Human peripheral blood was collected from healthy volunteers with written informed consent. This study was conducted in accordance with the Declaration of Helsinki and all protocols were approved by the Ethics Committee of the University Hospital Jena (permit number: 273-12/09). All animals used for the project were held in accordance with the European Convention for the Protection of Vertebrate Animals Used for Experimental and Other Scientific Purposes and the experiments performed in accordance with European and German regulations. The sacrifice of mice for blood withdrawal was performed under §4 “removal of organs” of the German Animal Welfare Act and was approved by the local animal welfare officer (no specific permit number issued). The systemic candidiasis model was approved by the Thuringian authority and ethics committee (Thüringer Landesamt für Verbraucherschutz, permit number 03-007/13 and HKI-19-003).

C. albicans GFP-expressing strains M137 (18) (pACT1-GFP) and C. albicans Δefg1Δcph1 + pADH1-GFP were used. To construct C. albicans Δefg1Δcph1 + pADH1-GFP a CaGFP-caSAT1 construct with homology regions for the integration into the C. albicans ADH1 locus was excised with AscI/SacI from the plasmid pSK-ADH1prom-CaGFP-SAT1 (4) and then transformed into the EFG1/CPH1 double mutant HLC52 (19). Transformation was performed with the established lithium acetate protocol (20). Transformants were grown for two days on YPD with 200 mg/ml nourseothricine and verified by PCR and microscopy. For experiments C. albicans over-night cultures grown in YPD medium (2% peptone, 1% yeast extract, 2% glucose) at 30°C and 180 rpm were diluted 1:50 in YPD medium and sub-cultured for another 3 to 4 h into the mid-log phase under the same conditions. Yeast cells were washed three times with PBS, counted and diluted in PBS to 5 × 10⁷/ml or 5 × 10⁶/ml, as indicated.

GFP-expressing S. aureus [6850/pALC1743 (21, 22)] and Escherichia coli ATCC 25922 (23) were cultivated overnight at 37°C, 180 rpm in LB medium. The overnight culture was inoculated 1:100 into fresh LB medium and incubated at 37°C, 180 rpm until O.D₆₀₀ 0.6 to 0.7 was reached. The cultures were then washed three times with PBS and bacterial cell numbers were calculated based on OD₆₀₀–CFU correlations. Cultures were diluted to desired concentrations with PBS before inoculation of whole blood.

Human peripheral blood from healthy donors was collected in hirudin-monovettes (Sarstedt, Germany, 2.7 ml volume). Hirudin was used as an anti-coagulant as it was previously shown to have no effect on complement activation (24). Due to their size, direct use of the hirudin-monovettes was not feasible for the collection of murine blood. Therefore, three hirudin-monovettes were rinsed with total volume of 500 µl sterile saline (0.9% NaCl) to solubilize the hirudin. Needles and syringes rinsed with hirudin-NaCl were then used to collected blood from female BALB/c mice (8–12 weeks old, Charles River, Germany) or from C57BL/6J mice (8–12 weeks old, Service Unit Experimental Biomedicine, Friedrich-Schiller-University Jena, Germany) by heart puncture immediately after sacrifice by intraperitoneal application of an overdose of ketamine (500 mg/kg) and xylazin (25 mg/kg). The blood was then immediately transferred to a falcon tube containing 500 µl hirudin-NaCl, in which blood from ten animals was pooled for each experiment. Pilot experiments analyzing fungal CFU in whole blood of female and male C57BL/6J mice revealed no sex-specific differences; thus, both male and female mice were used for the experiments.

The human whole-blood infection assay was performed as described previously (4), and murine blood was infected in the same way. Briefly, 1/50 volume fungal or bacterial suspensions prepared as described above were added to murine or human blood, resulting in an infectious dose of 1 × 10⁶/ml for most experiments or 1 × 10⁵/ml for lower infection dose experiments. The incubation of the whole blood was carried out at 37°C under slow constant motion on a rolling device (Phoenix instrument RS_TR05) for time points from 10 min to 480 min. Survival of the pathogens was determined by plating on agar plates (YPD for C. albicans, LB for E. coli and S. aureus) using serial dilutions in PBS and counting colony forming units. To determine the effect of ketamine/xylazin on phagocytosis and association of C. albicans with immune cells, 40 µg/ml ketamine and 2 µg/ml xylazin were added to human blood directly before the experiment. 40 µg/ml ketamine is equivalent to fivefold the maximal serum concentration in mice after intraperitoneal application of 100 mg/kg ketamine (25).

