM) analysis of small numbers of sand fly–transmitted L. major-RFP parasites (<10) with supporting evidence from flow cytometric analysis of CD11b⁺ phagocytic cells in the skin at acute time points (1–10 h) following intradermal needle inoculation with ≥1 × 10⁴ parasites. Under these latter conditions, the vast majority of infected CD11b⁺RFP⁺ cells are Ly6G⁺Ly6C^int neutrophils, and these have also been characterized as lysozyme-high cells employing LysM-GFP reporter mice (Fig. 1A, 1B) (3, 10, 69, 70). Neutrophils are defined as Ly6C intermediate in the context of the Ly6C high expression found on monocytes (70, 71).

Detecting RFP⁺ infected cells by flow cytometric analysis at early time points following sand fly transmission is challenging, likely because only 2.3% of experimentally infected flies will transmit the ≥1 × 10⁴ dose of parasites required to reliably identify and enumerate infected cells by flow cytometry (65, 68). Although more recent flow cytometric analysis following sand fly transmission of a more virulent strain of L. major, L. major-RYN, has been more successful (72), we attempted to substantiate our original observations with the healing L. major-FV1 strain by exposing ears to the bites of 10–15 L. major-RFP–infected sand flies, with the intention to capture a bite site containing larger numbers of parasites (≥10) amenable to 2P-IVM and semiquantitative analysis early after transmission. Employing needle inoculation of LysM-GFP mice, we confirmed that RFP⁺LysM-GFP^hi infected cells visible by 2P-IVM are virtually all Ly6G⁺Ly6C^int neutrophils (Fig. 1C, leftmost bar) and that the vast majority of total live RFP⁺ cells are CD11b⁺Ly6G⁺Ly6C^int or CD11b⁺LysM^hi (Fig. 1D) (3). Following infected sand fly exposure we analyzed a bite site with high numbers of inoculated parasites that was amenable to 2P-IVM (Fig. 1E–G). Qualitative analysis of the x,y images collected across the z plane of the bite site shown in Fig. 1E revealed hundreds of RFP⁺ parasites within LysM-GFP^hi neutrophils (Fig. 1F, Supplemental Video 1). Quantitative imaging analysis of the number of RFP⁺ parasites shown in Fig. 1E that were tightly associated with LysM-GFP^hi cells (white dots), LysM-GFP^int cells (green dots), or no GFP association (LysM-GFP⁻, red dots) revealed that of the 1788 total parasites in the field of view, 1463 (81.8% of total parasites; 96.4% of GFP associated parasites) were very likely inside a LysM-GFP^hi neutrophil (Fig. 1G), substantiating with a large number of sand fly inoculated parasites that neutrophils are the initial host cell for most L. major-FV1 parasites following sand fly transmission.

To determine the relative contribution of C3 and CD11b to L. major phagocytosis by neutrophils in vivo, we needle inoculated WT, C3^−/−, or CD11b^−/− mice (38, 40–42, 73) (Fig. 1H–K). Importantly, we employed purified metacyclic promastigotes for infection, as this form of the parasite is the infectious life cycle stage of the parasite transmitted by the sand fly (2). This is in contrast to employing stationary phase culture-derived parasites that contain procyclic promastigotes that lack a fully developed and elongated LPG and gp63-enriched glycocalyx and are more susceptible to complement-mediated lysis (74).

Our analysis revealed that C3 did not play an overt role in acute neutrophil recruitment but was required for optimal parasite uptake as indicated by a 3.1-fold decrease in the total number of infected neutrophils and a significantly lower frequency of neutrophils at the dermal site with at least one parasite based on RFP expression at 90 min p.i. (Fig. 1H, 1J). The slight decrease in total neutrophil numbers in C3^−/− mice shown in Fig. 1J (left panel) never reached statistical significance in four separate experiments. We also found that CD11b^−/− mice had equivalent numbers of total neutrophils at the infection site, a 3.3-fold decrease in total RFP⁺ neutrophils, and a significantly lower frequency of RFP⁺ neutrophils (Fig. 1I, 1K). These observations suggest that the C3/CD11b pathway is the initial and/or primary means by which neutrophils phagocytose Leishmania parasites in vivo. We found a large variation in the number of infected neutrophils at 90 min p.i. This is because maximum uptake takes ∼10 h (10, 70, 71), and therefore the variation is likely a representation of varying kinetics of recruitment and phagocytosis in individual ears.

