Mice

BALB/c, BALB/c RAG2^-/- and congenic CB17 SCID (CB17/lcr-Prkdc^SCID/lcrlcoCrl) mice that lack T and B cells due to a genetic autosomal recessive mutation in the Prkdc^SCID allele on chromosome 16 [22, 23] were bred and maintained in the animal facilities of the Bernhard Nocht Institute for Tropical Medicine, Hamburg, and housed in a biosafety level 3 facility for experimentation. The facilities are registered by the Public Health Authorities (Behoerde für Gesundheit und Verbraucherschutz, Hamburg). All experimentations and procedures were approved by the Public Health Authorities (Behoerde für Gesundheit und Verbraucherschutz, Hamburg: no 88/13) and performed according to the German Animal Welfare Act.

Culture, purification and freezing of R. typhi

R. typhi (Wilmington strain) was cultivated in L929 mouse fibroblasts (ATCC CCL-1) in RPMI1640 (PAA, Cölbe, Germany) supplemented with 10% FCS (PAA, Cölbe, Germany), 2 mM L-glutamine (PAA, Cölbe, Germany) and 10 mM HEPES (PAA, Cölbe, Germany) without antibiotics under biosafety level 3 conditions. 1×10⁷ L929 cells were seeded in 175 cm² culture flasks (Greiner Bio-One, Frickenhausen, Germany) and γ-irradiated (1966 rad at 560sec). One day later cells were infected with R. typhi and incubated for 5 to 7 days. Stocks of purified bacteria were prepared from L929 cell lysates. Therefore, cells were resuspended in 1.5 ml PBS and vortexed thoroughly for 1 min with 200 μl sterile siliceous particles (60/90 grit silicon carbite, Lortone inc., Mukilteo USA) in a 2 ml SafeSeal tube (Sarstedt, Nümbrecht, Germany). The crude lysate was strained through a 2 μm cell strainer (Puradisc 25 syringe filter 2 μm; GE Healthcare Life Sciences, Freiburg, Germany), mixed in a ratio of 1:1 with 2-fold concentrated storage medium (FCS/15% DMSO) and transferred into Cryo.S tubes (Greiner Bio-One, Frickenhausen, Germany) in liquid nitrogen. Thawed bacterial stocks were centrifuged at 7826xg for 5 min at room temperature, washed once with PBS and analyzed for bacterial content by quantitative real-time PCR (qPCR). Spot forming units (sfu) as a measure for the amount of living bacteria in the preparation were determined by immunofocus assay. For this purpose, L929 cells were incubated with titrated amounts of R. typhi in 24well plates. After 4h of bacterial adherence the medium was exchange against semi-solid medium containing 1% methylcellulose and cells were further incubated for 8–10 days. For detection of R. typhi, cells were fixed in PBS/4% formaldehyde/0.1% TritonX100 (Sigma-Aldrich, Deisenhofen, Germany) for 20 minutes followed by permeabilization in PBS/0.5% TritonX100 for 20–60 minutes. Cells were blocked with 200 μl PBS/10% FCS for 1 hour. Monoclonal anti-R. typhi antibody (BNI52) was added at 1 μg/ml in PBS/10% FCS overnight at 4°C. Cells were washed in H₂O and with goat anti-mouse HRP (Dako, Hamburg, Germany; 1:400 in PBS/10% FCS) for 1-2h in the dark at RT. Finally, cells were washed and plates were developed with Immunoblot 3,3′,5,5′-tetramethylbenzidine (TMB) substrate solution (Mikrogen, Neuried, Germany; 200 μl) and analyzed with a BZ9000 Keyence microscope (Keyence, Neu-Isenburg, Germany).

Infection of mice, neutrophil depletion and brefeldin A treatment

Mice were subcutaneously (s.c.) infected into the base of the tail with 2×10⁶ sfu R. typhi in 50 μl PBS. For neutrophil depletion, 200 μg anti-Ly6G were injected every 3 days intraperitoneally into CB17 SCID mice starting 6 days after infection with R. typhi. A control group of R. typhi-infected CB17 SCID received equal amounts of IgG2a isotype antibody. A second control group of animals was not infected and received anti-Ly-6G only. A third group of control mice was not infected and treated with PBS. Depletion of neutrophils was monitored by flow cytometry of blood cells. For flow cytometric detection of intracellular IFNγ and TNFα expression by different cell populations directly ex vivo 100 μg brefeldin A (#B7651; Sigma, Deisenhofen, Germany) were intravenously injected into R. typhi-infected CB17 SCID mice 12h prior to spleen cell isolation. Non-infected control animals that had received PBS instead of R. typhi were treated the same way. These analyses were performed on day 12 post infection or PBS treatment, respectively.

Clinical scoring and survival

Based on the findings described in the first part of the results section a clinical score was defined to monitor the health status of R. typhi-infected animals. The following five criteria were assessed: posture (0: normal, 1: temporarily curved, 2: curved), fur condition (0: normal, 1: staring in the neck, 2: overall staring), activity (0: normal, 1: reduced, 2: strongly reduced), weight loss (0: < 10%, 1: 10–14%, 2: > 15%) and food and water uptake (0: normal, 1: reduced, 2: none). Mice were considered healthy with a score < 5, moderately ill with a score of 5–7 and severely ill with a score of 8–10. Mice were euthanized reaching a total score of ≥8 or showing weight loss of ≥20%. This was determined as the time of death. The state of health of the animals was assessed by clinical scoring every 2 days.

Collection of blood samples

Blood was taken submandibular or by cardiac puncture after euthanasia with CO₂. For plasma samples blood was collected in EDTA coated tubes (KABE Labortechnik GmbH, Nümbrecht-Elsenroth, Germany) and centrifuged at 5654×g. Serum samples were obtained by agglutination for 15–20 at RT followed by centrifugation for 10 min 5654×g.

DNA preparation from purified bacteria, cell culture and organs

10 mg tissue was homogenized in 500 μl PBS in Precellys ceramic Kit tubes (Peqlab. Erlangen, Germany) in a Precellys 24 homogenizer (Peqlab. Erlangen, Germany) with following cycle parameters: 6000 rpm two times for 45 sec with a 60 sec break. DNA was prepared from 80 μl homogenized organs or up to 1×10⁶ L929 cells using the QIAamp DNA Mini Kit (Qiagen, Hilden, Germany) according to the manufacturer’s guide.

Quantitative real-time PCR (qPCR)

R. typhi was quantified by amplification of a 137 bp fragment of the PrsA gene (RT0565) with the forward primer 5´-ACA GCT TCA AAT GGT GGG GT-3´ and reverse primer 5´-TGC CAG CCG AAA TCT GTT TTG-3´ in a standard SYBR green real-time PCR. To determine R. typhi copy numbers a standard template plasmid (pCR2.1-PrsA) containing the same PrsA DNA fragment was used. Reactions were performed in a Rotor Gene 6000 (Qiagen, Hilden, Germany) in a total volume of 10 μl with 1×HotStar Taq DNA Polymerase Buffer comprising 1.5 mM MgCl₂, 0.175 mM dNTPs, 100 nM primers, 0.05x SYBR green I nucleic acid gel stain (SIGMA Life Science, Deisenhofen, Germany) and 0.25 U HotStar Taq DNA Polymerase (Qiagen, Hilden, Germany) under following conditions: 15 min pre-heating at 95°C followed by 40 cycles of denaturation (94°C for 20 sec), primer annealing (53°C, 30 sec) and elongation (72°C for 20 sec).