Generally hematology analysis of murine samples was performed using a BC-5300Vet (Mindray) configured for murine blood. For murine samples, blood smears were prepared for each time point and stained by May-Gruenwald-Giemsa staining (Roth). Filamentation and interaction with platelets were observed by light microscopy. Further analyses were performed by differential staining and subsequent measurement with a flow cytometer. For murine blood, differential staining was performed with CD45-PerCP (Cat. no. 557235, clone 30-F11, 12 µg/ml, BD Biosciences), CD19-APC-Cy7 (Cat. no. 115530, clone 1D3; 12 µg/ml, Biolegend) for B-cells, CD3e-V500 (Cat. no. 560771, clone 500A2, 12 µg/ml, BD Biosciences) for T-cells, Ly6G-PE (Cat. no. 127608, clone 1A8, 12 µg/ml final conc; Biolegend) for neutrophils, NK1.1-BV421 (for C57BL/6J mice: Cat. no. 108731, clone PK136, 12 µg/ml, Biolegend) or CD335-BV421 (for BALB/C mice: Cat. no. 562850, Clone 29A1.4, 12 µg/ml, BD Biosciences) for NK cells, CD11b-APC (Cat. no. 130-091-241, clone M1/70.15.11.5, 30 µg/ml, Miltenyi) for monocytes and neutrophils, CD41 (Cat. no. 133906, clone MWReg30, 12 µg/ml, Biolegend) for platelets, and CD69-PE vio770 (Cat. no. 130-103-944, clone H1.2F3, 30 µg/ml, Miltenyi) and CD62-PE-Vio779 (Cat. no. 130-105-537, clone REA 344, 30 µg/ml, Miltenyi) as platelet activation markers. In parallel, staining with the appropriate isotype controls (APC Rat IgG2b, κ Isotype Control, clone RTK5430, Cat. no. 400611, Biolegend; V500: Syrian hamster IgG2 κ Isotype Control, clone B81-3, Cat. no. 560785, BD Bioscience; PE: rat IgG2a κ Isotype Control, clone RTK2758, Cat. no. 400507, Biolegend; PerCP rat IgG2b κ Isotype Control, clone A95-1, Cat. no. 552991, BD Bioscience; APC-Cy7 rat IgG2a κ Isotype Control, clone RTK2758, Cat. no. 400524, Biolegend; V450 rat IgG2a κ Isotype Control, clone R35-95, Cat. no. 560377, BD Bioscience; V450 mouse IgG2a κ Isotype Control, clone G155-178, Cat. no. 560550, BD Bioscience; PE Vio770 hamster IgG1 Isotype Control, clone G235-2356, Cat. no. 553956, BD Bioscience; PE rat IgG1 κ Isotype Control, clone RTK2071, Cat. no. 400407, Biolegend; APC human IgG1 Isotype Control, clone QA16A12, Cat. no. 403505, Biolegend) was performed as a binding specificity control. After staining the erythrocytes were removed using the BD FACS Lysing solution and fixed samples were analyzed on a FACSVerse (BD Biosciences) flow cytometer after two washing steps in PBS supplemented with 3% FCS.

Human whole-blood was analyzed with a BD FACSCanto II flow cytometer. Platelets were specifically identified by CD42b⁺ staining (mouse anti-human CD42b-APC antibody, clone HIP-1, Cat. no. 303912, BioLegend). Co-staining with mouse anti-human CD66b-V450 antibody (clone G10F5, Cat. no. 561649, BD Bioscience) identified platelets associated with neutrophils. Activation of human platelets was investigated by changes in surface CD62P expression (mouse anti-human CD62P-PE antibody, clone AK-4, Cat. no. 555524, BD Bioscience). In parallel, staining with the appropriate isotype controls (APC mouse IgG1, κ Isotype Ctrl, clone MOPC-21, Cat. no. 400122, BioLegend; V450 mouse IgM, κ Isotype Control, clone G155-228, Cat. no. 560861, BD Bioscience and PE Mouse IgG1, κ Isotype Control, clone MOPC-21, Cat. no. 555749, BD Bioscience) was performed as a binding specificity control. Stained blood samples were treated with BD FACS Lysing solution followed by washing and harvesting cells in BD CellWASH solution. FlowJo10 was used for analysis of all samples.