Employing C3^−/− mice, we were also able to determine the long-term impact of the absence of C3 on infection following sand fly transmission, the natural mode of infection (Fig. 1A). Exposure of WT and C3^−/− mice to the bites of infected sand flies revealed reduced lesion sizes in C3^−/− mice from the very outset of infection (p ≥ 0.02 at 3 wk p.i.) that was maintained over the course of the experiment (Fig. 1L, left panel) and associated with a significant, 10-fold reduction in the number of infected cells at 20 wk p.i. (Fig. 1L, right panel, p = 0.037). Despite the observed reduction in lesion size, a lack of C3 did not significantly reduce the likelihood that transmission would occur (Fig. 1M, top panel) or impact the likelihood that a lesion (of any size) would appear if infected (Fig. 1M, bottom panel), suggesting that parasites are able to establish an infection in the absence of C3, but that C3 plays a proparasitic role when present.

Of note, the relatively flat mean lesion curve in WT mice starting at 3 wk p.i. (Fig. 1L, green circles in Fig. 1N) is the result of the wide variation in disease progression in individual ears following exposure to infected sand fly bites (open circles in Fig. 1N). For example, within the 31 WT infected ears in two separate experiments, the earliest a lesion resolved was at 9 wk p.i. (red circles in Fig. 1N) whereas the latest a lesion appeared was at 10 wk p.i. (blue circles in Fig. 1N).

We next employed flow cytometric and microscopic analysis of in vitro L. major-RFP-neutrophil coculture to further define the impact of Leishmania infection on neutrophils (Fig. 2A, 2B). To validate our use of RFP expression by flow cytometry to assess infection, we directly compared same sample–purified bone marrow–derived neutrophils cocultured with serum-opsonized L. major-RFP employing both flow cytometric and direct microscopic analysis of cytospin preparations as performed previously (70). We found a parasite/neutrophil infection ratio–dependent linear correlation between the percentage of RFP⁺ neutrophils by flow cytometry and infected neutrophils by light microscopy (R² = 0.9477, Fig. 2C), as well as between RFP median fluorescence intensity (MFI) and the number of parasites per infected neutrophil (R² = 0.8455) (Fig. 2D). These observations confirm that RFP positivity and MFI as determined employing flow cytometry are the result of parasite internalization and parasite load per cell.

To investigate the role of serum opsonization and CD11b on uptake, we employed neutrophils isolated from WT and CD11b^−/− mice cocultured with serum-opsonized or nonopsonized parasites (Fig. 2E) and found that phagocytosis was highly dependent on CD11b (diamonds), regardless of serum opsonization, infection ratio, or duration of coculture (30 min versus 2 h). At early time points (open circles), opsonization was also required for optimal parasite uptake and this was most apparent at lower infection ratios, whereas at late time points (filled circles), neutrophils were able to phagocytose increasing numbers of parasites in the absence of opsonization. To determine the role of C3 in enhanced parasite uptake following opsonization, we employed mouse serum (MS) from WT (WT normal MS [NMS]) or C3^−/− mice (C3^−/− MS), or with HI serum from each strain, and analyzed uptake by 2P-IVM (Fig. 2F, Supplemental Videos 2, 3) or flow cytometry (Fig. 2G). Visualization of neutrophil parasite interactions revealed that optimal parasite phagocytosis occurred after opsonization with WT NMS but not with WT HI-NMS (p < 0.0001 at L. major; neutrophil ratios of ≥2 in Fig. 2G), as previously suggested (43), and was highly dependent on the parasite/neutrophil ratio. Opsonization of parasites with MS from C3^−/− mice gave the same result as HI-NMS (Fig. 2F, 2G), and pretreatment of neutrophils with LPS, a potent mediator of neutrophil activation (75), did not enhance phagocytosis (Fig. 2G, lower panel). The increases in the percentage of infected neutrophils following coculture with C3^−/−- or HI-MS–opsonized parasites at high L. major/neutrophil ratios suggest that non–C3-mediated mechanisms of parasite phagocytosis exist, as has been previously shown for macrophages (37), but that this pathway likely has limited importance to neutrophil uptake, as most parasites are taken up by neutrophils within a few minutes of contact (Fig. 2F, 2G) (3). These observations indicate that neutrophils almost exclusively employ the CD11b receptor for parasite phagocytosis and that this is enhanced by serum opsonization via C3.