Antibodies and reagents

Anti-R. typhi (BNI52) is a monoclonal mouse antibody that was generated at the Bernhard Nocht Institute for Tropical Medicine, Hamburg, Germany. It was used at 1 μg/ml for immunofluorescent stainings of infected cell cultures and flow cytometry. For flow cytometry the following antibodies were used at the indicated dilutions and concentrations: anti-NOS2 (iNOS)-PE (clone CXNFT, 1:200), anti-NOS2 (iNOS)-PE/Cy7 (clone CXNFT, 1:300), anti-MHCI (H-2d)-PE (clone SF1-1.1.1, 1:200), rat IgG2a κ Isotype PerCp-Cy5.5 (eBR2a) and anti-mouse NKp46-PE (clone 29A1.4, 1:200) from eBioscience, Frankfurt, Germany; anti-CD11b-PerCp-Cy5.5 (clone M1/70, 1:200; 1:800 for bmMΦ) from BD Bioscience, Heidelberg, Germany; anti-Ly6-C-PerCP/Cy5.5 (clone HK1.4, 1:200), anti-Ly-6G-APC (clone 1A8, 1:166,7), anti-GR1 (L6G/Ly6C)-APC (clone GR-1/RB6-8C5, 1:500), anti-CD80-PE/Dazzle594 (clone 16-10A1, 1:100), anti-CD11b-FITC (clone M1/70, 1:100), rat IgG2a κ isotype PE (RTK2758; 1:200), rat IgG1 κ isotype PE-Cy7 (RTK20711; 1:200), anti-mouse IFNγ PE/Dazzle (clone XMG1.2; 1:333), anti-mouse TNFα BV510 (clone MP-6-XT22; 1:80) and rat IgG1 κ isotype PE/Dazzle 594 (RTK 2071; 1:200) from BioLegend (London, UK); unlabeled mouse IgG3 κ isotype (clone B10; 1 μg/ml) and anti-mouse-IgG3-FITC from SouthernBiotech (#1100–02; Birmingham, USA; 1:200). Histological stainings were performed with the following antibodies and reagents: R. typhi patient serum (1:100) from the diagnostics department of the Bernhard Nocht Institute for Tropical Medicine, Hamburg, Germany; anti-mouse iNOS (ABIN373696, 1:75) from Abcam, Cambridge, USA; anti-mouse IBA1 (#019–19741; 1:500) from WAKO, Neuss, Germany; anti-mouse Ly-6G (clone 1A8; 1:1000) from BD Biosciences, Heidelberg, Germany; anti-mouse-IgG3-FITC (#1100–02; 1:200) from SouthernBiotech, Birmingham, USA; anti-human IgG-FITC (#H10101C; 1:200), anti-rabbit Alexa555 (#A31572; 1:300) and anti-rat Alexa568 (#A11077; 1:300) from Thermo Fisher Scientific, Braunschweig, Germany; anti-FITC-Alexa488 (1:1000) from LifeTechnologies, Darmstadt, Germany; DAPI (4´,6-diamidino-2-phenylindole dihydrochloride; 1:1000) and CohnII human IgG fraction (5% in PBS) from Sigma, Deisenhofen, Germany. Isotype antibodies were used at concentrations corresponding to the respective staining antibodies. For neutrophil depletion anti-Ly-6G (clone 1A8) and rat IgG2a (clone 2A3) isotype control antibody (BioXCell, New Hampshire, USA) were used.

Flow cytometry

Single cell suspensions were prepared from spleen, liver, blood samples or cell culture. Erythrocytes were eliminated from spleen, liver and blood samples by incubating the cells in erythrocyte lysis buffer (10 mM Tris, 144 mM NH₄Cl, pH 7.5) for 5 minutes at room temperature. Afterwards, cells were washed twice with PBS. Fc receptors were blocked with 50 μl 5% CohnII human IgG fraction (Sigma-Aldrich, Deisenhofen, Germany) in PBS or Perm/Wash solution. BD Cytofix/Cytoperm and BD Perm/Wash solutions (BD Biosciences, Heidelberg, Germany) were used for intracellular staining of iNOS and R. typhi. Intracellular cytokines were detected employing the FoxP3/Transcription Factor Staining Buffer Set (eBioscience, Frankfurt, Germany). Procedures were performed according to the manufacturer´s instructions. CD80 and MHCI on bmMΦ were first stained extracellularly followed by intracellular staining of iNOS and R. typhi. Antibodies were diluted in 50 μl of either PBS or Perm/Wash solution. Cytokines (TNFα and IFNγ), NKp46, CD11b and GR1 were stained simultaneously intracellularly in Permeabilization Buffer of the FoxP3/Transcription Factor Staining Buffer Set. After staining, cells were washed and resuspended in PBS/1% PFA or PBS/10% FCS prior to flow cytometry. Analyses were performed with a BD Accuri C6 or BD LSR II flow cytometer (BD Biosciences, San José, USA) and FlowJo single cell analysis software (FlowJo LLC, Ashland, USA).

Detection of reactive oxygen species (ROS)

ROS release was determined in blood cells of infected mice by the formation of the fluorescent compound rhodamine-123 from dihydrorhodamine-123 (DHR-123, AAT Bioquest, California, USA). Staining was performed with 30 μl EDTA blood samples after erythrocyte lysis. Surface markers were stained as described above prior to ROS detection. Cells were incubated with 30 μg/ml DHR-123 in PBS for 20 min at 37°C in the dark. After washing with 4 ml of cold PBS 40 μl of cell suspensions were immediately analyzed using BD Accuri C6 (BD Biosciences, San José, USA) and FlowJo single cell analysis software (FlowJo LLC, Ashland, USA).

Detection of cytokines in plasma and cell culture supernatants

Plasma cytokines were quantified by bead-based LEGENDplex immunoassay (BioLegend, London, UK) according to the manufacturer’s protocol using cluster tubes (ThermoScientific, Loughborough UK). 12.5 μl of plasma from EDTA blood samples was used diluted 1:2 in PBS. Supernatants from bmMΦ were used non-diluted. Analyses were performed using a BD Accuri C6 (BD Biosciences, San José, USA) and LEGENDplex analysis software (BioLegend, San Diego, USA).

Detection of serum Glutamate pyruvate transaminase (GPT)

Serum levels of GPT were evaluated using Reflotron GPT (ALT) stripes and Reflotron Plus device (Roche Diagnostics, Mannheim, Germany) according to the manufacturer’s instructions. Serum samples were diluted 1:3 in PBS prior to analyses.

Generation and infection of bmMΦ in vitro

Bone marrow was isolated from femur and tibia of BALB/c mice. 2×10⁶ cells were differentiated in petri dishes for 12 days in IMDM (PAA, Cölbe, Germany) supplemented with 10% FCS, 2 mM L-glutamine, 5% horse serum (Biochrom, Berlin, Germany) and L929 fibroblast medium as a source of M-CSF. Fresh medium was applied every 3 days. After 12 days of differentiation bmMΦ were harvested and washed twice with PBS. For analysis of NO and cytokine release as well as flow cytometry, bmMΦ were seeded into 24-well tissue culture plates at 5×10⁵ cells per well and infected in duplicates with 5, 10 or 25 R. typhi particles as determined by qPCR per cell. Control cells were incubated with medium or stimulated with 0.5 μg/ml E. coli (strain 055:B5) lipopolysaccharide (LPS) (Sigma, Deisenhofen, Germany). After incubation of 24 and 48 hours cells were analyzed by qPCR and flow cytometric staining of R. typhi, MHCI, CD80 and iNOS. Cytokines and nitric oxide (NO) were quantified in the supernatants as described below.