Neutrophils were isolated from bone marrow of femur and tibia of three to four female 8- to 12-week-old BALB/c mice as previously described (26). Briefly, bone marrow was flushed with RPMI 1640 supplemented with Penicillin/Streptomycin (Sigma) and homogenized using a 40 µm cell strainer. Lysis of erythrocytes was carried out for 1 min on ice using a lysis buffer containing 8 mg/ml NH₄Cl, 1 mg/ml K₂CO₃ and 0.01% EDTA. Neutrophils were purified using a Percoll gradient of 52%, 69%, and 78% in PBS and cells were collected from the 69% to 78% interface. After washing with HBSS neutrophils were resuspended in HBSS, analyzed on a BC-5300Vet (Mindray) and stored on ice until usage for a maximum of 1 h. The purity of isolated neutrophils was roughly 70%.

Six- to eight-week-old female specific-pathogen-free BALB/c mice (16 to 18 g) purchased from Charles River (Germany) were housed in groups of five in individually ventilated cages with free access to food and water. Mice were infected with 2.5 × 10⁴ C. albicans CFU/g body weight in 100 µl DPBS via the lateral tail vein at day 0. Groups of mice were sacrificed at the indicated time points post infection by intraperitoneal application of an overdose of ketamin (500 mg/kg) and xylazin (25 mg/kg). 100 µl of blood were collected under terminal anesthesia by retro-orbital bleeding, immediately transferred into a tube containing 10 µl EDTA solution (1.6 mg/ml), and gently mixed. 20 µl of the sample were analyzed on a BC-5300Vet (Mindray) to determine platelet counts and mean platelet volume. Platelet-rich plasma (RPP) was prepared from 40 µl of whole-blood by centrifugation at 135 g for 15 min at room temperature. To detect platelet activation, platelets were stained for 30 min with fluorescence-labeled antibodies (BioLegend) directed against CD41 (clone HIP8, Cat. no. 303710, 0.1 µg/ml) as platelet marker and CD63 (clone H5C6, Cat. no. 353006, 8 µg/ml) as activation marker, followed by fixation with 1% formaldehyde. Surface expression of CD62P, fibrinogen binding and C3c binding were determined as described for CD63 using fluorescence-labeled antibodies (CD62P: BioLegend, clone AK4, Cat. no. 304906, 1.5 µg/ml; fibrinogen: BioRad, polyclonal, Cat. no. 4440-8004F, 100 µg/ml; C3c: Dako, polyclonal, Cat. no. F0201, 400 µg/ml). Non-activated unstained cells and non-activated and activated stained cells were used to calibrate the flow cytometer using single stains and combined stains. Plasma was prepared by centrifugation of 40 µl whole-blood at 1500 g for 15 min at room temperature. Plasma concentrations of soluble CD62P were determined using the Quantikine^® ELISA Mouse sP-Selectin/CD62P kit (R&D Systems, USA) performed according to manufacturer instructions.

Venous blood of healthy human volunteers was collected in sodium citrate monovettes (Sarstedt) and directly centrifuged at 80 × g for 20 min (without break). Supernatants were collected and treated with 0.25 mg/ml acetylsalicylic acid (Sigma Aldrich) for 30 min followed by addition of 1 mM Prostaglandin E1 (Sigma Aldrich), both to prevent pre-activation of the containing platelets. Centrifugation at 400 × g for 8 min pelleted platelets that were afterwards suspended in HEPES-Tyrodes buffer (10 mM HEPES, 137 mM NaCl, 2.8 mM KCl, 1 mM MgCl₂6H₂O, 12 mM NaHCO₃, 0.4 mM Na₂HPO₄2H₂O, 5.5 mM Glucose, 0.35% BSA, without Ca²⁺). Purified platelets were treated again with 1 mM Prostaglandin E1, centrifuged at 400 × g for 10 min and suspended either in RPMI 1640 medium containing 20% autologous active or heat-inactivated plasma. Autologous human plasma was collected after centrifugation of Hirudin-anticoagulated blood from the same donor at 16,000 × g for 10 min. To inactivate complement proteins, autologous human plasma was incubated for 1 h at 56°C. Murine platelets were prepared from freshly collected blood of healthy mice as concentrates by thrombocytapheresis with Amicus cell separator (Baxter, Vienna, Austria) by the Department of Immunology and Blood Transfusion (Innsbruck Medical University, Innsbruck, Austria). The platelet concentration was determined by a hemocytometer and adjusted to a concentration of 1.2 to 1.4 × 10⁹/ml. Murine serum was collected by centrifugation of blood samples from (i) SPF mice without any treatment, (ii) mice systemically infected with a sublethal dose of 1 × 10⁴ C. albicans CFU/g body weight on days 8 to 21 after infection (see 2.6), (iii) C3-deficient mice (B6.129S4-C3^tm1Crr/J). Pooled serum of 5 to 12 mice per group was used for experiments with isolated platelets.

