Bronchoscopic fibered confocal fluorescence microscopy for longitudinal  in vivo  assessment of pulmonary fungal infections in free-breathing mice

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Abstract

Respiratory diseases, such as pulmonary infections, are an important cause of morbidity and mortality worldwide. Preclinical studies often require invasive techniques to evaluate the extent of infection. Fibered confocal fluorescence microscopy (FCFM) is an emerging optical imaging technique that allows for real-time detection of fluorescently labeled cells within live animals, thereby bridging the gap between in vivo whole-body imaging methods and traditional histological examinations. Previously, the use of FCFM in preclinical lung research was limited to endpoint observations due to the invasive procedures required to access lungs. Here, we introduce a bronchoscopic FCFM approach that enabled in vivo visualization and morphological characterisation of fungal cells within lungs of mice suffering from pulmonary Aspergillus or Cryptococcus infections. The minimally invasive character of this approach allowed longitudinal monitoring of infection in free-breathing animals, thereby providing both visual and quantitative information on infection progression. Both the sensitivity and specificity of this technique were high during advanced stages of infection, allowing clear distinction between infected and non-infected animals. In conclusion, our study demonstrates the potential of this novel bronchoscopic FCFM approach to study pulmonary diseases, which can lead to novel insights in disease pathogenesis by allowing longitudinal in vivo microscopic examinations of the lungs.

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Cryptococcal Cell Morphology Affects Host Cell Interactions and Pathogenicity

Laura Okagaki, Anna K. Strain, Judith N. Nielsen … (2010)

Introduction Unicellular organisms exhibit morphological changes under a wide variety of environmental conditions. In many pathogenic fungi, the ability to switch cell morphology is integral to the infection cycle. Dimorphic fungi, such as Blastomyces dermatitidis and Histoplasma capsulatum, grow in the environment in a hyphal form. When a susceptible host inhales spores, these fungi grow as yeasts. This change in morphology is induced by the high mammalian body temperature [1], [2], [3]. Other pathogenic fungi, such as Candida albicans and Coccidioides immitis, change to specific cell morphologies based on environmental cues or stage of infection [4], [5], [6]. Morphological changes in the pathogenic fungus C. albicans affect tissue tropism and dissemination. Hyphal cells are important in the invasion of host tissues, while yeast cells can easily disseminate through the blood and lymph systems to spread the infection [5], [7]. Additionally, phagocytosis of yeast cells induces differentiation into hyphal cells [6]. Cryptococcus neoformans is an opportunistic fungal pathogen that is most commonly associated with disease in immunocompromised patient populations, such as HIV/AIDS patients, transplant recipients, patients with lymphoid disorders, chronic treatment with corticosteroids, or patients undergoing certain types of chemotherapies [8], [9], [10]. C. neoformans presents clinically as skin lesions, pneumonia, or meningitis [11]. Over 30% of the HIV/AIDS population in Sub-Saharan Africa present with cryptococcal meningitis and cryptococcosis is currently the fifth leading cause of fatalities in this region [12]. Infection with C. neoformans begins when desiccated yeast cells or spores are inhaled and lodge in the alveoli of the lungs. Cryptococcosis occurs when yeast cells disseminate to the bloodstream and ultimately penetrate the blood-brain barrier (BBB) [10], [13]. While the exact mechanism for trafficking from the lungs to the central nervous system (CNS) remains unknown, interactions with host phagocytes and the endothelial cells of the BBB have been shown to be important in this process [14], [15], [16], [17], [18], [19], [20], [21], [22]. Morphogenesis in C. neoformans has primarily been observed as a result of pheromone signaling and mating [23], [24]. There are two varieties of C. neoformans: neoformans and grubii. Historically, mating has been studied in vitro in var. neoformans even though the vast majority of human cryptococcosis cases are caused by var. grubii. C. neoformans has two mating types: a and α. Mating is initiated when pheromone (a or α) secreted by one mating type binds to the pheromone receptor, Ste3α or Ste3a respectively, of the other mating type to trigger a mitogen-activated protein kinase (MAPK) signaling cascade [23], [25]. Pheromone signaling results in morphological changes in var. neoformans, including germ tube formation by mating type α cells and enlargement of mating type a cells [23], [24]. Pheromone-induced MAPK signaling ultimately results in fusion of a and α cells followed by dikaryotic filamentation. Dikaryotic hyphae eventually give rise to basidia where nuclear fusion occurs and meiosis produces haploid spores [23], [26]. In var. grubii, no in vitro morphogenesis in wild-type strains has been observed during early pheromone signaling, although hyphal formation and basidium production mimic that seen in var. neoformans [27]. In this study, we show that cell enlargement is observed in vivo in var. grubii, and that this cell enlargement can be regulated by pheromone signaling. Additionally, we show that these morphological changes in cell size affect pathogenicity by altering phagocytosis and dissemination to the central nervous system (CNS). Finally, we characterized DNA content of this novel cell type to reveal that these enlarged cells are polyploid. Results Morphological changes in C. neoformans var. grubii cells in vivo Pheromone signaling in C. neoformans is known to cause morphological changes including formation of conjugation tubes, dikaryotic filaments, and production of basidia and spores [23], [24], [26], [28]. Mating type a cell enlargement has also been observed in confrontation assays [23]. Cell enlargement has been observed in both human and mouse specimens [29], [30], [31], [32]. Thus, we systematically analyzed cellular morphology in various tissues of mice intranasally infected with var. grubii mating type a or α strains or mice coinfected with both mating types to determine the effect of pheromone signaling and mating type on in vivo cell morphology. Histopathologic tissue sections from the lungs, heart, spleen, liver, kidneys, and brain at 1, 2, 3, 7, 14, and 21 days post-infection were examined for changes in cryptococcal cell morphology. Dramatic changes in cryptococcal cell size were observed in the lungs, although a few cells with increased cell size were also observed in the spleen and brain at late time points ( Figure 1A , Figure S1 ). Most fungal cells in the lungs remained small (5–10 µm in diameter) resembling yeast cells grown in rich medium in vitro. However, a proportion of the cryptococcal cells in the lungs were much larger. For ease of reference, we designated this group of enlarged cryptococcal cells as “titan” cells. These titan cells were >10 µm in diameter, with some cell sizes approaching 50 to 100 µm in diameter ( Figure 1A ). Titan cell diameter measurements were based on actual cell body size and excluded capsule changes which were highly variable. Titan cells were observed as early as 1 day post-infection in the lungs, accounted for approximately 20% of the cryptococcal cells in the lungs by 3 days post-infection, and remained relatively constant throughout the rest of the infection ( Figure 1B, 1C ; Figure S1 ). Titan cells were occasionally observed in the spleen and brain but at low levels ( Figure S1 ). In contrast, coinfection with both mating types resulted in an increase in titan cell production to almost 50% of the cells present in the lungs ( Figure 1B, 1C ). 10.1371/journal.ppat.1000953.g001 Figure 1 Titan cells in the lungs of coinfected mice. A) Mice were coinfected with an approximate 1∶1 ratio of a:α intranasally at a final concentration of 5×104 cells. Lung sections were stained with periodic acid Schiff (PAS) 14 days (top) or 3 days (bottom) post-infection. White arrow denotes C. neoformans cells >10 µm in diameter. Black arrow denotes cells ≤10 µm in diameter. Top: bar = 100 µm, bottom: bar = 10 µm. B) The number of small cells (≤10 µm) and titan cells (>10 µm) were quantified in single and coinfections at 7 days post-infection. >500 cells were counted per treatment per mouse. Error bars indicate SD from 3 mice per treatment. Asterisk indicates p 80 cells were analyzed per mouse per treatment. Data are representative of three independent experiments with three mice per treatment. Error bars indicate SD. Asterisk indicates p 500 cells were examined per animal. Error bars indicate SD from four mice per treatment. Asterisk indicates p 0.2 were observed for other pair-wise comparisons. Because pheromone signaling induces mating type a cell enlargement during in vitro mating of var. neoformans, we hypothesized that the increase in titan cell formation during coinfection was specific to mating type a cells. To test this hypothesis, we differentially stained a cells with AlexaFluor 488 (green) and α cells with AlexaFluor 594 (red) prior to intranasal inoculation of mice. Mice were sacrificed at 1–3 days post-infection, and unstained histopathological sections were examined for cryptococcal cell fluorescence ( Figure 2 ). At 1 day post-infection, no difference in the proportion of a or α titan cells in individual or coinfections was observed (data not shown). However, at 2–3 days post-infection, the proportion of mating type a titan cells in coinfections increased while the α titan cell proportion remained equivalent to the individual infections ( Figure 1C ). Almost half of the stained mating type a cells in coinfected lungs had converted to titan cells. 10.1371/journal.ppat.1000953.g002 Figure 2 Fluorescently labeled a (green) and α (red) cells in the lungs during coinfection. C. neoformans a and α strains were combined with AlexaFluor 488 (green) or AlexaFluor 594 (red), respectively, and incubated for 20 minutes. Cells were washed with PBS to remove excess dye. Mice were inoculated with an approximate 1∶1 ratio of a:α cells at a final concentration of 5×107 cells. At 2 days post-infection animals were sacrificed, lungs extracted, fixed in 10% buffered formalin, paraffin-embedded, and 5 µm sections generated. Host tissues are autofluorescent at both wavelengths resulting in a yellow color upon overlay. White arrows denote fluorescent C. neoformans cells. Bar = 20 µm. To further quantify titan cell formation during coinfection, cells were differentially stained green with AlexaFluor 488 prior to intranasal instillation with the following treatments: a only (green), α only (green), a(green)/α, or a/α(green). At 3 days post-infection, bronchoalveolar lavage (BAL) was performed. The resulting mix of cryptococcal and mouse cells was immediately fixed and the proportion of green titan cells was determined by microscopic examination ( Figure 1D , see below). Similar to the tissue sections, approximately 20% titan cells were observed in the individual infections with no difference in titan cell formation between the two mating types (p = 0.2, Figure 1D ). In the coinfections, mating type α titan cell formation remained at the basal level (p>0.64, Figure 1D ) while mating type a titan cell formation increased (p 0.6, Figure 1D ). Thus, the increase in titan cell formation by mating type a cells during coinfection requires the Ste3a receptor. However, the presence or absence of the Ste3a pheromone receptor has little effect on the basal level of titan cell formation observed in individual infections. Pheromone signaling alters dissemination to the central nervous system To examine the role of titan cell formation in pathogenicity, individual and coinfections with the wild-type and ste3 a Δ mutant strains were compared. The Cryptococcus infectious cycle can be divided into three stages: an initial pulmonary infection (lungs), dissemination (spleen), and penetration of the CNS (brain). Previous studies with C. neoformans var. grubii congenic strains showed no differences in virulence between the a and α mating types [26]. However, coinfection with both mating types simultaneously resulted in reduced a cell penetration of the CNS [33]. Interestingly, while a cell CNS penetration was reduced compared with α cells, both cell types had equivalent accumulation at the first two stages of infection. During coinfection, only mating type a cells displayed an increase in titan cell formation and a subsequent reduction in CNS penetration. Thus we hypothesized that pheromone signaling and the resulting increase in titan cell formation reduces a cell CNS penetration. To determine whether pheromone signaling affected dissemination to the brain during coinfection, we compared wild-type and ste3 a Δ mutant strains for CNS penetration when coinfected with α ( Figure 3 ). In both wild-type and ste3 a Δ coinfections, the number of a and α cells recovered from the spleen and lungs was equivalent to the proportion of the two cell types in the initial inocula (p>0.1). These data show, even at late time points, alterations in titan cell production in response to pheromone signaling do not affect persistence of the cells in the lungs. However, a significant decrease was seen in the proportion of wild-type a cells recovered from the brain (p = 0.001, Figure 3A ). In contrast, coinfections with the ste3 a Δ mutants restored a cell accumulation in the CNS to levels equivalent to the initial inocula (p>0.4, Figure 3B, C ). Both independent ste3 a Δ mutants showed similar results. Together, these data suggest that pheromone signaling during a/α coinfection affects the pathogenicity of a cells by increasing titan cell formation which inhibits the ability of a cells to establish a CNS infection. 