Detection of NO

NO concentrations were determined by Griess reaction in supernatants of bmMΦ. Assays were performed in microtiter plates (Greiner Bio-One, Frickenhausen, Germany). 100 μl of sample were mixed with 50 μl Griess 1 reagent (0.5 g sulfonamide in 50 ml 1M HCl) and 50 μl Griess 2 reagent (0.15 g naphtylethylendiamine-dihydrochloride in 50 ml H₂O). A serial dilution of sodium nitrite (NaNO₂) in culture medium was used as a standard (c_max 125 μM). The absorbance was measured at 560 nm with a Dynex MRXII spectrophotometric microplate reader (Dynex Technologies, Chantilly, USA).

Immunofluorescence staining of R. typhi in bmMΦ

5×10⁴ bmMΦ were seeded into 8well Nunc Permanox chamber slides (Sigma-Aldrich, Munich, Germany). Living R. typhi or heat-inactivated R. typhi particles (30 min, 56°C) were added at 10 copies per cell. Control cells were not infected. Medium was exchanged after 4 h of bacterial adherence. Cells were further incubated for 48 h and permeabilized by addition of ice-cold acetone:methanol (1:3) for 10 minutes at -20°C. Cells were washed 3 times in PBS and staining procedures were then performed at 37°C. Fc receptors were blocked with 5% CohnII in PBS for 15 minutes and 1 μg/ml anti-R. typhi (BNI52) was added for additional 30 minutes. After washing, cells were stained with anti-mouse IgG3-FITC (1:200) in PBS for 30 minutes followed by staining with anti-FITC-Alexa488 and DAPI (both 1:1000). Finally, cells were washed in PBS and slides were covered with Permafluor Mounting Medium and cover slips (ThermoScientific, Loughborough UK). Images were taken with the Axioskop MC-80 microscope (Zeiss, Oberkochen, Germany).

Histological stainings

For immunohistochemistry (IHC) tissues from infected mice were fixed in 4% formalin in PBS and embedded in paraffin. Deparaffinization of the sections was performed using standard methods. Sections were first heated at 63°C for 30 minutes in a heating cabinet followed by treatment with Xylol for 30 minutes and EtOH (3x 100% EtOH, 3x 96% EtOH, 80% EtOH, 70% EtOH). Each step was performed for 3–5 minutes. Slides were finally washed in H₂O. Deparaffinized sections were boiled for 30 minutes in 10 mM citrate buffer (10 mM sodium citrate, 0.05% Tween20, pH6.0) for antigen retrieval. Staining was performed using a Ventana Benchmark XT apparatus (Ventana, Tuscon, USA). Antibodies were diluted in 5% goat serum (Dianova, Hamburg, Germany) in Tris-buffered saline pH7.6 (TBS) and 0.1% Triton X100 in antibody diluent solution (Zytomed, Berlin, Germany). Rabbit anti-mouse IBA1 (1:500), rabbit anti-mouse iNOS (1:75) and rat anti-mouse Ly-6G (1:1000) were used. R. typhi was detected employing serum from a R. typhi-patient (1:100). Slides were incubated with primary antibodies for 1 h. Histofine Simple Stain MAX anti-human, anti-rabbit, anti-mouse or anti-rat peroxidase-coupled antibodies (Nichirei Biosciences, Tokyo, Japan) were used as secondary antibodies. Detection was performed with ultraview universal DAB detection kit (Ventana, Tuscon, USA). For immunofluorescent stainings donkey anti-rabbit Alexa555 (1:300), goat anti-rat Alexa568 (1:300), goat anti-human IgG-FITC (1:200) and anti-Alexa488-FITC (1:1000) were used as secondary antibodies. Nuclei were stained with DAPI (1:1000). Sections were covered with Tissue-Tek embedding medium (Sakura Finetek, Staufen, Germany). Images were taken with a BZ9000 Keyence microscope (Keyence, Neu-Isenburg, Germany).

Statistical analyses

Statistical analyses were performed with GraphPad Prism 5 software (GraphPad Software, Inc., La Jolla, USA). The proportion of surviving animals was analyzed with the Log-rank Mantel Cox test. Normality test was performed with D'Agostino-Pearson normality test for n≥8. For comparison between two groups two-tailed Students t-test for parametric samples or Mann-Whitney U test for non-parametric samples were applied. To assess differences between multiple groups Kruskal-Wallis test followed by Dunn´s post-test or Two way ANOVA followed by Tukey´s post- est were applied.

T and B cell-deficient CB17 SCID mice are highly susceptible to R. typhi and show disseminated infection

To determine the susceptibility of CB17 SCID mice to R. typhi infection, we infected the animals with titrated amounts of R. typhi (2×10⁶, 2×10⁴ and 2×10² sfu) s.c. into the base of the tail. A control group of CB17 SCID mice received PBS instead of R. typhi. Immunocompetent congenic BALB/c wild-type mice were infected with the highest dose of R. typhi as an additional control. As expected, BALB/c wild-type mice did not show clinical symptoms of disease at any point in time and all animals survived the infection (Fig 1A and S3B Fig). In contrast, CB17 SCID mice were highly susceptible. 100% of the mice that received 2×10⁶ sfu and 94% of the mice that were infected with 2×10⁴ sfu succumbed to the infection while 60% of the mice that obtained 2×10² sfu survived the infection (Fig 1A). The survival period was 16.93±3.75 days for mice that were infected with the highest dose, 22.07±2.74 days for mice infected with the median dose and 24.5±3.54 days for those that received the lowest dose. Staring fur was the first sign of illness and appeared around day 7–10 post infection in animals that received the highest dose of R. typhi (2×10⁶ sfu) while the onset of disease was later in mice that were infected with 2×10⁴ (day 12–14). Disease then rapidly progressed and the animals showed hunchback appearance and inactivity. This correlated with body weight loss that further progressed until death while PBS-treated CB17 SCID control mice remained healthy and gained weight (Fig 1A). Based on these findings the lethal dose of 2×10⁶ sfu was used for all further experimentation.

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Fig 1. CB17 SCID mice succumb to R. typhi infection and develop systemic infection.

CB17 SCID mice were infected s.c. with 2×10⁶, 2×10⁴ or 2×10² sfu R. typhi into the base of the tail (n = 10 for each group). BALB/c mice received 2×10⁶ sfu R. typhi via the same route (n = 11). Control CB17 SCID mice received PBS (n = 10). Survival rates (left; y-axis) and weight loss were monitored (right; y-axis) (A). Bacterial burden (y-axis) was determined in liver, spleen, brain and lung of CB17 SCID after s.c. infection with 2×10⁶ sfu R. typhi (n = 10–11) by PrsA qPCR at the time of death (B). Data represent combined results from two independent experiments and show the mean ± SEM.

https://doi.org/10.1371/journal.pntd.0004935.g001

Next, bacterial dissemination was analyzed. Bacterial load was determined in liver, spleen, brain and lung by qPCR. High amounts of R. typhi were found in all organs in CB17 SCID mice at the time of death. The highest bacterial load was detected in spleen followed by the brain, lung and liver (Fig 1B), demonstrating disseminated infection in these animals. These results show that CB17 SCID mice are highly susceptible to R. typhi infection and fail to control bacterial growth.