Given our previous observation that NADPH-dependent ROS mediate the induction of neutrophil apoptosis as defined by annexin V binding and had a significant impact on reducing lesion size but did not impact parasite load, we next determined the impact of infection and mechanism of uptake on the detection of intracellular ROS employing the colorimetric DHR123 assay and flow cytometry (Fig. 3A). Neutrophils alone, infected neutrophils in the absence of DHR123, and neutrophils stimulated with PMA, a pharmacological activator of NADPH-dependent ROS production, were employed as controls. We have previously shown that our L. major-RFP parasite loses fluorescence as the parasite progresses toward parasite death (3). Employing RFP expression as a readout of viable neutrophil infection (Fig. 2C) we found that NADPH-dependent ROS had a statistically significant but negligible impact on Leishmania survival in neutrophils at 1 and 3 h following in vitro infection (Fig. 3B), as previously shown in vivo (50). To ensure that RFP⁺ neutrophils did indeed contain viable parasites, we cell sorted RFP⁺ neutrophils and titrated infected cells in parasite growth medium. We found that the frequency of RFP⁺ neutrophils from both WT and Phox^−/− samples containing at least one viable parasite was not significantly different from the theoretical value of 100% viable (Wilcoxon signed rank test; WT, p = 0.25; Phox^−/−, p > 0.9999) and were not different from each other (Fig. 3C). We also determined the RFP MFI and, similar to our percent RFP⁺ data, found a negligible but significant 1.6-fold (1 h) and 2.25-fold (3 h) decrease in RFP MFI in WT versus Phox^−/− RFP⁺ neutrophils (Fig. 3D). The minimal level of NADPH-dependent parasite elimination occurred despite robust infection-dependent neutrophil activation as indicated by a rapid increase in CD11b expression, a marker of neutrophil activation (51), on WT RFP⁺ versus RFP⁻ neutrophils at the 8:1 infection ratio or neutrophils cultured alone (Fig. 3E).

We next determined the kinetics of ROS production by neutrophils following coculture and found that both the percentage of ROS⁺ neutrophils (Fig. 3F, top panel) and amount of ROS per neutrophil (Fig. 3F, bottom panel) were greatest at 30 min p.i. Comparison of RFP⁻ versus RFP⁺ neutrophils from the same well revealed that intracellular infection was associated with a higher frequency of ROS⁺ neutrophils (Fig. 3G, top panel) and higher levels of ROS production per neutrophil (Fig. 3G, bottom panel), indicating that intracellular infection was required for optimal ROS production. The frequency of ROS⁺ uninfected RFP⁻ neutrophils also increased versus neutrophils alone (open circles in Fig. 3G, upper panel). This could be due to the small number of neutrophils that are able to kill the parasite, resulting in an RFP⁻ROS⁺ phenotype, NADPH activation by cell surface contact with the parasite, or diffusion of ROS from RFP⁺ to RFP⁻ cells. In any case, although these RFP⁻ cells are ROS⁺, the amount of ROS that is produced is very low as determined by ROS MFI versus RFP⁺ cells (Fig. 3A and Fig. 3G, 3I, lower panels).

To determine the degree to which ROS production was dependent on phagocytosis of live parasites, we exposed neutrophils to live or fixed parasites as well as PE latex beads. Although phagocytosis of serum-incubated beads and serum-opsonized fixed parasites was less efficient compared with live parasites (Fig. 3H), we were able to detect a sufficient number of PE/RFP⁺ neutrophils following 30 min of coculture. Phagocytosis of serum-exposed latex beads led to minimal ROS production, demonstrating that ROS production does not occur following internalization by default (Fig. 3I). In contrast, phagocytosis of fixed Leishmania resulted in similar frequencies and ROS production compared with live parasites. Therefore, respiratory burst was associated with phagocytosis of Leishmania but not potential downstream activation mediated by live infection.