10.1371/journal.ppat.1000953.g003 Figure 3 C. neoformans pheromone receptor mutant strains penetrate the CNS during coinfection. Mice were coinfected intranasally with an approximate 1∶1 ratio of A) a:αNAT, B) ste3 a Δ#1:α, or C) ste3 a Δ#2:α at a final concentration of 5×104 cells. The actual proportion of a cells in the infecting inoculum was determined by growth on selective medium and is plotted as a horizontal dashed line. At 21 days post-infection animals were sacrificed, the lungs, brain, and spleen were homogenized and serial dilutions plated. >500 colonies per organ per mouse were isolated and assayed for drug resistance to determine mating type. The proportion of a cells is plotted with open circles denoting values from individual animals and bar height representing the geometric mean. To determine P-values, Wilcoxon rank sum analysis was performed on the measured number of a and α cells compared with the expected number, assuming that both strains remained at the initial inoculum proportions. Coinfection does not affect blood brain barrier penetration upon IV injection An in vivo murine tail vein injection model was employed to determine whether coinfection disrupts CNS penetration by reducing a cell interactions with the endothelial cells of the BBB [34], [35]. In this model, cells bypass the lungs and are injected directly into the bloodstream via the mouse tail vein. The cells then lodge in the small capillaries of the brain and cross the endothelial cell layer of the BBB. To test whether interaction with the BBB was directly affected by mating type or coinfection, a and α cells were fluorescently labeled and examined for their interactions with the BBB. Both cell types were able to traffic to the small capillaries of the brain ( Figure 4A ) and quantification revealed equal proportions of the two mating types in the capillaries (data not shown). During coinfection, the two mating types were observed in close proximity approximately 25% of the time, consistent with random interactions between cells in a mixed population. The finding that cells of opposite mating type are found in close association would enable pheromone signaling to occur between them in the capillaries of the brain ( Figure 4B ). Both mating types could induce phagocytosis by the endothelial cells of the BBB ( Figure 4C ). Capsule structural changes are important for interactions with the endothelial cells of the BBB [35]. These structural changes can be characterized by alterations in anti-capsular antibody binding. The binding patterns to the cryptococcal capsule for two monoclonal antibodies, E1 and CRND-8, recognizing distinct epitopes on the capsular polysaccharide were studied over time and found to be similar for both mating types. Cells observed in the capillaries shortly after inoculation and up to 6 hours post-infection exhibited only E1 antibody binding. In contrast, cells observed in the brain parenchyma were mostly labeled with CRND-8, as described previously for KN99α [35]. No difference in the capsular antigen staining or the kinetics of capsular changes upon crossing of the BBB were observed between the a and α cells during interactions with the endothelial cells of the BBB – either alone or during coinfection (data not shown). These data suggest that the inability of a cells to penetrate the CNS during coinfection is not due to innate differences between the two cell types or their interactions with the BBB itself, but instead may be due to an inability of the a cells to traffic appropriately from the lungs to the brain. 10.1371/journal.ppat.1000953.g004 Figure 4 KN99a and KN99α cells interact with endothelial cells of the blood-brain barrier during coinfection. a or α were combined with AlexaFluor 350 (blue), AlexaFluor 488 (green) or AlexaFluor 594 (red) and incubated for 20 minutes. Mice were inoculated by tail vein injection with an approximate 1∶1 ratio of a:α at a final concentration of 2×107 cells. At 1 day post-infection, animals were sacrificed and 50 µm frozen brain sections were obtained. A) Sections from mice infected with a (green) and α (blue) were immunostained with anti-collagen IV primary antibody (endothelial cell membrane) with a TRITC (red) labeled secondary antibody. Bar = 20 µm B) Sections from mice infected with a (green) and α (red) were imaged by confocal microscopy and sections were compiled as a projection. Bar size = 20 µm C) Frozen sections from mice infected with a (red) and α (green) were treated with Hoechst (host cell nuclei), imaged by confocal microscopy, and sections were compiled as a 3D rendering. The U-shaped nuclei are indicative of endothelial cells containing cryptococcal cells. Bar = 10 µm. Titan cells are resistant to phagocytosis One of the first lines of defense by the host immune system is phagocytosis and the resultant killing of pathogens by mononuclear macrophages and monocytes in the lungs. These host cells identify pathogens, phagocytose them, and either kill the pathogen outright via oxidative and/or nitrosative bursts or present antigens to T cells for further activation of the host immune response [36]. Recent studies suggest phagocytosis by monocytes or macrophages is important for subsequent CNS penetration [16], [18], [20], [22]. Thus, we examined titan cell interactions with lung host immune cells. Fixed BAL samples were analyzed microscopically for yeast cell interactions with host phagocytic cells. Titan cells were never observed inside host phagocytes, presumably due to their large size. Engulfed small cryptococci were observed inside phagocytic host cells ( Figure 5A, B ). No difference in phagocytosis was observed between mating types (p = 0.82, Figure 5D ). The percentage of intracellular α cells during coinfection was similar to that observed in single mating type infections (p>0.89, Figure 5D ). In contrast, a decrease in the percentage of phagocytosed a cells was seen during coinfection (p 0.53 Figure 5D ). Interestingly, titan cells were often surrounded by one or more host immune cells ( Figure 5C ). Yet complete phagocytosis of titan cells was not observed upon characterization of these cellular interactions by confocal microscopy (data not shown). Taken together, these data indicate titan cell formation was negatively correlated with phagocytosis by host immune cells. 10.1371/journal.ppat.1000953.g005 Figure 5 Titan cell formation and phagocytosis in the lungs of infected mice. Mice were intranasally infected with either a, α, or ste3 a Δ cells labeled with AlexaFluor 488 (green) or coinfected with one labeled and one unlabeled strain (four mice per treatment). Cells obtained by BAL were fixed, stained with DAPI, and examined by microscopy for green fluorescence (cell type) and cell size. >500 cells were examined per animal. Bar = 10 µm A) C. neoformans a cells (green) ≤10 µm in diameter were visible inside host phagocytes. Host cells were identified by large blue DAPI stained nuclei. B) Several small α (≤10 µm) cells (green) can be seen inside a single host cell. C) Mating type a titan cells (>10 µm) are seen in contact with host phagocytes but are too large to be phagocytosed. D) Cells obtained by bronchoalveolar lavage (BAL) were fixed and examined by microscopy for green fluorescence and percent phagocytosis. >500 cells were examined per animal. Error bars indicate SD from four mice per treatment. Asterisk indicates p 0.4 were observed for other pair-wise comparisons. Both macrophages and neutrophils employ oxidative and nitrosative bursts as a means of killing pathogens (Janeway et al., 2008). Titan cell resistance to these stresses was characterized by comparing the growth of purified titan and small cells isolated by cell sorting of BAL samples. Both cell types showed equivalent growth in the absence of oxidative or nitrosative stress ( Figure 6A ). Treatment with sodium nitrate (NaNO3) slowed the growth of the small cell population compared to the titan cell population ( Figure 6B ). Treatment with tert-butyl hydroperoxide (TBHP) resulted in killing of small cells, represented by a decrease in cell counts relative to the initial time point ( Figure 6C ). In contrast, titan cells exhibited continued growth in the presence of these oxidative stresses ( Figure 6C ). Similar results were observed with stabilized hydrogen peroxide treatment. Thus, titan cells are more resistant than normal cells to both oxidative and nitrosative stresses similar to those employed by cells of the host immune system. 10.1371/journal.ppat.1000953.g006 Figure 6 Titan cells are resistant to oxidative and nitrosative stress. Mice were coinfected intranasally with 4×107 cryptococcal cells. At 3 days post-infection, BALs were performed and cells were sorted by FACS based on size. 2×104 titan cells or small cells were resuspended in 100 µL RMPI. Cryptococcal cells received A) no treatment, B) 10 mM NaNO3, C) 1 mM TBHP. At 0, 6, 16, or 24 hours, aliquots of each treatment were plated on YPD agar and colony forming units (CFU) were determined. Error bars indicate SD from three replicates. Titan cells are polyploid In yeasts, cell enlargement is often associated with either cell cycle arrest or increased DNA content [37], [38], [39]. Pheromone sensing in the model yeasts Schizosaccharomyces pombe and Saccharomyces cerevisiae is known to trigger a cell cycle arrest. We examined titan cells for their progression through the cell cycle by characterizing their ability to bud and produce daughter cells. In addition, we determined the DNA content of titan cells. Titan cells produced in vivo were obtained from BAL of mice with single or coinfections. The cells were immediately fixed and stained with DAPI. Microscopic examination of titan cells revealed a single nucleus ( Figure 7A ). Analysis of titan cell nuclear structure by confocal microscopy and z-stack sectioning showed the nucleus had an elongated tubular shape instead of the classic round shape observed in smaller cells (data not shown). Because of its elongated shape, only a portion of the nucleus was observed in each focal plane. Several stages of the cell cycle were identified. In early bud formation ( Figure 7B ), titan cells had a nucleus in the mother cell while the daughter cell lacked a nucleus. The mother cell nucleus was observed at the bud site and entering the daughter cell ( Figure 7C ). After nuclear division the mother and daughter each contained single nuclei ( Figure 7D ). Finally, after cytokinesis was complete, individual nuclei were visible in the mother and associated daughter