R. typhi-infected CB17 SCID mice develop splenomegaly which is predominantly caused by the accumulation of neutrophils and MΦ

We further investigated pathological changes in CB17 SCID mice. R. typhi-infected CB17 SCID mice showed a very strong increase in spleen size and weight (288.2±23.3 mg) at the time of death compared to 33.1±2.6 mg in control CB17 SCID mice (Fig 2A). The cellular composition of the spleen of R. typhi-infected CB17 SCID mice was further assessed by flow cytometric analysis. Cells were stained for CD11b and GR1 to distinguish CD11b⁺GR1^hi neutrophils from CD11b⁺GR1^low MΦ/monocytes as depicted in Fig 2B (left). R. typhi-infected CB17 SCID mice showed a significantly increased frequency of MΦ/monocytes. At the time of death CD11b⁺GR1^low MΦ/monocytes constituted 27.6±4.3% of the spleen cells compared to 12.1±1.1% in CB17 SCID control mice (Fig 2B, middle). A similar trend was true for CD11b⁺GR1^hi neutrophils that represented 9.5±1.9% of the cells in the spleen of control mice and 15±2.5% of the spleen cells in R. typhi-infected CB17 SCID mice (Fig 2B, right). Thus, together MΦ/monocytes and neutrophils represented approximately 40% of the spleen cells.

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Fig 2. CB17 SCID mice develop splenomegaly which is largely due to the accumulation of MΦ and neutrophils.

CB17 SCID mice were infected s.c. with 2×10⁶ sfu into the base of the tail. Control CB17 SCID mice received PBS. The photograph shows a representative spleen of a control mouse (left) and two R. typhi-infected CB17 SCID mice at the time of death (middle and right). Spleen weight (y-axis) was determined. The graph shows combined results from two independent experiments. Each dot represents a single mouse (n = 10–11). The mean ± SEM is presented. Data were analyzed with unpaired t test. Asterisks indicate statistically significant differences (***p<0.001) (A). The percentage of CD11b⁺GR1^low MΦ/monocytes and CD11b⁺GR1^hi neutrophils among spleen cells (y-axis) in CB17 SCID control animals (open circles) and R. typhi-infected CB17 SCID mice (black circles) was assessed at the time of death of the animals by flow cytometry. A representative staining for CD11b and GR1 and gating of cells is shown for a control mouse and a R. typhi-infected CB17 SCID mouse. Numbers indicate the percentage of the CD11b⁺GR1^hi neutrophils and CD11b⁺GR1^low MΦ/monocytes. Graphs show combined results from two independent experiments. Each dot in the graphs represents a single mouse (n = 10–11). The mean ± SEM is depicted. Statistical analysis was performed with student`s t-test after D´Agostino and Pearson normality test. Asterisks indicate statistically significant differences (** p<0.01) (B). BALB/c (n = 5–7) and CB17 SCID mice (n = 5–7) were infected s.c. with 2×10⁶ sfu into the base of the tail. Total numbers of CD11b⁺GR1^low MΦ/monocytes and CD11b⁺GR1^hi neutrophils among spleen cells (y-axis) were assessed by flow cytometric staining as described above during the course of infection. Graphs show the total numbers of CD11b⁺GR1^low MΦ/monocytes and CD11b⁺GR1^hi neutrophils (y-axis) at the indicated point in time (x-axis). The mean ± SEM is depicted. Statistical analysis was performed with Kruskal-Wallis test followed by Dunn´s post-test comparing samples from infected mice with samples from control animals (day 0) (C).

https://doi.org/10.1371/journal.pntd.0004935.g002

We further analyzed the absolute numbers of MΦ/monocytes and neutrophils in the spleen during the course of infection and compared CB17 SCID mice and BALB/c wild-type mice. In BALB/c wild-type mice numbers of CD11b⁺GR1^low MΦ/monocytes remained unaltered during the course of infection (Fig 2C, left) while numbers of CD11b⁺GR1^hi neutrophils were slightly enhanced early in infection on day 3 (control: 7.22×10⁵, R. typhi-infected: 1.73×10⁶) and returned to basal counts until day 7 (Fig 2C, right). In R. typhi-infected CB17 SCID mice numbers of CD11b⁺GR1^hi neutrophils and CD11b⁺GR1^low MΦ/monocytes steadily increased during the course of infection beginning on day 3 and resulting in an approximately 30-fold increase of MΦ/monocytes (control: 2.5×10⁵, R. typhi-infected: 8.2×10⁶; Fig 2C, left) and 50-fold increase of neutrophils immediately prior to death (control: 1.18×10⁵, R. typhi-infected: 6.74×10⁶; Fig 2C, right).

These results demonstrate that splenomegaly in R. typhi-infected CB17 SCID mice is largely due to disproportionate increase of MΦ/monocytes and neutrophils.

CB17 SCID mice develop severe liver necrosis upon R. typhi infection

Next to the massive enlargement of the spleen R. typhi-infected CB17 SCID mice developed severe liver necrosis which was already visible by eye. In addition, the gall bladder of R. typhi-infected CB17 SCID mice was dark, indicating endothelial damage and bleedings in the organ (Fig 3A, left). Liver damage was measurable by significantly elevated levels of GPT in the serum of R. typhi-infected CB17 SCID mice (Fig 3A, middle). In addition, liver weight was enhanced in several animals at the time of death although these differences were not significant (1162±166.8 mg compared to 1035±11.7 mg in control mice; Fig 3A, right). Numerous necrotic areas were visible in histological stainings of the liver of R. typhi-infected CB17 SCID mice compared to control animals that received PBS instead of bacteria and several spots of infiltrating cells were detectable that were not present in the liver of healthy control mice (Fig 3B). Cellular infiltration began around day 7 post infection when the bacteria were predominantly found in endothelial cells and necrotic lesions were still absent. Infiltrating cells further increased until death around day 15. At this point in time the bacteria were detectable in foci of cellular infiltrates (S1 Fig). Further flow cytometric analysis of cellular isolates from the liver at the time of death confirmed a significant increase of cellular infiltrates (Fig 4A, left). Among these a significantly enhanced percentage of both CD11b⁺GR1^low MΦ/monocytes (12.6±2.5% compared to 5.9±1.2% in control mice; Fig 4A, middle) and CD11b⁺GR1^hi neutrophils (16.9±2.4% compared to 3.7±0.7% in control mice; Fig 4A, right) was observed, demonstrating that MΦ/monocytes as well as neutrophils infiltrate the liver to a comparable extent.

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Fig 3. R. typhi-infected CB17 SCID mice develop severe liver necrosis.