Confrontation of isolated platelets with C. albicans was performed at a ratio of platelets to fungal cells of 10:1 for the indicated time points (37°C, constant rolling at 5 rpm). Mock-infected platelets with the two different media served as controls.

Plasma samples were generated from whole-blood aliquots that were incubated on ice for 45 min to 1 h and subsequently centrifuged at 4°C for 10 min at 10,000 × g. Supernatants were stored at −80°C until further use. Concentration of IFN-γ, IL-1β, TNF-α, IL-6, and KC (CXCL1) in murine samples was determined by ELISA (Invitrogen, Thermo Fisher Scientific) performed according to manufacturer’s instructions.

The fate of C. albicans upon exposure to murine blood was analyzed in whole blood collected from two of the most commonly used mouse strains for infection experiments, BALB/c and C57BL/6J. As described by Hünniger et al. for the human whole-blood model (4), whole blood was inoculated with 1 × 10⁶/ml of yeast-grown C. albicans and fungal survival was determined by counting colony forming units (CFU) at various time points over a time course of 3 h. In contrast to human blood in which a 50% decrease of the initial CFUs within the first hour of contact was observed (4), the fungal burden did not decrease during incubation in neither BALB/c nor C57BL/6J mouse blood but remained stable throughout the experiment ( Figures 1A, B ). Of note, ketamine and xylazin were used to euthanize mice prior to blood collection. As ketamine has been shown to affect antimicrobial functions of macrophages and neutrophils in a dose-dependent manner (27–30), we added ketamine and xylazin to human blood and quantified fungal killing. At the used dose (40 µg/ml ketamine and 2 µg/ml xylazin) ketamine/xylazin treatment did not affect fungal killing ( Figure S1 ).

In order to determine whether the higher fungal survival in murine compared to human blood was associated with differences in the interaction with immune cells, we analyzed the number of immune cells and association of C. albicans with leukocytes. Quantification of total leukocyte numbers using a hematology analyzer showed a moderate, non-significant decline over time in both mouse strains ( Figure S2A ). While the numbers of CD45⁺ leukocytes in relation to all blood cells showed no obvious changes throughout the infection ( Figure S2B ), the numbers of neutrophils decreased by almost 50% within the first 10 min after addition of C. albicans ( Figure S2C ). This was not the case for mock-infected control samples ( Figure S2D ). In contrast, the number of other types of leukocytes remained stable over time ( Figure S3 ). Note that the relative abundance of neutrophils was significantly lower in the blood of B57BL/6J (4.56 ± 1.3%) than of BALB/c mice (8.63 ± 1.58%; p = 0.02).

Using a strain constitutively expressing green fluorescent protein the association of C. albicans with leukocytes was determined by flow cytometry. Within 30 min, more than 50% of C. albicans cells were associated with immune cells in BALB/c whole blood ( Figure 1C ). The proportion of fungal cells associated with immune cells remained relatively stable until the end of the experiments ( Figure 1C ). A similar trend was observed in blood of C57BL/6J mice; however, the rate of association was lower and did not exceed 40%. Surprisingly, a large proportion of C. albicans cells were found to be associated with B- and T-cells in both mouse lines ( Figures 1C, D ). This contrasts results from the human whole-blood model in which monocytes and neutrophils were the dominant types of immune cells physically interacting with C. albicans (4). Since the ratio of the different immune cell populations in blood differs significantly between humans and mice, we speculated that the association to lymphocytes could be the consequence of the higher abundance of these types of immune cells in murine blood rather than the result of specific interactions. We thus calculated the percentage of host cells interacting with C. albicans within the different immune cell populations ( Figures 1E, F ). Although only a small fraction of C. albicans cells were associated with neutrophils, up to 50% and 40% of all neutrophils associated to fungal cells in blood from BALB/c and C57BL/6J mice, respectively. In contrast, only a minor fraction of the B- and T-cell populations interacted with the fungus, suggesting that the observed association is indeed a stochastic physical event rather than the result of specific interactions.