cell ( Figure 7E ). Budding of the titan cells was readily observed from in vivo samples suggesting complete cell cycle arrest would not explain the increased titan cell size. 10.1371/journal.ppat.1000953.g007 Figure 7 Titan cells can undergo cell division. Mice were infected with 5×107 cells by inhalation of an approximate 1∶1 ratio of a:α cells. At 3 days post-infection, mice were sacrificed, BALs performed, and the resulting cells were fixed and DAPI stained for nuclear content. A) Titan cell containing a single nucleus. B) Titan cell early bud formation. C) Nuclear transfer from a mother (titan cell) to a daughter cell. D) Titan cell late bud formation. E) Cytokinesis of a daughter cell from a titan cell. Bar = 10 µm. Increases in cell size in plants, or gigantism, is often correlated with increased ploidy [40]. Because titan cells contain only one nucleus, we quantified their DNA content by flow cytometry and quantitative PCR. Fluorescently labeled cells from individual or coinfections were isolated by BAL and immediately fixed and stained with DAPI. The fixed cell suspensions were then analyzed using an imaging flow cytometer to define cell populations ( Figure S3 ). Two distinct populations of fluorescent cryptococcal cells were identified: cryptococcal cells alone and cryptococcal cells inside host cells. Because phagocytosed cryptococcal cell size cannot be accurately measured with flow cytometry, only single non-phagocytosed yeast cells were examined further. The single non-phagocytosed yeast cells were then divided into three populations based on cell diameter: ≤10 µm, >10 µm but ≤20 µm, and >20 µm. The ≤10 µm cell population was designated as small cells of typical size for Cryptococcus. The group of cells >20 µm were designated as the titan cell population. The intermediate cell population, >10 µm but ≤20 µm, contained a mixture of small and titan cells, thus could not be accurately characterized by flow cytometry. Flow cytometry and cell sorting of 50,000 cells were used to obtain an accurate representation of the DNA content for each population ( Figure 8 ). DNA content determinations were based on DAPI fluorescence in haploid cells grown in vitro in Dulbecco's modified eagle medium (DMEM) at 37°C and 5% CO2 (non-titan-inducing conditions) ( Figure S4 ). The small cell population isolated from coinfected mice showed a prominent peak consistent with a majority of the cells in the population containing two copies (2C) of DNA. These data would suggest that most of the small cell population in vivo were in G2 of the cell cycle ( Figure 8A ). In contrast, the titan cell population showed two peaks consistent with 4C or 8C DNA content ( Figure 8A ). No differences in titan cell DNA content were observed between the two mating types or in individual versus coinfections, indicating that titan cell DNA content was not altered by coinfection ( Figure S4 ). 10.1371/journal.ppat.1000953.g008 Figure 8 Titan cells have increased DNA content. Mice were intranasally infected with 5×107 cells with an approximate 1∶1 ratio of a:α cells labeled with AlexaFluor 488 (green). At 3 days post-infection, mice were sacrificed, BALs performed, and the resulting cells were fixed. A) Cells were stained with DAPI to measure nuclear content by flow cytometry. Left panel indicates small (≤10 µm) and titan (>20 µm) cell population gates. Right panel indicates DNA content based on DAPI fluorescence for small (dark gray) and titan (white) populations normalized to cell number (% maximum). Dashed lines indicate predicted 1C, 2C, 4C, and 8C DNA content based on DAPI intensity of the 1C and 2C control cells stained and analyzed in the same experiment. B) Cells were grown in vitro in spent DMEM liquid medium for 7 days at 30°C. Cells were fixed and stained with DAPI. Left panel indicates small (≤10 µm), intermediate (>10, ≤20 µm), and titan (>20 µm) cell population gates. Right panel indicates DNA content based on DAPI fluorescence for small (dark gray), intermediate (light gray), and titan (white) populations normalized to cell number. Dashed lines indicate predicted 1C, 2C, 4C, and 8C DNA content based on DAPI intensity of the 1C and 2C control cells strained and analyzed in the same experiment. C) Fixed BAL samples were sorted into small and titan cell populations by fluorescence activated cell sorting. DNA was purified from the sorted populations, normalized to cell number, and chitin synthase 1 (CHS1) gene copy number was determined by comparison to a log phase control sample with a known ratio of 1C:2C cells with a total gene copy number equivalent to 1.4. Analysis of the in vivo samples suggested that both the small cells and titan cells could be undergoing active cell growth and replication, making characterization of titan cell ploidy difficult in these in vivo samples. To determine the ploidy of titan cells, we identified in vitro conditions that stimulated titan cell production. Titan cell formation was only observed in cryptococcal samples grown in spent media previously used to culture mammalian cells ( Figure S5 ). Differences in titan cell formation were observed based on the media used, the temperature of incubation, and mammalian cell type. Optimal in vitro titan cell production was observed when cryptococcal cells were grown in spent DMEM derived from MH-S alveolar macrophages at 30°C. When grown to stationary phase for 5 days in this medium approximately 4% of the total population was titan cells. On average, titan cells generated in vitro were smaller than those observed in vivo, ranging from 15 µm to 30 µm in diameter. Due to the smaller size of the in vitro titan cells, the intermediate cell population (>10 µm but ≤20 µm) was included in the flow cytometric DNA content analysis ( Figure 8B ). In contrast to the in vivo samples, the DNA content of the in vitro small cell population at 5 days was consistent with 1C cells, suggesting that the cells were in stationary phase ( Figure 8B ). Cells grown to stationary phase in a standard growth medium were also 1C (data not shown). The intermediate cell population had a single peak consistent with 4C cells and the larger titan cell population (>20 µm) had a single peak consistent with 8C cells ( Figure 8B ). Thus, titan cells in stationary phase appeared to be either tetraploid or octoploid based on cell size. Quantitative PCR was used to determine the average copy number per cell of the chitin synthase 1 (CHS1) gene as an additional molecular characterization of DNA content in the in vivo small and titan cell populations. Quantitative PCR was performed on the isolated DNA from three cell populations (small, titan, control). This quantitative PCR analysis confirmed that the titan cells had increased CHS1 DNA content compared with the small cells (p 3 times in sterile PBS to remove unbound dye. The cells were resuspended in PBS at a concentration of 1×108 based on hemocytometer count. Three mice per treatment (a, α, or coinfection) were infected intranasally with 5×106 fungal cells. The concentration of yeast cells in the inoculum was confirmed by plating serial dilutions and enumerating CFU and the proportion of a cells in the coinfection inoculum was determined by mating assay [27]. Infected mice were sacrificed at 1, 2, or 3 days post-infection by CO2 inhalation. Lungs were extracted and fixed as described above and unstained sections were examined for cell size, morphology, and fluorescence. Data presented are representative of three independent experiments with two or three mice per treatment per experiment. ste3aΔ mutant strains Two independent ste3aΔ mutant strains were generated by gene disruption as previously described [73].The nourseothricin transgene (NAT) was used to replace the STE3 a gene coding region. PCR was used to generate the 5′ (KN0035 and KN0036) and 3′ (KN0037 and KN0040) flanking regions containing linkers to a NATr cassette and overlap PCR generated the NAT insertion allele ( Table 1 ). The mutant allele was introduced by biolistic transformation into KN99a to generate ste3 a Δ#1 and into the spontaneous ura- strain JF99a [74] to generate ste3 a Δ#2. Transformed colonies resistant to nourseothricin (100 µg/ml) were identified by PCR amplification and sequencing of PCR products spanning a region upstream of the 5′ flanking region into the NAT cassette (KN0079 and KN0031) and from the NAT cassette to downstream of the 3′ flanking region (KN0032 and KN0109). Gene deletion was further confirmed by mating the mutant strains with KN99α on V8, pH 5 media for >14 days at 25°C in the dark. The mutant strains were sterile. The ste3 a Δ#2 was passaged on SD-ura media to isolate a URA+ revertant for use in virulence tests. 10.1371/journal.ppat.1000953.t001 Table 1 PCR Primers. Primer Designation Sequence STE3a Knockout construct KN0035 GCCCTAGCAATGTCGATACCC KN0036 AGCTCACATCCTCGCAGC GCACGTCCGGAGTACACG KN0032 GCTGCGAGGATGTGAGCT KN0031 GGTTTATCTGTATTAACACGG KN0037 CCGTGTTAATACAGATAAACCCTGTATGGCGCTCCTTGGAAG KN0040 CACAGCAAAGGCACATTCGCAAG Outside PCR Checks KN0079 GGAGTTGACGCACGTTTATGGCAA KN0109 CACTGGTGGAGCATTCATGTCG qPCR with primers for CHS1 KN104 GTCCCAGGAGGACTCCTTTC KN105 TGTCGTTCAGGTCGAGTGAG In vivo analysis of ste3aΔ strains Groups of 5–10 mice were infected with 5×104 cells in an approximate 1∶1 ratio of ste3 a Δ#1:KN99α, ste3 a Δ#2:KN99α, or KN99a:KN99αNAT. The actual proportion of a cells in the infecting inoculum was determined by growth on selective media. At 21 days post-infection, animals were sacrificed. The lungs, spleen, and brain from each animal were homogenized in 2–4 ml PBS and serial dilutions were plated on YPD for CFU enumeration. >500 colonies per organ were isolated and assayed for antibiotic resistance on YPD containing 100 µg/ml nourseothricin to determine mating type. Interactions with the blood-brain barrier (BBB) KN99a and KN99α cells were fluorescently labeled as described above. Three mice per treatment were inoculated by tail vein injection with KN99a, KN99α, or an approximate 1∶1 ratio of KN99a:KN99α at a final concentration of 2×107 cells. At 1 day post-infection animals were sacrificed, perfused with 20 ml PBS then 20 ml 4% paraformaldehyde (PFA). Brains were harvested, placed in 4% PFA then 40% w/v sucrose solution in PBS, frozen in isopentane and liquid nitrogen, stored at −80°C, and 50 µm sections were generated. For immunohybridizations, slides were washed in PBS for 15 min followed by incubation with 100 µl trypsin-EDTA (Invitrogen) at 37°C for 10 minutes. Slides were then washed in PBS containing 20% fetal calf serum (Invitrogen) for 10 minutes, blocked with PBS containing 20% FCS, 0.1% bovine serum albumin (BSA) and 0.1% triton X-100 (Sigma, St. Louis, MO) for 20 minutes, then washed with PBS containing 0.1% triton X-100. Anti-collagen IV antibody (Santa Cruz Biotechnology, Santa Cruz, CA) was diluted to a 1/50 concentration in PBS with 0.1% BSA and 0.1% Triton X-100. Antibody-treated slides were incubated overnight at 4°C followed by washing in PBS. Cy3 labeled goat anti-rabbit antibody was diluted to a 1/200 concentration and added to the slides. After 5 hours of incubation at 37°C, slides were washed three times in PBS for 15 minutes. Hoechst medium was diluted to a 1/500 concentration and added to the slides for 30 seconds. Slides were washed for 5 minutes in PBS and mounted in Vectashield mounting medium. Capsule antigen staining was as described in Charlier et al., 2005 using the CRND-8 and E1 antibodies. Slides were imaged by fluorescence microscopy (Zeiss Axioplan) or by 2-photon confocal microscopy (Zeiss LSM 510 equipped with a Coherent Mira 900 tunable laser) with sections compiled as a projection or as a 3D rendering. Bronchoalveolar lavage (BAL) Four mice per treatment were infected as described above with 5×106 AlexaFluor 488 labeled KN99a, KN99α, and ste3 a Δ#1, or an approximate 1∶1 ratio of one stained and one unstained strain. Infected mice were sacrificed at 3 days post-infection by CO2 inhalation. Lungs were lavaged with 1.5 mL sterile PBS three times using a 20 gauge needle placed in the trachea. For flow cytometry, cells in the lavage fluid were pelleted at 16,000 g, resuspended in 3.7% formaldehyde, and incubated at room temperature for 30 minutes. Cells were then washed once with