CB17 SCID mice were infected s.c. with 2×10⁶ sfu into the tail base while control mice received PBS. The photograph shows a representative liver of a R. typhi-infected CB17 SCID mouse at the time of death (left). GPT was measured in sera (y-axis) of control mice (open symbols) and R. typhi-infected CB17 SCID mice (black symbols) (middle) and liver weight (y-axis) was determined at the time of death (right). The graphs show combined results from two independent experiments (n = 10). Each dot represents a single mouse. The mean ± SEM is given. Statistical significance was analyzed by Mann Whitney U test after D´Agostino and Pearson normality test (A). Representative histological stainings of the liver from a control mouse (left) and a R. typhi-infected CB17 SCID mouse (right) with HE are depicted. Images were taken at 2-fold magnification. Black arrows point to necrotic lesions. White arrows indicate foci of infiltrating cells (B).

https://doi.org/10.1371/journal.pntd.0004935.g003

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Fig 4. CD11b⁺GR1^low MΦ/monocytes as well as of CD11b⁺GR1^hi neutrophils infiltrate the liver.

CB17 SCID mice were infected s.c. with 2×10⁶ sfu into the base of the tail. Control mice received PBS. At the time of death of R. typhi-infected mice the numbers of cellular isolates from liver (left) and the percentage of CD11b⁺GR1^low MΦ/monocytes (middle) and CD11b⁺GR1^hi neutrophils (right) (y-axis) were determined by flow cytometry. Graphs show combined results from two independent experiments and each dot represents a single mouse (n = 10–11). The mean ± SEM is depicted. Data were analyzed by Student`s t-test (left and right) or Mann Whitney test (middle) after D´Agostino and Pearson normality test. Asterisks indicate statistically significant differences (*p<0.05, *** p<0.001) (A). Serial sections of the liver from a control mouse and a R. typhi-infected CB17 SCID mouse were stained for IBA1, Ly-6G, iNOS and R. typhi as indicated. Pictures show a necrotic area. (B). In addition, several foci of infiltrating IBA1⁺ MΦ from the periphery were observed (C). Immunofluorescent co-stainings were performed for R. typhi (green) and Ly-6G or IBA1 (red) from the liver of an infected CB17 SCID mouse. Nuclei were stained with DAPI (blue) (D).

https://doi.org/10.1371/journal.pntd.0004935.g004

We further performed serial histological sections to clarify the localization of these cells as well as of R. typhi in the liver. For this purpose sections were stained for IBA1 which is exclusively expressed by MΦ [24] and Ly-6G as a marker for granulocytes [25]. In addition, iNOS was stained as an indicator for cellular activation. Enhanced numbers of IBA1⁺ MΦ were found equally distributed in the liver parenchyma of R. typhi-infected CB17 SCID mice, indicating hyperplasia of Kupffer cells. IBA1⁺ MΦ accumulated around necrotic areas (Fig 4B). In addition, several foci of infiltrating IBA1⁺ MΦ from the periphery were observed (Fig 4C). In contrast to MΦ, infiltrating Ly-6G⁺ granulocytes almost exclusively clustered in foci often found nearby necrotic regions (Fig 4B). Expression of iNOS was detectable in IBA1⁺ MΦ as well as in Ly-6G⁺ neutrophils (Fig 4B), demonstrating an activated phenotype of both cell populations. R. typhi was detectable in clusters of infiltrating Ly-6G⁺ neutrophils as well as IBA1⁺ MΦ but not within necrotic tissue (Fig 4B and 4D).

Neutrophils and MΦ harbor R. typhi and show an activated phenotype

Activated phagocytes that exert bactericidal functions release reactive oxygen species (ROS) and express iNOS to produce nitric oxide (NO) [26]. To further assess these functional properties of MΦ/monocytes and neutrophils in R. typhi infection we performed flow cytometric analyses. CD11b⁺GR1^low MΦ/monocytes and CD11b⁺GR1^hi neutrophils from R. typhi-infected CB17 SCID mice were analyzed at the time of death of the animals for ROS release, iNOS expression and bacterial uptake. Non-infected mice that received PBS instead of R. typhi were used as a control. Fig 5A (left) shows a representative flow cytometric analysis of ROS content in CD11b⁺GR1^low MΦ/monocytes (top) and neutrophils (bottom) in the blood. ROS were detectable in around 7% of the CD11b⁺GR1^low MΦ/monocyte population and 3% of the CD11b⁺GR1^hi neutrophils in R. typhi-infected CB17 SCID mice (Fig 5A). We further analyzed bacterial uptake and iNOS expression in CD11b⁺GR1^low MΦ/monocytes and CD11b⁺GR1^hi neutrophils in spleen and liver. Representative dot plots of intracellular staining of R. typhi and iNOS in cells from both organs are shown in Fig 5B and 5C (top). R. typhi was detectable in a high proportion of CD11b⁺GR1^low MΦ/monocytes (9.0±1.2% in the spleen and 2.1±0.7% in the liver) and CD11b⁺GR1^hi neutrophils (11.8±1.9% in the spleen and 2.1±0.5% in the liver), demonstrating ingestion of R. typhi by both cell populations (Fig 5B and 5C). Furthermore, CD11b⁺GR1^low MΦ/monocytes as well as CD11b⁺GR1^hi neutrophils expressed iNOS in spleen (31.5±6.2% of CD11b⁺GR1^low MΦ/monocytes and 2.4±0.3% of CD11b⁺GR1^hi neutrophils) and liver (11.1±2.9% of CD11b⁺GR1^low MΦ/monocytes and 1.3±0.5% of CD11b⁺GR1^hi neutrophils) with MΦ representing the main iNOS-expressing cell population in both organs (Fig 5B and 5C). Thus, MΦ and neutrophils exhibit a bactericidal phenotype.

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Fig 5. Neutrophils and MΦ release ROS and express iNOS.

CB17 SCID mice were infected s.c. with 2×10⁶ sfu into the base of the tail. At the time of death blood, spleen and liver cell samples were analyzed by flow cytometry for ROS release, R. typhi uptake and iNOS expression. A representative dot plot of ROS staining in CD11b⁺Ly-6C^+/-Ly-6G^- MΦ/monocytes (upper panel) and CD11b⁺Ly-6G⁺ neutrophils (lower panel) from a control mouse (left) and a R. typhi-infected mouse (right) is shown. The graph depicts the mean ± SEM of the percentage of ROS-releasing cells (y-axis) among MΦ/monocytes and neutrophils (x-axis) determined in blood samples from two independent experiments (n = 10–11) (below) (A). Representative dot plots of spleen (B) and liver cells (C) gated on CD11b⁺ cells and further stained for GR1, intracellular R. typhi (upper panel) and iNOS (lower panel) are shown. Graphs show the percentage of R. typhi⁺ and iNOS⁺ cells (y-axis) among CD11b⁺GR1^low MΦ/monocytes and CD11b⁺GR1^hi neutrophils (x-axis). Data show the mean ± SEM of combined results from two independent experiments (n = 10–11). Statistical analysis was performed with Mann-Whitney U test (*p<0.05, **p<0.01, ***p<0.001) (B,C).