Neutrophils are the most active immune cells interacting with C. albicans in both human and murine blood, and have been shown to be critical for killing of the fungus in the human whole-blood model (4). However, the absolute number of neutrophils is approximately 10-fold lower in murine (0.4 × 10⁶/ml) than human blood (4 × 10⁶/ml; according to reference values and determined by a hematology analyzer). Using the same infection dose (1 × 10^6/ml) in both models thus resulted in a pathogen to neutrophil ratio of 2.5 in murine but 0.25 in human blood. We hypothesized that the higher microbe to neutrophil ratio in the murine whole-blood model overwhelmed the capacity of the neutrophils to interact with and control C. albicans. Therefore, we analyzed fungal survival in whole blood of BALB/c mice using a 10-fold lower infection dose (1 × 10^5/ml), which is comparable to the ratio used previously in human whole blood (4). With the reduced infection dose there was a moderate reduction of the fungal burden by 30% within 3 h of infection ( Figure 2A ). A similar effect was observed when murine blood was supplemented with neutrophils isolated from bone marrow to reach cell numbers comparable to human blood (4.4 ± 0.51 × 10⁹/L of cells, determined by a hematology analyzer) ( Figure 2B ). Thus, both the addition of external neutrophils and the lower infection dose led to increased fungal killing, but survival of C. albicans after 180 min was still substantially higher (>40%) than in a human whole-blood model (10% (4);). Furthermore, fungal burden decreased only slowly in murine blood, whereas 50% of fungal cells were killed within the first 60 min in human blood (4). To exclude effects mediated by ketamine/xylazin, ketamine and xylazin were added to human blood; this did not affect association rates with neutrophils and monocytes ( Figure S4 ).

The lower number of neutrophils in murine compared to human blood did not fully explain the lower killing of C. albicans in murine blood. However, functional differences between human and murine neutrophils have been described: A previous study from Ermert et al. (31) described a reduced efficiency of murine neutrophils to kill Candida species due to the fact that mice lack β-defensins and produce lower amounts of myeloperoxidase. They also showed that internalized C. albicans can escape from murine but not from human neutrophils by outgrowing neutrophils through filamentation and subsequent rupture of the neutrophil membrane (31). We therefore analyzed C. albicans morphology in whole blood by microscopy. Following infection with yeast cells, after 60 min the majority of C. albicans (~70%) in murine blood were hyphae and after 180 min nearly all (>90%) fungal cells grew as filaments ( Figure 3A ). In contrast, hyphal morphology was observed only for 10% and 20% of all C. albicans cells in human blood after 60 and 180 min, respectively ( Figure 3B ).

C. albicans morphology has been shown to affect recognition by neutrophils (5, 32) and monocytes (33). Furthermore, while human neutrophils prevent C. albicans filamentation and escape following phagocytosis (31), human monocytes and macrophages do not, and intracellular filamentation leads to immune cell lysis and fungal escape (34). Given the higher rate of C. albicans filamentation observed in murine blood, and the role of morphology for immune escape, we used a yeast-locked C. albicans Δefg1/Δcph1 mutant expressing GFP (19) in the murine whole-blood infection model to determine if the higher filamentation rate could explain higher survival of C. albicans in murine blood. Surprisingly, no decrease in CFU counts was observed with this strain and fungal numbers even increased 2.5-fold from 60 to 180 min ( Figure 4A ). The increasing fungal burden correlated with a decrease in CD45⁺ cells ( Figure 4B ), and especially neutrophils ( Figure 4C ), at later time points (150 min and 180 min). The percentage of C. albicans Δefg1/Δcph1 cells not associated with any of type of immune cell (~80%, Figure 4D ) was significantly higher compared to the wild type strain (~50%, Figure S2A ). This is in contrast to experiments with human blood, in which similar association of wild type and mutant with immune cells was observed (4, 5). However, more than 25% of all neutrophils were associated with this mutant after 2 h ( Figure 4E ), and the release of cytokines upon infection with the mutant was comparable to or higher than C. albicans wild type infection ( Figure S4 ). Thus, the large number of free Δefg1/Δcph1 cells cannot be explained by an inability of immune cells to recognize mutant yeast cells.