We next determined the degree to which ROS production is dependent on phagocytosis mediated by C3/CD11b. As expected, given the almost absolute requirement for CD11b for parasite uptake by neutrophils (Fig. 2E) and the tight correlation between intracellular infection and ROS production (Fig. 3G), ROS production was highly dependent on CD11b when the total neutrophil population was analyzed (Fig. 3J). As expected, this was not the case when neutrophils were stimulated by PMA. Of interest, even when the small but detectable population of RFP⁺CD11b^−/− neutrophils was analyzed for ROS production versus WT neutrophils, a highly significant decrease in ROS production was observed (Fig. 3J, RFP⁺ group), demonstrating that respiratory burst is enhanced when Leishmania gains access to neutrophils via CD11b. We also analyzed ROS production following phagocytosis of parasites opsonized with C3^−/− serum over time (Fig. 3K). We once again gated on RFP⁺ infected cells and found that the early induction of ROS was greatly reduced in the absence of C3 (Fig. 3K). Interestingly, at the 120 min time point, when parasites begin to gain access to neutrophils in a CD11b-dependent but C3/opsonization-independent manner (Fig. 2J), this difference was lost (Fig. 3K). However, this late and low-level C3-independent ROS production following parasite internalization was not mediated by phagosomal NADPH, as ROS was detected in gp91 PHOX^−/− neutrophils (Fig. 3K), suggesting that it is derived from the mitochondria (76) and not the phagolysosome.

It has been suggested that Leishmania actively subverts respiratory burst or that internalization via CD11b is “silent.” However, employing increasing infection ratios, a comparison of RFP MFI and ROS MFI within RFP⁺ infected cells revealed that increasing multiplicities of infection resulted in greater frequencies of infection and amounts of ROS production per infected cell, providing further evidence that the parasite does not subvert respiratory burst (Fig. 3L). To investigate this further, we also determined the cellular localization of ROS within neutrophils employing direct microscopic evaluation of the ROS detection reagent NBT. In remarkable fashion, NBT deposition was exclusively associated with the parasite inside the phagolysosome and this was not observed employing gp91^−/−, NADPH/NOX2-deficient neutrophils (Fig. 3M). Note that the NBT reagent quenches the RFP signal in NBT⁺ infected WT neutrophils. These observations confirm that L. major induces robust production of NADPH/NOX2-dependent ROS that are deposited directly onto the parasite surface in the phagolysosome.

Having determined that Leishmania phagocytosis induced neutrophil activation and respiratory burst in a CD11b/C3-dependent manner, we next determined the impact of infection on neutrophil cell death employing annexin V staining. Annexin V binds to PS from the inner leaflet of the plasma membrane that becomes exposed on the outer cell membrane during apoptosis (Fig. 4A). Both the frequency and amount of annexin V staining were significantly increased on RFP⁺ infected cells versus RFP⁻ uninfected cells from the same well, gated as shown in Fig. 4A (lower panels), and this was even greater when compared with wells with neutrophils alone (Fig. 4B), similar to our previous observations employing later time points and the L. amazonensis parasite (50). Increased annexin V staining also occurred when fixed parasites were employed for infection (Fig. 4C). In contrast, uptake of latex beads resulted in reduced frequencies and binding levels of annexin V versus Leishmania uptake, although this was still greater than neutrophils alone. In agreement with our previous studies on the role of ROS in the acute neutrophil apoptotic phenotype (≤2 h p.i.) (50), this early expression of PS was not dependent on ROS, as PHOX^−/− neutrophils also had increased levels of PS expression/annexin V binding (Fig. 4D). The increase (p = 0.04) in annexin V staining on WT versus PHOX^−/− uninfected RFP⁻ neutrophils may be due to a requirement for NADPH/NOX2 in a bystander effect of coculture on uninfected cells. In contrast, early PS exposure occurred following phagocytosis of Leishmania and did not require live infection or NADPH/NOX2 (Fig. 4C, 4D). The reduced frequencies of annexin V⁺ cells following uptake of beads suggest that the size or complexity of the parasite is associated with the degree of membrane flipping induced by phagocytosis. To determine the impact of infection and subsequent increased PS expression on long-term neutrophil viability, we also counted the number of live neutrophils per well following culture of neutrophils alone versus neutrophils plus Leishmania (Fig. 4E). Surprisingly, we found that despite the early expression of PS on infected neutrophils, suggestive of apoptosis, neutrophil-Leishmania coculture wells contained higher, not lower, numbers of total viable neutrophils at 24 and 48 h p.i. Analysis of infected (RFP⁺) versus uninfected (RFP⁻) neutrophils or neutrophils alone at 48 h after initiation of coculture revealed that infected neutrophils were still viable at 48 versus 2 h, whereas uninfected neutrophils were significantly reduced, suggesting that infection increased neutrophil survival.