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Neutrophil depletion neither alters the course of infection nor bacterial load

Recent data show that neutrophils largely contribute to bacterial elimination in the liver [27–30]. Having shown that neutrophils infiltrate the liver of R. typhi-infected CB17 SCID mice, we further elucidated the contribution of neutrophils to bacterial elimination and protection against R. typhi. For this purpose neutrophils were depleted in R. typhi-infected CB17 SCID mice by the application of anti-Ly-6G antibody beginning at day 6 post infection before neutrophils start to rise (Fig 2B). Anti-Ly-6G treatment was repeated every 3 days. R. typhi-infected control mice received equal amounts of isotype antibody instead. Additional control groups of CB17 SCID mice were treated with either PBS or anti-Ly-6G but were non-infected. Success of neutrophil depletion was examined by flow cytometry of blood cells from infected CB17 SCID mice 1 day after the second antibody application. At this point in time (day 10 post infection), CD11b⁺Ly-6G^hi neutrophils represented approximately 65% of the CD11b⁺ cells in the blood of R. typhi-infected control mice and were completely absent in R. typhi-infected CB17 SCID mice that received anti-Ly-6G antibody (Fig 6A). However, neutrophil depletion did not alter the course of disease as monitored by clinical scoring (Fig 6B, left) and mice succumbed to the infection with similar kinetics as control mice (Fig 6B, right). Furthermore, neutrophil-depleted animals showed comparable bacterial burden in spleen, liver, brain and lung as R. typhi-infected control animals that were treated with isotype antibody (Fig 6C).

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Fig 6. Neutrophil depletion completely prevents liver damage in CB17 SCID mice but does not protect from death.

CB17 SICD mice were infected s.c. with 2×10⁶ sfu into the base of the tail. Neutrophils were depleted by intraperitoneal application of anti-Ly-6G antibody. Control animals received rat IgG2a isotype antibody. Depletion was first performed on day 6 post infection and then repeated every 3 days. Efficacy of depletion was assessed by flow cytometric staining of blood cells 1 day after the second depletion (day 10 post infection). A representative dot plot of blood cells from an isotype antibody-treated mouse (left) and a neutrophil-depleted mouse (right) stained for Ly-6G and CD11b is depicted (A). The health status of the mice was assessed over time (x-axis) using a clinical score (y-axis). Four groups of mice were analyzed (non-infected mice that were treated with PBS only (open circles) or PBS and anti-Ly-6G (black circles) and R. typhi-infected mice that were treated with isotype (open squares) or anti-Ly-6G antibody (black squares); n = 9–10 for each group). The data show the mean ± SEM of the clinical score obtained from two independent experiments. Survival rates (y-axis) of anti-Ly-6G- and isotype antibody-treated mice were compared using the Log-rank Mantel Cox test (p = 0.7778; non-significant). Spleens of R. typhi-infected neutrophil-depleted mice were still strongly enlarged. The inserted photograph shows two spleens from non-infected control mice that received PBS (left) and three spleens from neutrophil-depleted R. typhi-infected mice (B). Bacterial burden was determined by PrsA qPCR (y-axis) in liver, spleen, brain and lung (x-axis) of R. typhi-infected isotype-treated (white bars) or anti-Ly-6G-treated CB17 SCID mice (black bars) at the time of death (n = 10). The mean ± SEM is depicted (C). The percentage of CD11b⁺GR1^hi neutrophils (left) and CD11b⁺GR1^low MΦ/monocytes (right) (y-axis) in the spleens from the very same mice were determined by flow cytometry. Each dot represents a single mouse. The mean ± SEM is depicted. Data were analyzed by one-way ANOVA followed by Tukey post-test (left) or Kruskal Wallis test followed by Dunn´s post-test (right) after D´Agostino and Pearson normality test. Asterisks indicate statistically significant differences (*p<0.05, **p<0.01, ***p<0.001) (D). GPT was determined in sera of infected CB17 SCID mice that received isotype (open squares) or anti-Ly-6G antibody (black squares) (n = 8–9) at the time of death. Control animals were treated with anti-Ly-6G but received PBS instead of R. typhi (black circles). Each dot represents a single mouse. Mean ± SEM from combined results of two independent experiments are shown. Data were analyzed by Kruskal-Wallis test followed by Dunn´s post-test after D´Agostino and Pearson normality test. Asterisks indicate statistically significant differences (**p<0.01). Photographs show a representative liver of an isotype-treated (left) and an anti-Ly6G-treated R. typhi-infected CB17 SCID mouse (right) (E). Representative HE stainings of histological sections of the liver of an isotype-treated and an anti-Ly-6G-treated R. typhi-infected CB17 SCID mouse at the time of death are shown. Black arrows point to necrotic lesions. Necrosis was not observed in anti-Ly-6G-treated mice. Open arrows point to cellular infiltrates (F).

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R. typhi-infected CB17 SCID mice still developed splenomegaly in the absence of neutrophils that was comparable to R. typhi-infected mice that were treated with isotype antibody (Fig 6B, insert). We further assessed the cellular composition of the spleen of all groups of mice including non-infected animals that received either PBS or anti-Ly-6G only. As expected, the CD11b⁺GR1^hi neutrophil population was absent in all mice that received anti-Ly-6G, whether infected or not (Fig 6D, left), while a reciprocally, although not significantly enhanced percentage, of CD11b⁺GR1^low MΦ/monocytes was observed in R. typhi-infected CB17 SCID mice in the absence of neutrophils. 53.5±3.8% of the spleen cells in neutrophil-depleted mice were CD11b⁺GR1^low MΦ/monocytes compared to 45.3±1.9% in control animals (Fig 6D, right), indicating that the absence of neutrophils is in part compensated by increased accumulation of MΦ/monocytes.

Neutrophils are responsible for liver damage

Although neutrophils are known to be crucial for pathogen elimination in bacterial infections [31], an impact of these cells in bacterial control was not observed in R. typhi-infected CB17 SCID mice (Fig 6C). However, activated neutrophils can also mediate harmful reactions and can be involved in hepatic injury [32, 33]. We therefore analyzed liver pathology by measuring serum GPT and performing histological stainings. Indeed, neutrophil-depleted R. typhi-infected CB17 SCID mice were completely protected from liver damage. In the absence of neutrophils, R. typhi-infected CB17 SCID mice showed normal serum GPT levels. Livers from these mice appeared healthy and the gall bladders were clear (Fig 6E). Moreover, necrotic lesions were not detectable anymore in histological stainings although still many foci of infiltrating cells were visible (Fig 6F). These were positive for IBA1 and, thus, MΦ (S2B Fig). These results clearly show that neutrophils are solely responsible for liver necrosis in R. typhi-infected CB17 SCID mice.

R. typhi-infected CB17 SCID mice show systemic inflammation which is unaltered in the absence of neutrophils

Although neutrophil-depleted R. typhi-infected CB17 SCID did not show liver damage anymore the animals succumbed to the infection. Thus, liver damage is not the cause of death which must have other reasons. One of these might be an overwhelming immune response. To gain insight into the immune response of CB17 SCID and wild-type BALB/c mice upon R. typhi infection we next analyzed cytokines in the blood. R. typhi-infected CB17 SCID mice showed steadily increasing release of inflammatory cytokines during the course of disease. This response was clearly dominated by IFNγ that reached plasma levels of 1034±180 pg/ml prior to death. In addition, slightly enhanced plasma levels of IL-6 (217±105 pg/ml), IL-12p70 (15±3 pg/ml), TNFα (48±7 pg/ml) and MCP-1 (160±25 pg/ml) were present (Fig 7A). GM-CSF was not significantly enhanced in R. typhi-infected CB17 SCID mice during the course of infection (S3 Fig). These results show that CB17 SCID mice mount a very strong systemic inflammatory response. In contrast, immunocompetent BALB/c wild-type mice produced significantly increased levels of IFNγ (191±70 pg/ml) in addition to MCP-1 (197±81 pg/ml) exclusively at day 3 post infection while IL-12p70, IL-6 and TNFα were not detectable at all in these mice (Fig 7A) and GM-CSF was not significantly elevated during the course of infection (S3 Fig).