PBS, resuspended in PBS containing 300 ng/ml 4′,6-diamidino-2-phenylindole (DAPI) (Invitrogen), incubated at room temperature for 10 minutes, washed with PBS, and resuspended in PBS. >500 cells per animal were analyzed for size and fluorescence by microscopy (AxioImager, Carl Zeiss, Inc). Confocal microscopy (LSM710, Carl Zeiss, Inc) and z-stack imaging (AxioImager with Apotome, Carl Zeiss, Inc) were used to examine interactions with host mononuclear cells. Images were analyzed using Axiovision and Zen software (Carl Zeiss, Inc). Crescent shaped and other fluorescently-labeled cryptococcal cell fragments (i.e. not round cells) were observed within host mononuclear cells. These cell fragments were not included in the analysis. Nitrosative and oxidative stress assays Twelve mice were intranasally infected with 2×107 cells in 50 µL PBS of an approximately 1∶1 ratio of KN99a and KN99α cells. At 3 days post-infection, mice were sacrificed by CO2 inhalation and BALs were performed. Cells were sorted by FACS using an iCyt Reflection cell sorter (iCyt, Champaign, IL). Cells were sorted based on size using forward scatter (FSC) into small cell and titan cell populations. Purity of samples was checked by flow cytometry and microscopy. Samples were resuspended in Roswell Park Memorial Institute (RPMI) medium 1640 (Invitrogen) supplemented with 10% fetal bovine serum (FBS) (ATCC, Manassas, VA), 4.5 g glucose/L (BD), 1 mM sodium pyruvate (Invitrogen), 0.01 M HEPES (MP Biomedicals, Solon, OH), 5% penicillin/streptomycin (Invitrogen) and 0.05 mM β-mercaptoethanol (Chemicon) to a concentration of 2×104 cells per 100 µL. Samples were then treated with 10 mM NaNO3 (Sigma-Aldrich, St Louis, MO), 3 mM H2O2 (Walgreens Co., Deerfield, IL), or 1 mM tert-butyl hydroperoxide (TBHP) (Sigma-Aldrich). At 0, 6, 16 or 24 hours post treatment, 10 µL aliquots of each sample were plated onto YPD agar for CFU enumeration. Flow cytometry Fixed BAL samples from 4 mice per treatment were generated as described above and analyzed using an ImageStream imaging flow cytometer and INSPIRE software (Amnis Corporation, Seattle Washington). Briefly, images for 5000 cells per sample were collected and analyzed for single cells (R1), doublets (R0), or aggregates of cells ( Figure S3A ). Only single cells (R1) were used in our analyses because cell aggregates would misrepresent cell sizes. Single cells were further analyzed for AlexaFluor 488 fluorescence and DAPI staining ( Figure S3B ). Due to the high nuclear content of mammalian cells, these cells had extremely high DAPI staining (R2 and R3). Non-phagocytosed yeast cells (R5) we identified based on their low DAPI staining. Visual confirmation of cell size in the flow cytometry images was used to identify small and titan cell populations (R6 and R7), that each gate contained only the target cells, and that no contamination between the populations was observed ( Figure S3C ). Data analysis and gating was performed using IDEAS software (Amnis Corporation). Cryptococcal cells grown in vitro in YPD or DMEM to log or stationary phase were used as controls to identify haploid cells (1C) and actively dividing cells (1C + 2C). To examine titan cell ploidy, fixed BAL samples from 4 mice per treatment were generated as described above. In vitro control samples were grown in YPD or Dulbecco's modified eagle medium (DMEM, 37°C, supplemented with 10% fetal bovine serum (FBS) (ATCC), 4.5 g glucose/L (BD), 1 M sodium pyruvate (Invitrogen), 0.01 M HEPES (MP Biomedicals, Solon, OH), 5% penicillin/streptomycin (Invitrogen) and 0.05 mM β-mercaptoethanol (Chemicon) for 6 hours (log phase) or 5 days (stationary phase). Spent DMEM or RPMI was collected from MH-S macrophages after 3–5 days culture at 37°C and 5% CO2. Spent endothelial cell (EC) media (complete EGM medium, Clonetics, San Diego, CA, USA) was collected from human umbilical vein endothelial cells (HUVEC) after 3–5 day culture at 37°C and 5% CO2. In vitro titan cells were grown in filter sterilized spent media at 30°C or 37°C for 7 days. In vitro and in vivo samples were fixed in 3.7% formaldehyde and stained with 300 ng/ml DAPI in PBS. Autofluorescence of non-DAPI stained fixed titan cells was measured and used to set the baseline for ploidy measurements. Cells were examined for cell size by forward scatter (FCS) and nuclear content by DAPI using an LSRII flow cytometer with FACSDiva software (BD) using gating defined by imaging flow cytometry. FCS cell sizes in each gate were verified by microscopy (Zeiss Axioplan). Data presented are representative of three independent experiments with four mice per treatment. 50,000 cells per treatment were analyzed to determine titan cell formation in vitro. In vitro titan cell formation was variable from experiment to experiment but trends between treatments remained constant. Data presented are representative of five independent experiments. Because the absolute number of cells in each population and in each mouse differed, the DAPI fluorescence for each population was normalized to the number of cells in that population in order to clearly visualize peaks on a histogram representation of the data ( Figure 8 , Figure S4 ). Cells were examined for cell size by forward scatter (FCS) and nuclear content by DAPI using an LSRII flow cytometer with FACSDiva software (BD) using the gating defined by imaging flow cytometry. FCS cell sizes in each gate were verified by microscopy to identify the ≤10 µm, >10 µm but ≤20 µm, and >20 µm cell populations (Zeiss AxioImager). Data presented are representative of three independent experiments with four mice per treatment. 50,000 cells per treatment were analyzed to determine titan cell formation in vitro. In vitro titan cell formation was variable from experiment to experiment but trends between treatments remained constant. Data presented are representative of five independent experiments. Cell sorting and qPCR Ten to fourteen mice were infected with 5×106 AlexaFluor 488-stained cells at an approximate 1∶1 ratio of KN99a:KN99α, as described above. Infected mice were sacrificed at 3 days post-infection and BALs were performed. BALs were pelleted and resuspended in 0.05% SDS in sterile water for 1 minute to promote host cell lysis. Cells were then fixed in 1 ml PBS containing 1% formaldehyde and incubated for 30 minutes at room temperature with mixing. Samples were incubated in 125 mM glycine for 5 minutes, centrifuged at 1500 g for 10 minutes, and the pellets were resuspended in ice cold TBS (20 mM Tris, pH 7.6, 150 mM NaCl) containing 125 mM glycine. Cells were washed once in TBS, resuspended in 1 ml PBS and the cell concentration was determined by hemocytometer count. Cell numbers were adjusted to 106 cells/ml, and 1% BSA was added to the fixed cell suspension. Cells were sorted using a FACSAria fluorescence activated cell sorter (FACS) using FACSDiva software (BD). Small and titan cell populations were isolated by FACS using gating as described above. DNA was isolated from 106 cells from small, titan, and 37°C DMEM (control) cell populations. A portion of the control cell population was DAPI stained and the number of haploid and diploid cells in the population was determined by flow cytometry ( Figure S3 ). Small cells were classified as ≤10 µm and titan cells were >10 µm. After sorting, the two cell populations were pelleted and resuspended in lysis buffer (50 mM HEPES, 140 mM NaCl, 1% Triton X-100, 0.1% Sodium deoxycholate, 1 mM EDTA). The cell suspensions were transferred to tubes containing 0.3 mm glass beads and vortexed for six 5 minute cycles at 4°C. The bottoms of the tubes were then pierced with a hot 21-gauge needle. The tubes were placed into 15 ml conical tubes and centrifuged at 1500 g for 5 minutes at 4°C. The pellets and supernatants were combined and transferred to new tubes. These mixtures were centrifuged for 10 minutes at 10,000 g at 4°C and the supernatants transferred to clean tubes. After a further 5 minute centrifugation, the DNA crosslinks were reversed by adding 200 µl TE (10 mM Tris, pH 7.5, 1 mM EDTA) containing 1% SDS to the clarified supernatants and incubating for 6 hours at 65°C. Samples were then incubated 2 hours at 37°C with 250 µl TE containing 0.4 mg/ml proteinase K. After adding 55 µl 4 M LiCl, the DNA was extracted with 0.5 ml phenol and the DNA was precipitated with 100% ethanol. The DNA pellets were washed with 70% ethanol, dried, and resuspended in TE containing 1.5 µl RNase (Ambion AM22886). Samples were stored at −20°C until analyzed by qPCR with primers KN104 and KN105 for chitin synthase (CHS1) ( Table 1 ). Gene copy number in the control sample was calculated based on the known number of 1C and 2C cells present in that sample (1.4C) based on flow cytometry. The small and titan cell gene copy numbers were normalized to the control sample. Statistical analysis All analyses were performed using Analyse-It (Analyse-it Ltd., Leeds, England). Wilcoxon rank sum analysis was used to analyze differences in coinfection data and P-values 10 µm in diameter) was determined by microscopic examination of >500 cells per sample per mouse. Sufficient cell numbers were unavailable in tissue sections from 1 dpi lungs and 1, 3, 7, and 14 dpi spleen and brain for quantification. Error bars indicate SD from six mice per time point. (2.29 MB TIF) Click here for additional data file. Figure S2 ste3aΔ survival assays. Mice were inoculated with 5×104 cells of either wild-type a, ste3aΔ#1 (left) or ste3aΔ#2 (right) cells and progression to morbidity was monitored. (2.43 MB TIF) Click here for additional data file. Figure S3 Imaging Flow Cytometry. C. neoformans a and α strains were combined with AlexaFluor 488 (green) and incubated at 25°C for 20 minutes. Cells were washed with sterile PBS to remove excess dye. Mice were inoculated with an approximate 1∶1 ratio of a:α cells. At 3 days post-infection animals were sacrificed and BALs were performed. The resulting cells were fixed, DAPI stained, and analyzed using an ImageStream flow cytometer using IDEAS software (Amnis Corporation). A) Cells were first examined for single cells (R1). Aggregates and doublets were excluded from further analysis. B) The R1 population was analyzed for DAPI intensity (X-axis) and AlexaFluor 488 intensity (Y-axis). DAPIhi host cells and phagocytosed cryptococcal cells (R2 and R3) as well as unstained yeast cells (R4) were excluded from further analysis. C) Diameter was used to divide the remaining population, R5, into cells 10 µm (R7). Samples from four mice per treatment were analyzed and gates determined by consensus among the samples. (8.29 MB TIF) Click here for additional data file. Figure S4 C. neoformans titan cells are polyploid. a and α strains were combined with AlexaFluor 488 (green) and incubated at 25°C for 20 minutes. Cells were washed with sterile PBS to remove excess dye. Mice were inoculated with an approximate 1∶1 ratio of a:α cells. At 3 days post-infection animals were sacrificed and BALs were performed. The resulting cells were fixed, DAPI stained and analyzed using an LSRII flow cytometer using FACSDiva software (BD). Fluorescently labeled yeast cells were first identified as 488hi and DAPIlow (left). Forward scatter (FSC) was used to identify small (≤10 µm) and titan (>20 µm) cells. Small (blue line) and titan (red line) cell populations were analyzed for DNA content (DAPI) and normalized for cell number (right). A–D coinfections E–G individual infections H) C. neoformans cells were grown for 5 days at 37°C and 5% CO2 in DMEM, fixed and DAPI stained. Both 1C and 2C peaks can be seen in this cell population. Absolute levels of DAPI intensity in these control cells varied from experiment to experiment thus were included as internal controls for every experiment. (2.31 MB TIF) Click here for additional data file. Figure S5 In vitro titan cell production. Cryptococcal cells were grown in spent DMEM (MH-S alveolar macrophages), RPMI (MH-S alveolar macrophages), or endothelial cell media (human umbilical vein endothelial cells, HUVEC) at 30°C or 37°C. Samples were fixed in 3.7% formaldehyde and 50,000 cells per sample were analyzed for cell size (forward scatter). Data presented are representative of five independent experiments. (2.12 MB TIF) Click here for additional data file.