To investigate this further we measured mRNA transcripts of several apoptosis- and cell survival–associated genes in neutrophil cultures alone or neutrophils cultured with Leishmania to determine whether infection altered the expression of genes involved in apoptosis (Fig. 5A). We found that mRNA transcription of casp3, casp8, casp9, and bcl-2 were all reduced in neutrophils cultured with Leishmania, suggesting that Leishmania may disrupt induction of caspase 3–mediated apoptosis initiated by both the intrinsic (caspase 9–mediated) and extrinsic (caspase 8–mediated) pathways. We then set out to measure caspase protein abundance in neutrophils by Western blot to substantiate our observations. Whereas we were able to successfully detect caspase 8 and caspase 9 in macrophages (Supplemental Fig. 1), we were not able to reliably detect these proteins employing infected neutrophils. In contrast, we were able to reliably detect caspase 3, and we found that L. major infection was associated with decreased procaspase 3 (Fig. 5B). Note that the increased β-actin signal in neutrophil+L. major wells, which increased with time, is the result of the tendency of neutrophils to cluster during in vitro Leishmania coculture as observed in Fig. 2B, 2F and Supplemental Video 2, resulting in more efficient cell recovery during washing steps. Therefore, our results in Fig. 5B (right panel) are adjusted for β-actin. To determine whether decreased procaspase 3 abundance was due to increased cleavage and determine the relationship between cleaved caspase 3 protein abundance and annexin V surface staining, we performed flow cytometric and Western blot analysis of neutrophils from the same culture wells (Fig. 5C, 5D). Although neutrophil internalization of parasites was once again associated with increased annexin V staining (Fig. 5C), we found reduced, not increased, levels of cleaved caspase 3, as determined by the amount of cleaved caspase-3 divided by available procaspase 3 (Fig. 5D). This reduction was consistent over time (Fig. 5D, 30 min–3 h, and 5E, 30 min–10 h) and occurred in multiple experiments (Fig. 5E, right panel, n = 13 independent experiments). In addition, the reduction in cleaved Caspase 3 also occurred when employing fixed parasites (Fig. 5D), suggesting that the inhibitory mechanism employed by the parasite does not require live infection. These results indicate that while phagocytosis of the Leishmania parasite results in rapid (within 30 min) and high levels of PS exposure on the outer membrane, this is unlikely due to the execution of a classical caspase 3–mediated apoptotic program and is not associated with increased cell death. To determine whether neutrophil acquisition of Leishmania can actively inhibit apoptosis, we determined the impact of infection on the induction of apoptosis by TNF-α, a known inducer of the extrinsic pathway of apoptosis via the TNF receptor and a biologically relevant cytokine (77). Actinomycin D (Act D), a potent inhibitor of transcription and inducer of apoptosis, was included as a positive control (78). We found that TNF-α induced PS expression on neutrophils alone whereas Act D also induced membrane permeability as indicated by uptake of the cell membrane permeability dye 7-AAD (Fig. 5F). Interestingly, analysis of RFP⁺ infected neutrophils exposed to TNF-α revealed decreased annexin V staining versus RFP⁻ neutrophils from the same well, suggesting that L. major uptake does actively inhibit phagocytosis-independent, TNF-α–mediated induction of apoptosis (Fig. 5G). In contrast, Leishmania had a more intermediate impact on Act D–mediated induction of apoptosis, resulting in a higher frequency of annexin V⁺7-AAD⁺ apoptotic cells but slightly lower frequencies of necrotic annexin V⁻7-AAD⁺ cells, suggesting that the parasite cannot prevent, and may enhance, Act D drug-induced apoptosis (Fig. 5H).

Current models of Leishmania infection suggest that neutrophils, extravasating into the site of tissue injury initiated by the sand fly proboscis, are among the first host cells to become infected by Leishmania metacyclic promastigotes in vivo and represent an initial point of contact between Leishmania and host cellular immunity (3, 14, 15, 79). Understanding these early interactions may provide novel insights into the mechanisms by which Leishmania parasitize host immune processes to establish a productive infection and contribute to improving vaccine-mediated immunity, which must be in place during this critical early window of the infection process to confer protection (80).