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Fig 7. Neutrophil depletion does not alter systemic inflammatory response.

Plasma cytokine levels (y-axis) were measured at indicated points in time (x-axis) in R. typhi-infected BALB/c (open symbols, n = 5–8) and CB17 SCID mice (black symbols, n = 5–8) by bead-based LEGENDplex assay. Data represent combined results from two independent experiments and are shown as mean ± SEM. Statistical significance was determined using Kruskall-Wallis test followed by Dunn´s post-test comparing samples from infected mice with samples from control mice (day 0) (A). Plasma cytokine levels (y-axis) from R. typhi-infected and PBS-treated CB17 SCID mice that either received isotype antibody or anti-Ly-6G as indicated on the x-axis were measured by bead-based LEGENDplex assay at the time of death. Combined results from two independent experiments are shown (n = 8–9). Each dot represents a single mouse. The mean ± SEM is shown. Statistical analysis was performed with Kruskall-Wallis test followed by Dunn´s post-test (B). Asterisks indicate statistically significant differences (*p<0.05, **p<0.01, ***p<0.001).

https://doi.org/10.1371/journal.pntd.0004935.g007

We further clarified the impact of neutrophil-depletion on the inflammatory immune response in R. typhi-infected CB17 SCID mice. Serum levels of IFNγ, IL-12p70, IL-6, TNFα and MCP-1 were increased in all R. typhi-infected animals compared to control mice. Neutrophil-depletion, however, did not alter cytokine production compared to R. typhi-infected CB17 SCID mice that received isotype antibody (Fig 7B), demonstrating minor contribution of neutrophils to systemic inflammation.

NK cells as well as MΦ produce IFNγ and MΦ represent the major source TNFα

Systemic inflammation in R. typhi-infected CB17 SCID mice was clearly dominated by IFNγ, a cytokine that is known to be predominantly produced by NK cells. Therefore, we further assessed the expansion of NK cells in addition to MΦ and neutrophils during the course of disease and cytokine expression by these cell populations. First, blood cells were stained for NKp46, CD11b and GR1 and absolute cell counts of NK cells (NKp46⁺), MΦ (CD11b⁺GR1^low) and neutrophils (CD11b⁺GR1^hi) after pregating on NKp46^- cells were determined. NK cells steadily increased during the course of infection and were enhanced approximately 3.5-fold on day 12 post infection in the blood (Fig 8A). Significantly enhanced numbers of MΦ were observed in the blood rather late in infection on day 12. The increase of these cells, however, was stronger (7-fold) compared to NK cells (3.5-fold) (Fig 8A). In contrast to NK cells and MΦ, neutrophils only transiently increased in the blood from day 3 to day 7 post infection and declined again until day 12 (Fig 8A) when the experiment was terminated.

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Fig 8. Expansion of NK cells, MΦ and neutrophils and cytokine expression by these cell populations.

CB17 SCID mice (n = 8) were infected s.c. with 2×10⁶ sfu into the base of the tail while control animals received PBS (n = 8). Blood cells were analyzed by flow cytometric staining of NKp46, CD11b and GR1. Cells were gated on NKp46⁺ (NK cells) and NKp46^- cells that were further differentiated into MΦ (CD11b⁺GR1^low) and neutrophils (CD11b⁺GR1^hi) as indicated in the example dot plots on the left. Absolute numbers of NK cells, MΦ and neutrophils were determined per μl blood (y-axis) at indicated points in time (x-axis). Measurements from non-infected PBS-treated mice were used as "day 0" control. Statistical analysis was performed with Kruskall-Wallis test followed by Dunn´s post-test (A). 12h before the experiment was terminated (day 12) the animals received brefeldin A to prevent cytokine secretion by the cells in vivo. Spleen cells were stained for NKp46, CD11b, GR1, IFNγ and TNFα and analyzed by flow cytometry. Cells were gated as described above. Absolute cell counts (y-axis) of NK cells, MΦ and neutrophils as indicated on the x-axis were determined. Statistical analysis was performed with Mann-Whitney U test (two-tailed) (B). The percentage of IFNγ- and TNFα-expressing cells among NK cells, MΦ and neutrophils in the spleen of PBS-treated control mice (n = 7) and R. typhi-infected animals (n = 8) was analyzed. The dot plots below show example stainings of the indicated cell population. Graphs on the left and in the middle show the statistical analysis of the percentage of IFNγ-expressing and TNFα-expressing cells (y-axis) among each cell population (x-axis). The graph on the right shows the analysis of the absolute cell count of IFNγ-expressing cells (y-axis) among NK cells and MΦ (x-axis). Statistical analysis was performed with Mann-Whitney U test (two-tailed) (C). Asterisks indicate statistically significant differences (*p<0.05, **p<0.01, ***p<0.001).

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We further analyzed cytokine expression by these cell populations and determined numbers of NK cells, MΦ and neutrophils in the spleen. For this purpose we prepreated the animals with brefeldin A 12 hours prior to the analysis to assess cytokine expression directly ex vivo. Spleen cells were then stained for NKp46, CD11b and GR1 to distinguish NKp46⁺ NK cells and CD11b⁺GR1^low MΦ and CD11b⁺GR1^hi neutrophils among NKp46^- as described above. In addition, cells were stained for intracellular IFNγ and TNFα. First, cell counts of NK cells, MΦ and neutrophils were determined. Numbers of both MΦ and neutrophils were strongly increased in the spleen of R. typhi-infected animals (MΦ: 9.56±1.09×10⁶; neutrophils: 5.00±1.08×10⁶) compared to control mice (MΦ: 0.48±0.13×10⁶; neutrophils: 0.47±0.07×10⁶). Thus, MΦ and neutrophil numbers were approximately 20-fold and 10-fold enhanced at this point in time (Fig 8B). Numbers of NK cells were generally higher compared to MΦ and neutrophils in CB17 SCID mice. 1.01±0.19×10⁶ NK cells were detectable in non-infected CB17 SCID control mice. NK cells significantly increased approximately 4-fold in the spleen of R. typhi-infected mice (4.08±0.72×10⁶) (Fig 8B). This increase corresponds to that observed in the blood (Fig 8A).

TNFα was predominantly detectable in MΦ. 7.08±0.64% of the MΦ expressed this cytokine. A lower proportion of neutrophils also expressed TNFα (2.65±0.28%) while TNFα expression by NK cells was negligible and not significantly enhanced (1.28±0.50%) (Fig 8C, left). However, 9.43±1.02% of the NK cells in the spleen expressed IFNγ. Surprisingly, IFNγ was also detectable in 4.04±1.02% of the MΦ while only 1.24±0.16% IFNγ-expressing cells were present among neutrophils (Fig 8C, middle). The absolute number of IFNγ-producing MΦ and NK cells was comparable in R. typhi-infected mice (Fig 8C). This is explained by the stronger increase of MΦ compared to NK cells (Fig 8B).

These results show that both NK cells and MΦ contribute to IFNγ production and that MΦ are a major source of TNFα in R. typhi-infected CB17 SCID mice.