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Changing epidemiology of systemic fungal infections.

M. Richardson, C Lass-Flörl (2008)

Species of Candida and Aspergillus remain the most common causes of invasive fungal infections, but other yeasts and filamentous fungi are emerging as significant pathogens. Opportunistic yeast-like fungi and moulds such as Zygomycetes, Fusarium spp. and Scedosporium spp. are increasingly being recognised in patient groups such as those with leukaemia and in bone marrow transplant recipients. Recognition of these epidemiological changes is critical to patient care. The key elements in selecting an appropriate antifungal agent are the type of patient (solid-organ or stem-cell transplant), severity of immunosuppression, history of prolonged exposure to antifungal drugs, and knowledge of the genera and species of the infecting pathogen and its typical susceptibility pattern.

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Fungal Cell Gigantism during Mammalian Infection

Oscar Zaragoza, Rocío García-Rodas, Joshua D. Nosanchuk … (2010)

Introduction The interaction between a microbe and a host involves a complex response by both the pathogen and the infected individual. The host has multiple defence mechanisms to avoid infection, damage and disease. Microbial pathogens adapt to survive in a host through multiple changes that include signalling pathways that confer the capacity to survive immune-mediated stresses. Both entities, the host and the microbe, interact and each contributes to the outcome of infection [1]. In the case of fungal pathogens, the interaction with the host frequently results in morphological changes. For example, Candida albicans forms pseudohyphae and true hyphae during infection, phenomena associated with virulence [2], [3], [4]. Other examples of fungal pathogens that form filaments during infection are Aspergillus species and the agents of zygomycosis. In contrast, Histoplasma capsulatum and Blastomyces dermatitidis manifest a temperature regulated dimorphism, such that at ambient temperatures they form filaments and at 37°C transform into yeast cells [5], [6], [7]. Although the role of these morphological transitions is not completely understood, it is believed that the phenomenon of fungal dimorphism plays an important function during the interaction of each of these microbes with their host. The fungus Cryptococcus neoformans is the causative agent of cryptococcosis, a disease responsible for over 600,000 deaths per year, which makes this pathogen a major global threat. Cryptococcosis is currently the fourth leading cause of death from infectious diseases in Sub-Saharan Africa [8]. C. neoformans is unique among the major fungal pathogens in that it possesses a polysaccharide capsule surrounding a yeast cell body [9]. Capsular polysaccharides are also released into host tissues [10], [11], [12], where they mediate numerous deleterious effects on host immune function [13], [14], [15]. In fact, the polysaccharide capsule is the factor that makes the greatest contribution to the virulence of C. neoformans [16]. Although C. neoformans can form pseudohyphae during mating [9], this pathogen is mainly found in host tissues as round yeast cells. However, there is a specific morphological change associated with cryptococcal infection that involves a significant increase in capsule volume. Capsule size in C. neoformans depends on the growth condition (reviewed in [17]). While capsule size is relatively small in standard laboratory media and in the environment, it undergoes a large increase in capsule size during pulmonary infection [18], such that it can comprise more than 90% of the total volume of the cell [19]. Capsular enlargement is believed to confer an advantage to the microorganism during its interaction with the host. For example, capsule growth interferes with complement-mediated phagocytosis [20] and protects the yeast cell against free radicals and antimicrobial agents [21]. Furthermore, increased capsule size makes the yeast more difficult to phagocytose by a variety of phagocytic cells, including amoebas that can prey upon C. neoformans in the environment [21]. We now report another morphological change whereby gigantic fungal cells are formed in tissue. This change is achieved, not only by a significant increase in capsule size, but also by an enlargement of the cell body. During pulmonary infection, we observed that a significant proportion of yeasts in the lung had cell volumes 900-fold larger than cells grown in standard laboratory conditions. In retrospect, giant cells have been noted in prior studies [18], [22], [23], [24], [25], but were never isolated or studied. The emergence of fungal giant cells poses a formidable problem for the immune system. In this study we present experimental evidence suggesting that C. neoformans gigantism may be a strategy that confers upon the organism the ability to survive within the host for long time periods. Results C. neoformans produce gigantic cells during infection While the typical size of C. neoformans cells ranges between 4–8 microns (Figure 1A), we confirm the existence and report the recovery of C. neoformans cells of enormous size formed during infection (Figure 1B). Although these cells manifested a large increase in capsule size, there was also a concomitant increase in cell body size. The cell body reached 25–30 µm in diameter, which was almost 7-fold greater than the 4.5 µm average size observed in vitro. This effect was more dramatic if the size of the capsule was included, with the giant cell size typically ranging from 40 to 60 µm in diameter, although extremely large cells with diameters around 70–100 µm were occasionally observed. If one considers volume applying the formula for a sphere (V = 4/3×π×(r3)), then giant cell formation involved an increase of 900-fold in cellular volume, compared to cells grown in Sabouraud medium. We investigated whether the phenomenon was found in different cryptococcal strains. Consequently, we infected different individual mice with ten different C. neoformans strains (both serotype A and D, including standard strains, such as H99, 24067 or B3501, and clinical isolates from the Yeast Collection of the Spanish Mycology Reference Laboratory). For all strains, we found giant cells after three weeks of infection, indicating that this phenomenon applied to diverse strains (results not shown). Hence, we focused our efforts on the model serotype A strain H99, and we arbitrarily defined giant cells as those with a cell diameter greater than 30 µm (capsule included), a size that is 5–6 times the usual size observed in vitro, and is virtually never encountered during in vitro experimental conditions. Using this strain, we observed giant cell formation in four different mouse strains (CD1, BALB/c, C57BL/6J and CBA/J), indicating that the emergence of giant cells was also not mouse strain specific. 10.1371/journal.ppat.1000945.g001 Figure 1 Morphological features of giant cells from infected mice. A) Cells grown in vitro in Sabouraud medium. B) Cell obtained from the lungs of a mouse infected with C. neoformans (105 cells/mouse) 5 weeks earlier. C) Photomicrographs of a giant cell illustrating the presence of multiple vesicles (highlighted with arrows) within giant cells. Scale bar, 10 µm. D, E and F, vacuole staining with MDY-64 dye. Yeast cells isolated from the lung of mice four weeks after infection with 105 yeast cells were stained with the specific vacuole marker MDY-64 as described in Materials and Methods. Localization of the signal was performed by confocal microscopy. D) Cell of regular size. E and F) Giant cells. In all the cases, regular light microscopy, fluorescence and the merge of both images are shown. Scale bars in the left panels applies to the corresponding middle and right panels. G, H and I, transmission electron microscopy of the capsule and cell wall of regular and giant cells. Cells grown in vitro or giant cells obtained from the lungs of mice three weeks after infection were fixed and processed for TEM. G, cell grown in vitro; H, giant cell; I, magnification of the cell wall region of the giant cell shown in B. CP, capsule; CW, cell wall; CY, cytoplasm. The rectangle indicates the width of the cell wall. J,K, staining of giant cells with anti-melanin mAb. J, light microscopy; K, fluorescence. Scale bar, 10 µm. Morphological features of the giant cells Giant cells had different cellular features than cells of regular size (Figure 1). Giant cells frequently contained multiple vesicles of unknown function that could reach more than 50 per cell (Figure 1C). In addition, there was usually a single enlarged vesicle that occupied a significant proportion of the cell body volume. To better identify these intracellular structures, we stained the cells with the vacuole specific marker MDY-64. In regular cells, we normally observed the presence of a single vacuole (Figure 1D). In giant cells, we observed two patterns of staining with this specific marker (Figure 1E,F). Multiple vesicles which stained with the vacuole marker were identified in approximately 50% of giant cells, whereas the remainder displayed staining mainly in a single large intracellular vesicle. These results suggest that in some giant cells, the vacuole fragmented into multiple vesicles or the smaller vesicles failed to coalesce. A peculiarity of the giant cells was the abnormally large width of their cell wall. This feature was most apparent when the cells were observed by transmission electron microscopy. Using this technique, we could determine that the cell wall of regular cells had a width between 50–100 nm (Figure 1G). In contrast, the width in giant cells was 20–30 larger, ranging from 2 to 3 µm (Figure 1H,I). In these pictures, it was also apparent that the density of the capsule differed between regular and giant cells. In the case of yeast obtained in vitro, the cells displayed a low density capsule with individual polysaccharide fibers attached to the cell wall (Figure 1G). In giant cells, the capsule was significantly denser in the regions close to the cell wall (Figure 1H,I). Fungal cell suspensions recovered from the lungs of infected mice had a dark brownish colour. We hypothesized that this phenomenon could be due to in vivo pigment accumulation at the cell wall level, in particular melanin [26]. To investigate this hypothesis, we stained giant cells with specific mAbs to melanin [27]. Giant C. neoformans cells bound mAb to melanin at the cell wall level (Figure 1J,K), suggesting that this structure was melanized. In addition, we observed that giant cells showed a high degree of autofluorescence (result not shown), which has been reported in cryptococcal cells grown in certain media [28]. Scanning electron microscopy images suggested that the capsule of giant cells was different from that of cells grown in vitro. For cells grown in standard Sabouraud medium we noted that the dehydration and fixing process resulted in polysaccharide shrinkage and aggregation of fibers such that regions of the cell wall became exposed (Figure 2A). In contrast, the architecture of the capsule of giant cells appeared intact and well preserved, revealing a highly cross-linked polysaccharide net (Figure 2B) that accumulated around the cell body as a very compacted layer (Figure 2C). In addition, we frequently observed the presence of “holes” in the capsule (Figure 2 D, E and F), which we interpreted as pathways formed during recent budding. Transmission electron microscopy images confirmed that the capsule of giant cells was denser than the capsule of in vitro cultivated cells (Figure 1 H, I). 10.1371/journal.ppat.1000945.g002 Figure 2 Scanning electron microscopy of cells grown in vitro and of giant cells. A) Cell in vitro grown in Sabouraud. B-F) Giant cells isolated from lung. Scale bar in panel B (10 µm) also applies to panels C-F. The higher degree of cross-linking in the capsule of giant cells relative to in vitro grown cells was confirmed by treating the cells with DMSO or γ-radiation, procedures known to strip the capsule of cells of regular size [29], [30], [31]. γ-radiation removed the majority of capsule of giant cells, but the inner region of the capsule remained attached to the cell (Figure 3). In contrast, DMSO treatment did not affect capsular size of giant cells while it routinely strips the capsule of cells grown in vitro (results not shown). The increased resistance of the inner capsule to radiation and the overall capsule to organic solvents is consistent with a higher degree of capsular cross-linking by the giant cells. 