In this study, we report that neutrophil internalization of Leishmania predominantly occurs via CD11b and is enhanced by opsonization with iC3b, the primary mechanism employed by macrophages (25, 26, 33, 43). L. major metacyclic promastigotes opsonized by employing C3-replete WT serum were rapidly (≤30 min) internalized by neutrophils whereas nonopsonized, HI serum-opsonized, or C3^−/− serum-opsonized L. major was internalized only after extended periods of coincubation (approximately equal to 2 h). Previous studies have demonstrated that LPG expressed on the Leishmania promastigote surface can bind directly to CD11b (CR3/Mac1) via an iC3b-independent mechanism (34, 81), albeit poorly, and this may also be the case for neutrophils at these later time points. In our hands, uptake of L. major by neutrophils was almost entirely CD11b-dependent both in vitro and in vivo, regardless of opsonization status. These results suggest that, unlike macrophages, which are known to possess alternative receptors for Leishmania internalization (27–32, 82), uptake of L. major by neutrophils is almost entirely CD11b-dependent and is enhanced by iC3b opsonization. It is likely that C3-CD11b–mediated uptake represents the biologically relevant mechanism of neutrophil internalization, as studies employing IVM and the in vitro observations reported in the present study demonstrated that parasites are rapidly internalized into neutrophils within minutes of initial parasite–neutrophil interactions (3). Similar to previous investigations on the role of C3 or CD11b on cutaneous leishmaniasis (40–42), we found that C3 was not required for infection, but it did play a significant proparasitic role following infected sand fly challenge, resulting in a 10-fold reduction in chronic parasite numbers and an ∼50% reduction in lesion sizes.

We next turned our attention to addressing the hypothesis that CD11b/iC3b-mediated uptake of Leishmania spp. metacyclic promastigotes may be unlinked from the production of NOX2-dependent ROS in the phagolysosomes of infected neutrophils (34–36). In contrast, our investigation clearly demonstrated that uptake of L. major metacyclic promastigotes by neutrophils resulted in a robust, parasite-dependent accumulation of intracellular ROS within 30 min of internalization. Employing NBT, we were also able to show that L. major uptake-associated ROS accumulation within infected neutrophils is due to a NADPH/NOX2-dependent intraphagosomal oxidative burst colocalized with the parasite surface.

Despite robust detection of ROS on the parasite surface, our data demonstrated a limited role for NOX2 ROS-mediated killing of L. major metacyclic promastigotes. The degree to which Leishmania parasites are susceptible to ROS has been reported to vary considerably by strain, but we would argue that this may be more related to variations in the efficiency of metacyclogenesis when nonpurified stationary-phase cultures are employed to infect phagocytes. Metacyclic promastigotes express the mature forms of the LPG molecule on the parasite surface and are believed to be more resistant to both ROS- and complement-mediated elimination. Why some parasites are eliminated may be because they are not fully differentiated or may be related to the “sacrificial parasite” hypothesis whereby some parasites undergo apoptosis to “trick” neutrophils into noninflammatory efferocytosis-like uptake. Therefore, it is important to consider that stationary phase, nonpurified parasites may be less representative of the metacyclic promastigote form of the parasites associated with mature transmissible infections by the sand fly vector (2, 67).

Although a role for NADPH-ROS on the modulation of Leishmania-infected neutrophil cell death has been established by Carneiro et al. (50), no published study has, to date, provided conclusive evidence for the mechanisms by which Leishmania infection modulates neutrophil cell death cascades or the precise role that oxidative burst plays in this process. Similar to previously published observations (10, 50), our investigation revealed that neutrophils infected with L. major stained positive for annexin V. While our previous work demonstrated a role for ROS in apoptosis at late time points of neutrophil infection (10 h), our earlier analysis reported in the present study (2 h) matched our previous finding that the initial expression of PS is NOX2 ROS-independent. Employing PE latex beads and live or fixed parasites, we were able to demonstrate that both ROS production and PS expression appear to be properties of neutrophils that have phagocytosed a live or dead parasite, but not beads, suggesting that these phenotypes do not require active infection and are instead related to parasite internalization. Analysis of the small number of CD11b^−/− neutrophils that acquired parasite also revealed a marked deficiency in ROS, again suggesting that CD11b is an important receptor in parasite internalization and activation of respiratory burst.