MΦ and neutrophils do not directly respond to R. typhi in vitro

As MΦ and neutrophils seem to be the dominant cell populations rising in R. typhi-infected CB17 SCID mice and show an activated phenotype, we further clarified if and how these cells react to R. typhi. In the experiment depicted in Fig 5, we first gated on iNOS⁺ cells among CD11b⁺GR1^low MΦ/monocytes and CD11b⁺GR1^hi neutrophils and analyzed the cells for R. typhi content. This analysis should demonstrate whether it is the R. typhi-harboring cells that express iNOS. Surprisingly, the vast majority of the iNOS-expressing CD11b⁺GR1^low MΦ/monocytes was R. typhi-negative (87.7±2.3% in the spleen and 92.8±1.3% in the liver (Fig 9A)) and the proportion of iNOS-expressing cells that contained bacteria was correspondingly small. Similar was also true for CD11b⁺GR1^hi neutrophils. Here, 72.3±4.1% of the iNOS-expressing cells in the spleen and 64.4±8.1% of those in the liver were negative for R. typhi (Fig 9A). Thus, activation of these cells does not correlate with bacterial phagocytosis.

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Fig 9. MΦ hardly respond to R. typhi and are uncapable to kill the bacteria.

To analyze a correlation of bacterial uptake and activation status of CD11b⁺GR1^low MΦ /monocytes and CD11b⁺GR1^hi neutrophils in vivo the respective cell population in the spleen and liver of the mice described in Fig 5 was first gated on iNOS⁺ cells and then analyzed for bacterial content. The graphs show the percentage of R. typhi-positive cells (R. typhi⁺, black bars) and R. typhi-negative cells (R. typhi-, white bars) among the iNOS-expressing cells (y-axis) of the indicated cell population (x-axis). Statistical analysis was performed with Student´s T test (***p<0.001) (A). bmMΦ were infected with indicated amounts of purified bacteria per cell. Control cells were either left untreated (control) or stimulated with 100 ng/ml LPS (x-axis). After 24h cells were analyzed for the presence of R. typhi by flow cytometric staining. The percentage of R. typhi⁺ cells is shown (y-axis). MHCI and CD80 expression and the release of NO, TNFα, IL-12p70, IL-6 and IL-1 were analyzed after 48h (y-axis). Graphs show combined results from four independent experiments. Data were analyzed by one-way ANOVA followed by Tukey´s post-test. Asterisks indicate statistically significant differences compared to untreated cells (*p<0.05, **p<0.01) (B). bmMΦ were plated in chamber slides and either treated with heat-inactivated or living R. typhi as indicated. 10 copies per cell were used. After 48 h cells were stained for R. typhi with anti-R. typhi (BNI52) (green). Nuclei were stained with DAPI (blue). Images were taken at 800x magnification. One view of bmMΦ containing heat-inactivated, degrading bacteria and four views of bmMΦ containing living R. typhi are shown (C). Growth of R. typhi in bmMΦ that were infected with 5 copies per cell was analyzed. Bacterial content (y-axis) was quantified by PrsA qPCR at indicated points in time (x-axis). The graph shows combined results from three independent experiments (D).

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Phagocytes such as MΦ usually recognize bacterial pathogens via pattern recognition receptors such as Toll-like receptors (TLR). These induce a classically activated phenotype including iNOS expression as observed in R. typhi-infected CB17 SCID mice, the release of inflammatory cytokines and the upregulation of costimulatory cell surface molecules [34]. The observation that MΦ and neutrophil activation did not correlate with bacterial uptake in vivo (Fig 9A), however, indicates that the cells may not directly react to the bacteria. To elucidate whether R. typhi activates MΦ, we generated bmMΦ from BALB/c mice and infected the cells with titrated amounts of R. typhi in vitro. Cells were analyzed for bacterial uptake and the expression of MHCI and CD80 by flow cytometry. In addition, NO, IL-6, IL-12 and TNFα were quantified in the supernatant by Griess reaction and LegendPlex assay. Stimulation with E. coli LPS was used as a control. After 24h approximately 12% of the cells that were infected with 25 R. typhi copies per cell were positive for R. typhi as determined by flow cytometry (Fig 9B). MΦ upregulated the expression of MHCI and CD80 48h after infection (Fig 9B). However, the cells neither released detectable amounts of inflammatory cytokines nor NO as it was observed upon stimulation with LPS (Fig 9B). These results show that MΦ do not react to R. typhi in a classical manner and further led to the question whether MΦ are capable to kill R. typhi. To clarify this issue we incubated bmMΦ with either living R. typhi or heat-killed R. typhi to be able to distinguish between living/replicating and degrading bacteria by immunofluorescence microscopy. As expected, fragments of degrading R. typhi particles were detectable after 48h in bmMΦ that received heat-killed bacteria (Fig 9C). In contrast, bmMΦ that were incubated with living R. typhi clearly contained intact and replicating bacterial particles at this point in time. In several cells proliferating bacteria clustered in rosette-like structures while other cells showed a more equal distribution of R. typhi in the cytosol. Interestingly, free R. typhi antigen not bound to particles was also observed in some cells in the cytosol (Fig 9C). Bacterial growth in bmMΦ in vitro was further confirmed by qPCR showing exponential increase of bacterial DNA in the cell culture within 96h (Fig 9D). Collectively these results show that R. typhi does not activate MΦ in a classical manner and that MΦ are incapable to kill the bacteria in vitro.

R. typhi-infected BALB/c mice survive R. typhi infection and show weak liver pathology

The results presented so far show that activated MΦ and neutrophils accumulate in immunodeficient CB17 SCID mice upon R. typhi infection and infiltrate the liver. Moreover, neutrophils revealed to be responsible for liver damage in these animals. To finally show whether these processes may also take place in immunocompetent mice, we again infected BALB/c wild-type mice. Although BALB/c wild-type mice do not show clinical symptoms of disease, histological analyses revealed that R. typhi infection affects the liver. Cellular infiltrates were visible in HE stainings of liver sections beginning on day 3 post infection, peaking on day 7 and declining until day 14 again (Fig 10A). At the peak of cellular infiltration on day 7, necrotic lesions were also visible in the livers of R. typhi-infected BALB/c mice (Fig 10A). Further stainings revealed that enhanced numbers of IBA1⁺ MΦ, Ly-6G⁺ granulocytes as well as T cells were detectable in the liver of R. typhi-infected animals compared to control BALB/c mice that were treated with PBS (Fig 10B). These cells were mainly found in the liver parenchyma. In addition, IBA1⁺ MΦ also accumulated around blood vessels (Fig 10B, below middle). The bacteria and/or bacterial antigen were detectable in endothelial cells (Fig 10B, below right) and in IBA⁺ MΦ (Fig 10B, below left) but not in the liver parenchyma. Liver damage in BALB/c wild-type mice, however, was generally mild. Necrotic lesions were small and rare and enhanced levels of serum GPT were not detectable (S3 Fig).

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Fig 10. Temporary liver damage in BALB/c wild-type mice upon R. typhi infection.

BALB/c mice were infected s.c. with 2×10⁶ sfu R. typhi. Liver sections were analyzed at indicated points in time post infection by staining with HE. Pictures show representative stainings of one out of five mice (A). Serial sections of the liver from PBS-treated BALB/c control mice or BALB/c mice infected s.c. with R. typhi as described above were stained for IBA1, Ly-6G, iNOS and R. typhi employing serum from a patient. For the staining of IBA1 two views are shown. In addition, co-staining of R. typhi (green), IBA1 (red) and DAPI (blue) was performed (B).

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