10.1371/journal.ppat.1000945.g003 Figure 3 Capsule release by γ-radiation. Cells of in vitro-enlarged capsule (A) and giant cells (B) were exposed to γ-irradiation, suspended in India ink, and observed under the microscope. Images of representative cells are shown before (control, left panel) and after irradiation (γ-irradiation, right panel). In C. neoformans, complement deposition is affected by the porosity and blocking capacity of the capsule [32]. Consequently, we characterized complement localization in the capsule of giant cells as a measure of capsule penetrability. As shown in Figure 4A, complement is known to deposit in the inner location of the capsule near the cell wall of typical yeast cells [20]. When giant cells were incubated in mouse serum, we observed that complement was not detected in the inner regions of the capsule (Figure 4B). The exclusion of complement from the inner capsule is consistent with reduced permeability resulting from increased fibril cross-linking. 10.1371/journal.ppat.1000945.g004 Figure 4 Differences in capsular structure between regular and giant cells. Yeast cells from lung extracts were incubated in mouse serum and labelled with antibodies to C3 and GXM. A) cell of a regular size; B) giant cell. Light microscopy and complement localization (green fluorescence) and capsule edge (red fluorescence) are shown. C–H) capsular features shown by fluorescence. C–F) Indirect immunofluorescence with GXM-binding mAbs. MAb 18B7 labelling of cells of regular size (C) or giant cells (D–F) isolated from lung of infected mice. Light microscopy and fluorescence pictures are shown for each cell. G–H) Binding of wheat germ agglutinin to C. neoformans. The presence of chitin-like structures in the capsule was studied by the binding of WGA to the cells as described in Materials and Methods. G) Cells grown in Sabouraud and then transferred to 10% Saboraud buffered at pH 7.3 with 50 mM MOPS buffer to induce capsule enlargement. H) A representative giant cell isolated from the lungs of infected mice. Arrows indicate the major regions where WGA binds to the cells. Scale bars in C–D in light microscopy panels denote 5 microns and apply to the fluorescence images. To ascertain whether giant cells manifested antigenic differences from cells grown in vitro we used indirect immunofluorescence with mAb 18B7. We compared cells of different size obtained from the lungs of infected mice, to avoid the possibility that factors of the immune system influenced the antigenic properties of the capsule. When stained with mAbs 18B7, cells of small size exhibited a uniform annular binding pattern (Figure 4C), which was identical to the binding of this mAb to cells grown in vitro [32]. In contrast, most of the fluorescence localized to the edge of the giant cell capsule, and this binding was diffuse and punctate (Figure 4D–F). Moreover, many cells showed a double ring, punctate pattern, with a more uniform inner ring and a rougher, more diffuse outer ring. Chitin-like structures in the capsule were recently demonstrated by the specific binding of fluorescent wheat germ agglutinin, which binds to sialic acids and β-1,4-N-acetylglucosamine (GlcNAc) oligomers [33]. We used WGA to ascertain whether these structures were also present in giant cells. Cells grown in vitro bound WGA, especially at the neck between the mother cell and the bud (Figure 4G). In giant cells, these structures were particularly prominent. Protrusions into the capsule were longer, reaching several microns (Figure 4H). Replication of giant cells Giant cells were viable, since they replicated when placed on fresh agar plates. Daughter cells emerging from giant cells were not trapped inside the thick polysaccharide capsule, but rather traversed it in less than 0.08 seconds (Figure 5A and B, supporting Videos S1 and S2). In some cells, movement through the capsule was much faster, taking less than 0.01 seconds (Figure 5C, supporting Video S3). By measuring the distance travelled through the capsule by emerging buds and the transit time we estimated that the daughter cells traversed the capsule at 20–50 m/h, which is a remarkably high velocity for a microscopic unicellular particle in a gelatinous environment. This data suggested the existence of a motive force propelling and separating the buds from the mother cell's capsule. We observed that giant cells could produce several daughter cells over brief periods of time (2–3 hours), with buds always emerging from the same cell site (data not shown). Despite ejection, the daughter cells remained close to the capsule of the mother cells where they replicated, producing abundant progeny around the giant cells. Even after the giant cells were surrounded by daughter cells, new buds were still ejected with significant force, since they were able to displace and move the surrounding cells upon impact (see supporting Video S4). 10.1371/journal.ppat.1000945.g005 Figure 5 Time lapse images of replicating giant cells in vitro. Giant cells were obtained from lungs of four weeks-infected mice and placed on Sabouraud plates. After 18 h, the growing colonies were monitored under the microscope. The digital videos were processed with Windows Movie Maker software. This software allowed the conversion of the videos into single pictures, each one represented as a different frame of the video. The number in each picture corresponds to the time in seconds. A, B and C correspond to three different budding events. Corresponding videos are included as supporting information files. When placed on agar, we observed that not all giant cells produced daughter cells after 24 h, suggesting that some of these cells were metabolically arrested, or had died prior to or during the isolation procedure. To measure the percentage of replicating cells we obtained giant cells from two mice and counted the proportion of giant cells producing colonies after 24 h on agar with a microscope. The percentage of giant cells reproducing was 60% and 73% for each mouse, respectively, indicating that the majority of giant cells were viable. As a secondary technique for testing giant cell viability we used the method based on the reduction of 2,3-bis(2-methoxy-4-nitro-5-sulfophenyl)-2H-tetrazolium-5-carboxanilide inner salt (XTT) by alive cells. Giant cells manifested a strong capacity to reduce XTT, which was approximately 100-fold greater than the activity shown by the same number of cells grown in vitro (data not shown). This result indicates that giant cells are metabolically active. In vitro cellular growth in minimal media and its relation to cell aging We tried to induce giant cell formation in vitro by incubating the cells in different media. When we incubated the cells in minimal media, around 4–5% of the cells showed a marked increased in cell size over 4 days. These cells reached up to 25–30 µm in diameter (capsule included), approximating, but not quite reaching the size of the giant cells recovered from mouse lungs. In addition, these in vitro giant cells showed other phenotypic differences with the giant cells obtained in vitro, such a smaller capsular size and a lack of enlargement of the cell wall (result not shown). Although this in vitro medium only partially reproduced the gigantism phenomenon, we used it to study if there was any relationship between cellular enlargement and the age of the cells. We hypothesized that massive cellular growth required a prolonged period of time, so the majority of the giant cells obtained would be originated from the initial inocula. To explore this hypothesis, we labelled the cells with complement. Complement proteins, especially C3, bind to the capsule covalently without inhibiting cell growth and do not segregate to buds after replication [34]. Consequently, we labelled cells grown in Sabouraud medium with mouse C3 and then incubated them in minimal medium (which induces a small population of giant-like cells) and Sabouraud medium. At time zero, all the cells were labelled with complement (Figure 6A), but after 4 days of incubation in minimal medium, only a few cells remained labelled (Figure 6B). When we measured the average size of the cells with complement bound after four days of incubation in minimal medium, we found that these cells had a significant larger size than the cells incubated in Sabouraud medium (Figure 6B). We repeated this experiment, placing the cells in parallel in minimal medium, which induces cell enlargement in some cells, and in Sabouraud medium, in which cell enlargement is not expected. Then, we measured cell size of complement-labelled and non-labelled cells. In minimal medium, we observed that the complement-stained cells were significantly larger than the unlabelled population (Figure 6C, D). Large cells were not found in Sabouraud medium after C3 labelling and there was no difference in the size of cells with and without C3 labelling (Figure 6C, D). This result indicates that cellular enlargement and giant cell formation is correlated with the age of the cells, such that the giant cells are the older cells in the culture. 10.1371/journal.ppat.1000945.g006 Figure 6 Giant cell formation in vitro. Cells from H99 strain were grown in Sabouraud, washed and labelled with mouse serum. C3 deposition was then detected by immunofluorescence (A). After incubation in mouse serum, cryptococcal cells were transferred to minimal medium (MM) for four days, and C3 was detected (B). Scale bar in A and B, 10 µm, and apply to the corresponding fluorescence panels. C) Cells were grown and labelled as in A (Day 0 sample), transferred to Sabouraud or MM for 4 days at 37°C, and the cell size was measured in the C3 labelled population (“MM, C3 labelled” and “Sab, C3 labelled”) and in the whole population, including both C3 labelled and unlabelled cells. The average and the standard deviation (error bars) are plotted and p-values for the highlighted comparisons are shown. At least, 20–50 cells were counted, except in the “Sab, C3 labelled” samples, where C3 positive cells were rarely identified due to the overgrowth of the culture. Kruskall-Wallis test was used to assess statistical differences. D) Scatter representation showing all the cells plotted in panel B. The line in each sample denotes the average of the distribution. DNA content We hypothesized that giant cell formation was a consequence of continued cell cycle progression without cellular fission. In other fungi, cell size can be related to DNA content [35], [36]. Using flow cytometry to measure DNA content, we found that giant cells had low permeability to propidium iodide by regular staining protocols (results not shown), but could be permeabilized by heating at 60°C for 45 minutes. As mentioned above, cells from the lungs of mice infected for 3–4 weeks showed strong autofluorescence, so we measured the intensity of the signal in the presence or absence of propidium iodide. We first analysed the difference in the forward scatter (FSC, cell size) and side scatter (SSC, cell complexity). When we compared these parameters, we observed a population of larger yeasts in the cells isolated from the lung that was not present in the yeast cells obtained in vitro, as was expected with the presence of giant cells in vivo (Figure 7A, B). This population was defined as region 1 (R1), which contained the giant cells present in the population. To estimate the DNA content, we added propidium iodide to these samples. When we measured the propidium iodide staining, we found that there was a high variation in the DNA content in the cell population obtained from lungs (Figure 7C), which implied a high variation in the DNA content of cells in vivo. Fungal cells isolated from the lungs of infected mice also displayed significant autofluorescence in the absence of propidium iodide staining. To assess the staining of giant cells, we subtracted the autofluorescence of the cells present in region 1 from the fluorescence value determined in the presence of propidium iodide. The mean fluorescence intensity for the giant cells was almost 103 fold higher than the signal measured in cells grown in vitro (Figure 7D). This result suggested that giant cells contain multiple copies of DNA. To quantify the ploidy level of the giant cells further, we performed real time PCR to amplify the ITS1 region from ribosomal DNA. We included controls of purified genomic DNA of known concentrations, which yielded a lineal relationship between crossing point values and DNA concentration. When we compared 300 giant and regular cells there was a difference in the Ct values of more than 3 cycles (36.95 in giant cells versus 40.00 in regular in vitro grown cells). When we estimated the amount of DNA present in each condition according to a standard curve generated using different concentrations of genomic DNA, we calculated that the amount of DNA in each giant cell was 1.3×10−7 ng. In contrast, in vitro cultivated cells contained 8.4×10−9 ng. This result indicated that giant cells contained 16× DNA than regular cells. This experiment was repeated using different cell concentrations and consistent results were obtained (result not shown). 