In addition to fungal filamentation, microscopic analysis revealed aggregates of platelets around C. albicans cells ( Figure 5A ). This interaction was confirmed and quantified by flow cytometry ( Figure 5B ): Within 10 min almost 60% of C. albicans cells were associated with platelets. The association rate slowly decreased over time to 40% after 3 h. At early time points, over 35% of the fungal-associated platelets expressed the activation marker CD69 on their surface ( Figure 5C ). We also observed increased expression of CD62 on the surface of C. albicans bound platelets, however in only 20% of all platelets ( Supplementary Figure S6 ). To exclude that platelet activation was an artefact of the ex vivo situation, blood samples of intravenously (i. v.) infected mice were analyzed. In vivo, platelet counts increased significantly over a course of three days post infection ( Figure 6A ). Transient platelet activation was observed at 6 h post infection characterized by a significant increase of complement C3c, indicating opsonization, the platelet aggregation factor fibrinogen, and the activation markers CD63 and CD62P on the surface of platelets ( Figures 6B–E ). While at later time points expression of these markers was comparable to control samples, increased concentrations of soluble CD62 were detectable in the plasma of infected mice from 1 to 3 days post infection by ELISA ( Figure 6F ). Taken together, these data demonstrate interaction and activation of platelets following infection with C. albicans in murine blood ex vivo and in vivo.

Our next aim was to determine if interaction with thrombocytes also occurs in human blood. Consistent with previous findings (4), we observed decreasing numbers of C. albicans cells not associated with host cells in human blood over time ( Figure 7A ). Only a small proportion (less than 10%) of fungal cells directly associated with platelets ( Figure 7B ). This was in sharp contrast to the results we obtained in murine blood. To investigate the potential influence of human blood components on fungal-platelet interaction, confrontation of isolated human platelets with C. albicans cells was performed in the presence of non-treated (active) or heat-inactivated autologous plasma and showed a time-dependent increase in platelet-C. albicans interaction when plasma proteins were inactivated ( Figure 7C ). In contrast, the presence of non-treated plasma containing active complement proteins resulted in a low platelet association. Consequently, we tested if opsonization of fungal cells interferes with platelet binding. Indeed, pre-opsonized C. albicans showed a significantly reduced interaction with isolated platelets compared to non-opsonized C. albicans ( Figure 7D ) indicating that opsonization of extracellular C. albicans during human whole-blood infection prevents the binding of platelets. In a comparable experiment using isolated murine thrombocytes and murine serum, active but not inactive serum significantly increased the binding of thrombocytes to C. albicans ( Figure 8A ).

The observation that inactivation of human plasma increases interaction of thrombocytes with C. albicans suggests that complement factors interfere with binding of human platelets to the fungus. While it remains unclear how complement becomes activated in active human plasma upon contact with C. albicans, the classical, antibody-mediated complement activation pathway might be involved. Specific-pathogen free mice, as used in this study, are usually not colonized by C. albicans (35) and show comparatively low colonization with fungi (36), resulting in very low detectable anti-Candida antibodies in SPF mice (37). Thus, due to the absence of antibodies, whole murine blood and serum might lack the level of opsonization mediated by normal human plasma and, in consequence, lower opsonization could result in higher interaction with platelets. To test this hypothesis, we compared the binding of isolated thrombocytes to C albicans in the presence of active serum from normal SPF mice (naïve mice) and mice that survived a sublethal systemic C. albicans infection (reconvalescent mice). The binding of thrombocytes to fungal cells was significantly higher with serum from naïve mice; furthermore, naïve serum and serum from C3-deficient mice induced similar binding of thrombocytes to C. albicans ( Figure 8B ), supporting the hypothesis that the lack of prior contact to the fungus in SPF mice contributes to the observed differences between human and murine whole blood regarding interaction with thrombocytes.

In order to determine whether the inability of murine blood to control infections ex vivo is restricted to C. albicans, we performed infections with S. aureus and E. coli as models for Gram-positive and Gram-negative bacteria, respectively. Similar to our observations with the filament-deficient C. albicans strain, CFU counts for both bacterial pathogens remained stable for the first 90 min followed by a steady increase ( Figure 9A ). This increase in pathogen load was even more pronounced than that observed for C. albicans Δefg1 Δcph1 ( Figure 4A ), and was also seen with a 10-fold lower infection dose, albeit at a lesser extent ( Figure S7 ). Also similar to infection with the yeast-locked C. albicans mutant was the reduction of leukocyte numbers ( Figure 9B ), and for S. aureus especially neutrophil counts ( Figure 9C ), at late time points. The association of bacteria to immune cells varied quite drastically between both bacterial strains: While almost 60% of neutrophils were rapidly bound to S. aureus within 10 min and almost all neutrophils were attached to these Gram-positive bacteria 1.5 h post infection ( Figure 9D ), bacterial host cell association occurred much later for E. coli and a maximum of 50% of neutrophils were attached to bacteria at 2 h post infection ( Figure 9E ).