Previous studies have demonstrated that CD11b-dependent uptake of serum-opsonized Escherichia coli and/or yeast particles induced neutrophils to initiate a ROS-dependent program of phagocytosis-induced cell death through a mechanism that involved the activity of caspase 8 and caspase 3, but did not involve caspase 9 (83). Despite undergoing respiratory burst and PS externalization, neutrophils from Leishmania cocultures had enhanced, not reduced, viability. We found robust PS expression following internalization of live or fixed parasites but a dramatic reduction in both the pro and cleaved form of the executioner Caspase, Caspase 3, as well as evidence for a reduction in caspase-3 cleavage from available procaspase 3. Analysis of Caspase mRNA gene expression also revealed marked reductions in caspase 8, 9, and 3 expression, indicating that L. major infection reduces the capacity for these neutrophils to execute the terminal effector function of the cell death initiation pathway. Although data demonstrating changes in neutrophil caspase 3 activity following Leishmania infection have been published (11), so far as we are aware, our data are the first to directly show an effect on caspase 3 protein regulation in neutrophils. Our analysis further demonstrated that coincubation of neutrophils with L. major reduced the detectable levels of procaspase 3 via a mechanism that was not dependent on cleavage to effector caspase 3 and that was reflected in mRNA levels. Additionally, L. major cocultured neutrophils also displayed downregulated expression of Bcl-2 but no change in mcl-1. Antiapoptotic proteins of the BCL-2 family (which include both BCL-2 and MCL-1) exist in tension with proapoptotic proteins of the BCL-2 family (such as BAX and BAK) (84). In the absence of direct inhibition by antiapoptotic BCL-2 family members, the proapoptotic BCL-2 family members create pores in the mitochondrial outer membrane to facilitate the release of cytochrome c (84) and subsequent initiation the cell-intrinsic pathway of apoptosis. Downregulated Bcl-2 by L. major cocultured neutrophils is suggestive that L. major infection may induce a generalized reduction in many molecules involved in regulating cell death, regardless of their proapoptotic or antiapoptotic roles (12).

PS externalization following Leishmania infection has been employed extensively as a marker of apoptosis (10, 11, 50), leading to the conclusion that infected neutrophils quickly become apoptotic. However, our data suggest that infection-induced PS externalization is unlinked from the cleavage of executioner caspase 3, and therefore may be unlinked from executioner caspase 3–mediated apoptosis. These data help to reconcile disparate reports that have been published by different groups, using different experimental models, which have concluded that Leishmania both enhances or stalls apoptosis. Our observations suggest that Leishmania uptake mimics apoptosis by inducing membrane flipping and PS exposure while simultaneously preventing caspase 3–mediated PCD. To a certain extent, this abrogation of apoptosis appears to be an “active state,” as infection reduced phagocytosis-independent induction of annexin V expression by TNF-α.

We have shown that C3-coated Leishmania major is rapidly internalized by murine neutrophils through a mechanism that is dependent on engagement of the neutrophil surface receptor, CD11b, and is enhanced by an interaction between CD11b and the iC3b opsonin. We have also shown that uptake of L. major induces ROS accumulation on the parasite surface in neutrophil phagolysosomes and externalizes PS, but it results in only limited parasite killing. Furthermore, we have demonstrated that infected neutrophils displayed elongated viability associated with decreased cleavage of the executioner caspase, caspase 3, leading us to conclude that infected neutrophils are modulated into a phenotypic state of “stalled apoptosis.” When viewed in the context of previous publications (10–12, 83, 85), we believe that these findings provide further evidence to support the Trojan horse neutrophil model of Leishmania infection.

The authors have no financial conflicts of interest.

We thank Dr. Robin Yates for B6.129S6-Cybb^tm1Din mice, H. Kuipers for use of the Cytek Aurora, which was funded through the Canadian Foundation for Innovation, K. Poon for technical assistance and training on the BD FACSCanto, and A. Lau and H. Chung in the Daniel Muruve laboratory for assistance with Western blotting.