10.1371/journal.ppat.1000945.g007 Figure 7 Determination of DNA content by cytometry and DAPI staining. Fungal cell samples were obtained from the lungs of one mouse 3 weeks after challenge with 105 C. neoformans cells. As a control, cells grown for 24 hours in Sabouraud were also used. The cells were fixed after incubation at 60°C for 45 minutes. Propidium iodide was immediately added at 10 µg/mL DNA content and the labelling was determined by the propidium iodide fluorescence intensity. Matched samples were subjected to cytometry without propidium iodide. A) FSC/SSC plot of cells grown in Sabouraud. B) FSC/SSC of cryptococcal cells from lungs. Region 1 delimits all the cells whose size and complexity are not found in cultures grown in vitro and presumably cover all the cells with increased cell size. C) Propidium iodide fluorescence intensity of the four samples analyzed: in vivo isolated cells (plus and minus propidium iodide) and in vitro cells (plus and minus propidium iodide). D) Propidium iodide fluorescence intensity of the cells from the lung present in R1. E–F) DAPI staining. Yeast cells obtained from the lungs of mice infected with 105 yeast cells were stained with DAPI. E) cell of regular size; F) giant cell. Corresponding light microscopy and fluorescence images are shown. Scale bar in the light microscopy panels apply to the fluorescence images. DAPI was used to directly observe the nucleus of the giant cells. This staining revealed that both regular (Figure 7E) and giant cells (Figure 7F) contained a single nucleus. Signal transduction pathways involved in giant cell formation We investigated the potential involvement of two of the major signal transduction pathways in C. neoformans (cAMP and Ras1 [37], [38]) in giant cell formation. Ras1-deficient cells produced giant cells in the lungs of infected mice (Figure 8A). In contrast, mutants unable to accumulate cAMP (lacking adenylate cyclase encoded by the CAC1 gene) did not produce giant cells during murine infection (Figure 8A), suggesting that this pathway was required for giant cell formation. 10.1371/journal.ppat.1000945.g008 Figure 8 Giant cell formation depends on cAMP, but not Ras1. C57BL/6J mice were infected with H99, ras1 (Ras1 mutant) and cac1 (adenylate cyclase mutant) strains (106 per mouse). After three weeks, the mice were sacrificed and fungal cells were isolated. Representative pictures of fungal cells are shown. A) India ink microscopy, scale bar in left panel applies to all the pictures. B) Cell size distribution of wild type, cac1 mutant and its reconstituted strain (cac1/CAC1) after 5 days of growth in minimal medium at 37°C. The cells were suspended in India Ink to delimit the capsule, and the diameter of the cells (capsule included) was measured microscopically. C) Forward Scatter/Side Scatter plot of cac1 mutants after incubation in minimal medium. The cells described in B were analysed by flow cytometry to obtain the corresponding forward scatter (FSC, correlated with cell size) and side scatter (SSC, correlated with cell complexity). The absence of giant cells in mice infected with the cAMP mutant was associated with a reduced fungal burden. Hence, the lack of giant cells with this mutant in vivo might be related to the ability of the host to rapidly clear the fungus. For this reason, we examined whether a cac1 mutant formed giant cells in minimal media. The cac1 mutant failed to produce giant cells, whereas a significant proportion of cells of the wild type (H99) and reconstituted (cac1/CAC1) strains manifested cellular enlargement (Figure 8B), confirming that cAMP pathway is involved in giant cell formation. To further characterize this phenotype, we analysed the forward and side scatter profile of the cells from the three strains by flow cytometry, since this type of plot allows for clear differentiations in cell size. As shown in Figure 8C, the cac1 mutant yielded a very homogenous population of relatively small FSC and SSC values. In contrast, the wild type and reconstituted strains produced more heterogeneous populations, in which cells with larger FSC and SSC values were measured, indicating the appearance of cells of larger size. Distribution of fungal cell size during infection and correlation with inflammation The distribution of cell sizes in vivo was extremely variable depending on the experimental conditions. Under our standard conditions (infection of 6–8 weeks old mice with 105 yeast cells) we consistently found that the proportion of giant cells in the lung was between 1–10%, with variation between individual mice and between experiments. However, in several experiments, we occasionally found that the proportion of giant cells was higher than 90% of the lung fungal cell population. Curiously, in those experiments where the proportion of giant cells was very high, there were no obvious signs of disease and the mice looked healthy. We decided to investigate this observation in more detail by studying the relationship between inoculum and giant cell formation. We hypothesized that infections with low inocula could reproduce chronic or latent asymptomatic infections. For this purpose, we performed infections with high (106/mouse) or a low dose (104 cells/mouse). Mice infected with high inocula consistently developed typical cryptococcal disease, as indirectly shown by progressive weight loss (Figure 9A). However, we found a high variation in outcome when the mice were infected with a low inoculum. Most of these mice (2 of 3) developed disease comparable with that seen after infection with a high dose. Severe disease was characterized by dense inflammation in the lungs, increasing the size and weight of these organs (1.3–1.8 grams) such that they accounted for 7–8% of the total body weight (Figure 9B). In contrast, the lung mass of asymptomatic mice (control mice and one of the mice infected with low inocula) was approximately 0.45 grams and ∼1% of total body weight. As expected, mice developing severe disease had a significantly higher number of CFUs (>106/lung) than the mice that did not manifest obvious signs of disease, where the number of CFUs remained very low (<104) during the experiment (Figure 9C). When we recovered the fungal cells from the lungs of these mice, we found profound differences in the size of the yeast cells. The average cryptococcal cell size in mice receiving a high inoculum was around 15–20 µm, which was significantly larger than the size reached when grown in vitro in rich medium (Figure 9D, Sabouraud medium). Although approximately 5–20% of the yeast cells met criteria for giant cells, the enlargement of the majority of cells isolated was mainly due to increase in the capsule size, so the size of these populations was only slightly different from the size reached when capsule size is induced overnight in vitro (see Figure 9D, in vitro enlarged capsule size). We were able to isolate around 1-3×103 C. neoformans cells from the lungs of asymptomatic, low dose infected mice and the average yeast cell size was around 40 µm (Figure 9D). Notably, approximately 70% of the isolated yeast cells were giant forms. 10.1371/journal.ppat.1000945.g009 Figure 9 Parameters of disease in mice infected with C. neoformans and giant cell proportion in the lungs. CD1 mice were infected with different inocula of C. neoformans, and different parameters were monitored during the course of the infection. A) Body weight of mice infected with a low dose (∼104 cells, black line, closed symbols, n = 3) or a high dose (106, grey line, grey symbols, n = 3) of C. neoformans. A control mouse, injected with PBS, is also shown (grey line, open symbols). B) Proportion of the lung weight in respect to body weight, as an indicator of lung inflammation. C) CFUs from the lungs of mice described in A and B. D) Cell size of C. neoformans in vitro and in vivo. Cell size (capsule included) was measured in fungal samples obtained from mice infected with high or low C. neoformans doses (see Figure 9A-C). As a control, cell size was measured in cells grown in vitro in Sabouraud or in 10% Sabouraud buffered at pH 7 with 50 mM MOPS to induce capsule enlargement. The number expresses the proportion of giant cells in each sample, defined as cell with a diameter above 30 µm. The line in each distribution represents the average of the population. E) Results of the CART analysis. Left panel, prediction of the correlation between inflammation and total fungal cell size. A cell size lower than 35.92 µm was strongly associated with increased inflammation. Right panel, corresponding ROC curve, showing a region under the curve of 0.8. We also analysed the proportion of giant cells in the lungs using Classification and Regression Trees (CART) analysis. Using this approach, we found that there was a strong association between the total fungal cell size in the lungs and the degree of inflammation, such that high inflammation was predicted when the average fungal cell size was below 36 µm (Figure 9E). This prediction is in accordance with our initial criteria of defining giant cells as those with a cell diameter greater than 30 µm. When we plotted the corresponding ROC curve, we found that the region under the curve was 0.80, which provides strong support for the prediction. On the other hand, the model could not efficiently predict the proportion of giant cells according to inflammation, due to the low number of yeast cells found in lungs without significant inflammation. The relationship between the proportion of giant cells and inflammation was confirmed histologically. In control mice, the lungs revealed the typical structure in which alveolar spaces were present throughout the lungs (Figure 10). In the infected mice, we only observed this benign histology in the asymptomatic mouse (mouse 1 of the group infected with 104 yeast cells, Figure 10). In the rest of the low dose (Figure 10) and all of the high dose infected mice (result not shown), dense inflammation was observed and alveolar spaces contained numerous yeast cells and inflammatory cells. 10.1371/journal.ppat.1000945.g010 Figure 10 Histological sections of mice infected with different C. neoformans inocula. Tissue sections from mice described in Figure 9 were stained with hematoxylin and eosin. Tissue section of control mouse (upper left panel) and three different mice infected with a low C. neoformans inocula. Mouse 1, upper right panel; mouse 2, middle and lower left panels; mouse 3, middle and lower right panels (for mice 2 and 3, two different regions are shown). For each panel, a magnification of a region is shown, which is delimited by an inset. Scale bar in upper left magnification, 25 µm, and applies to the rest of amplified panels. We performed another experiment using older mice (14–16 weeks old), which are more resistant to infection, and a lower infective dose than in the experiment previously described. We infected with either a low (103 cells/mouse, 10 mice) or a high dose (105 cells/mouse, 3 mice), and measured fungal cell size after a month of infection. In this new model, the mice did not develop any visible sign of disease. When the mice were sacrificed after one month, only one of the mice infected with the high dose showed inflammation in the lungs (table 1, 105, mouse 2). In the group infected with low dose, we did not find yeast cells in 3 mice, indicating that the infection had been cleared. In the other seven mice, we found a low number of yeast cells, suggesting a chronic asymptomatic infection. In six of the seven mice, the average size of the fungal cells was above 30 µm, and the proportion of giant cells was between 50–90%. When the mice were infected with a higher dose, the two mice in which no inflammation was detected showed fungal cell sizes above 30 µm, with a proportion of giant cells around 70–80%. In the mouse with inflammation (mouse number 2), the average fungal cell size was smaller (14 µm), and the proportion of giant cells was less than 5%. This data supports the notion that the highest proportion of giant cells is found in hosts with chronic and longstanding infection. 10.1371/journal.ppat.1000945.t001 Table 1 Fungal cell size and proportion of giant cells in 16-weeks old mice infected with low and high doses of H99 strain. Yeast dose (cells/mouse) 103 105 Mouse number 1 2 3 4 5 6 7 1 2 3 Lung weight (mg) 240 390 326 340 339 240 309 550 860 413 Mean ± St. Dev (µm) 34±6 33±6 31±13 20±5 49±9 46±10 44±11 34±12 14±5 40±10 Giant cells (%) 61 74 50 9 91 80 90 67 4 82 Lower 95% CI 32.3 29.5 21.4 15.5 42.1 30.4 39.4 30.1 13.3 34.1 Upper 95% CI 36.3 37.3 40.5 25 57.6 62.4 49.8 37 16 46.1 Ten CD1 mice (16 weeks old) were infected with 103 yeast cells/mouse, and three mice were infected with 105 yeast cells/mouse. The animals were killed after a month, and the weight of the lungs was measured. Fungal cells were isolated as described in Materials and Methods, and the mean value, standard deviation, proportion of giant cells (above 30 µm), and the lower and upper 95% confidence interval were calculated. In three mice of the group infected with a low dose, we did not find any yeast cells, so the data of the other seven mice is represented. Susceptibility to oxidative stress We measured the susceptibility of giant cells to oxidative stress produced by incubation in H2O2. Giant cells were significantly more resistant to killing by oxidative stress than cryptococcal cells grown in vitro, with survival rates of 19%±10 and 46%±14 for cells grown in vitro and giant cells, respectively (p = 0.014). In vitro interaction of giant cells with phagocytic cells To further characterize the interaction of giant cells with host effector cells, we incubated macrophage-like cells with giant cells and observed the outcome of the interaction using live-imaging microscopy. When small-sized cryptococci were exposed to macrophages, we observed rapid and avid phagocytosis, yeast cell transfer between macrophages, fusion of infected macrophages after division, and intracellular replication of the C. neoformans cells, as described previously [39], [40], [41], [42], [43] and shown in supporting Videos S5, S6 and S7. None of these phenomena were observed when macrophage-like cells were exposed to C. neoformans giant cells, indicating that the interaction between these fungal cells and macrophages was different and had different outcomes. Although in some cases the macrophages seemed to adhere to the giant fungal cells, there was no phagocytosis or macrophage fusion after division (supporting Videos S8 and S9), indicating that macrophages could not cope with the giant cells. Discussion Cryptococcus neoformans giant cells have been occasionally described in the literature primarily as curiosities in histological tissue sections [22], [23], [24], but their importance in pathogenesis has remained obscure. Apart from the fundamental problems in cell biology posed by the mechanisms responsible for the transition to gigantism, we considered that the presence of fungal giant cells would pose a major problem for the immune system simply by virtue of their size. Giant cell formation was associated with several changes to the capsule relative to the typical cells observed in vitro. These changes represented an exaggerated response in capsule, cell body, and cell wall size during infection. Moreover, the resistance to capsule shedding after γ-radiation exposure suggests a more compact and dense structure. Such an increase in capsular compactness could confer a survival advantage in vivo since the capsule is known to protect against oxidative fluxes of the types produced by immune effector cells [21]. Consistent with this idea, we have observed that giant cells are more resistant to oxidative stress. Giant cells maintained their enormous size ex vivo, although they produced smaller cells in agar at replication rates similar to those observed in vitro. Another remarkable aspect of the budding process is the rapidity with which the buds traversed the capsule, especially considering the denseness and compactness of the polysaccharide noted by scanning and transmission electron microscopy. However, we did observe holes in the capsules of giant cells with dimensions that approximated the size needed for daughter cells to emerge. Similar capsule holes have been occasionally described in encapsulated cells grown in vitro [44]. The strong binding of WGA to these cells also suggests the presence of chitin-like structures, which have been proposed to be involved in the movement of the bud through the capsule of the mother cell [33]. Given prior work noting tunnel-like structures formed around buds [33], [34], it is possible that the rapid egress of buds from the mother cells represent movement along such structures that provide a non-obstructed conduit through the capsule. The cellular mechanisms by which cryptococcal cells enlarge to gigantic sizes are not known and a complete understanding of this phenomenon is beyond the scope of the current work. Nevertheless, we explored the potential mechanism of cell division without fission as a way for progressively increasing mass. Cell growth is intimately dependant on the cell cycle. The cells need to reach a critical size for cell cycle to progress, and there is a constant ratio between the mass of the cell and its DNA content (reviewed in [45], [46], [47], [48]). In plants, the phenomenon is striking, because their cells can enlarge in size by 100- or even 1000-fold, and this is achieved by endoreduplication, which is the process in which the cell increases the ploidy of the cells through several rounds of DNA replication [49]. Recently, it has been shown that bacteria from the genus Epulopiscium, which grow to lengths of 200–300 µm and widths of 40–50 µm, have extreme polyploidy, generating tens of thousands of copies of their genome [50]. In a process that may be relevant to cryptococcal gigantism, there are some symbiotic bacteria that undergo an important differentiation process achieved by genome amplification by endoreduplication in plant nodules resulting in significant cell enlargement [51] and provide beautiful examples of how some factors of symbiotic plants regulate the cell cycle of their symbiotic microorganisms. We hypothesized that giant C. neoformans cells achieved their size by repeatedly entering G1 cycles without dividing. To investigate this possibility we stained cells for DNA. Cells recovered from infected animals produced a noisy FACS profile that was interpreted as being consistent with cell-to-cell variation in DNA content. By analyzing cell size and fluorescence intensity we showed that cell size correlated with DNA content, thus establishing that larger cells have more DNA, a result confirmed by real-time PCR. These findings are consistent with a mechanism for DNA replication without cell fission. The proportion of C. neoformans giant cells in infected mouse lung was a function of total microbial burden and pulmonary inflammation. Okagaki et al also found differences in giant/titan cell proportions during infection experiments with MATa and MATα strains, with the proportion of titan cells being higher when mice were co-infected with both mating types (Okagaki et al, see related article in the current PLoS Pathogens issue). These authors have concluded that titan cell formation is induced by the pheromone signalling pathway. Taking our and their results together, we can conclude that gigantism is a morphological response to host environments that impact cAMP and pheromone signalling pathways, which could regulate the cell cycle with the final purpose of generating giant cells during infection. The fact that the survival of the host is not compromised when the proportion of giant cells is high suggests that giant cells can survive in a local environment in the host for protracted periods of time without disseminating in the setting of intact host immunity, a finding in agreement with Okagaki's report. This notion is consistent with reports that a moderate increase in cell size due to capsule enlargement interferes with C. neoformans dissemination from the lung [52], [53]. Various studies have suggested that C. neoformans dissemination is associated with intracellular survival inside macrophages [54], [55], [56], [57], but this model cannot be applied to giant cells since they exceed the size of macrophages. Hence, the increased size of the giant cells is likely to be an impediment for their dissemination as they are simply too large to cross biological barriers and/or transverse capillary diameters. Nevertheless, such cells are viable and capable of producing small sized variants when placed in suitable conditions, such as rich agar. Taken together, our findings suggest that giant cell formation could provide the fungus with a strategy for prolonged survival in a host. Cells could conceivably survive through the life of the host and then return to soils when an animal dies. Alternatively, the giant cells could await permissive host conditions such as in the setting of advanced HIV infection, immunosuppression after organ transplant, or other conditions impairing host immune responses that lead the fungus to proliferate to produce abundant progeny. In this context, it is noteworthy that micro-yeast forms have been described in the lung of infected mice [18], [58], suggesting another form of size polymorphism at the other end of the scale. All these findings indicate that during infection, C. neoformans can display a wide variation in cell sizes, ranging from micro-forms to giant cells, and suggest that each of these morphotypes have different roles in the pathogenesis of persistence and dissemination. We are aware of occasional reports of gigantic cells in other fungal species. For example, giant Candida albicans cells with diameter up to 30 µm have been described [59], [60], and similar large cells have been described for other pathogenic fungi during infection. The arthroconidia of Coccidioides immitis and Coccidioides posadasii swell to form giant spherules (typically 30–150 µm in diameter) during mammalian infection and the spherules produce a large number of endospores derived from the cell membrane, each with a single nucleus [35]. When the spherule is mature, the cell membrane is dissolved and the endospores are released. Another example of fungal giant cells occurs in species from the genus Emmonsia, responsible for adiaspiromycosis in humans. Emmonsia crescent cells reach up to 200–700 µm during infection and these forms are multinucleate. Emmonsia parva forms cells up to 40 µm, and they also contain several nuclei [36]. It is possible that gigantism is a general property of unicellular fungi that is expressed under certain conditions. If this is the case, the reproducibility of giant cell formation during cryptococcal infection provides an excellent experimental system for the study of this phenomenon. In summary, C. neoformans cells can achieve gigantic dimensions during infection and the phenomenon suggests that gigantism may be considered a new form of fungal dimorphism. The occurrence of extraordinarily large cells may enable an adaptation for persistence in certain hosts. The findings for C. neoformans together with the similar reports in other fungi suggest that this may be a general mechanism for fungal survival under certain environments and possibly contribute to persistence during host-pathogen interactions. Materials and Methods Yeast strains and growth media For most experiments, serotype A H99 strain was used [61]. In some experiments the following strains were also used: 24067 (serotype D, ATCC); B3501 (serotype D, [62]); RPC3 (cac1::URA5, [37]); RPC7 (cac1::URA5/CAC1, [37]); LCC1 (ras1::ADE2, [38]) and different clinical isolates from the Yeast Collection of the Spanish Mycology Reference Laboratory (CL2132, CL4860, CL5154, CL5632, CL5707, CL5066 and CL4979). Yeasts were grown in Sabouraud liquid medium at 30°C with moderate shaking (150 r.p.m.). In some cases, the yeast cells were grown in minimal media (29.4 mM KH2PO4, 10 mM MgSO4, 13 mM Glycine, 3 µM thiamine, 15 mM glucose, pH 5.5). For melanization, L-DOPA containing medium was prepared as in [63]. In other experiments, the cells were transfer from the original Sabouraud culture to 10% Sabouraud medium pH 7.3 with 50 mM MOPS buffer, as described in [17], to induce capsule enlargement in vitro. Mouse strains and infection models Six to eight weeks old female BALB/c, C57BL/6J (Jackson Laboratories, Bethesda, MD) and CD1 mice (Charles River Laboratories) were used in this study. In some experiments, older CD1 mice (16 weeks old) were also used. C. neoformans strains were grown at 30°C, washed with sterile PBS, and suspended at specific cell densities. Fifty microliters of the selected yeast cell suspension were injected intratracheally into mice previously anesthetized with a xylazine/ketamine mixture, as described [64]. Histology sections of lung tissues Lungs were excised from mice at different infection times and fixed in formalin for 48 h at room temperature. The tissues were then dehydrated and embedded in paraffin using an STP120 Tissue Processor (Microm International, Walldorf, Germany). Then, 5 µm tissue sections were obtained using a Leica RM2245 microtome and placed on glass slides. Hematoxylin/eosin staining of the tissue sections was performed using standard protocols. Fungal cells isolation from lungs of infected mice Mice were euthanized at different times after infection and the lungs were removed. Lung tissue was then homogenized in 10 mL of PBS with 1 mg/ml collagenase (Roche, Mannheim, Germany). The cell suspension was incubated for 1 h at 37°C with occasional vortex agitation, and washed several times with sterile distilled water. The cells were suspended in sterile distilled water, and immediately placed in fixative for microscopy, in fresh medium for microscopy observation, or in Sabouraud agar at 30°C overnight to observe in vitro budding. Microscopy techniques Cells were viewed with different microscopes. In some experiments, an Olympus AX70 microscope was used and pictures were taken with a digital camera using QCapture Suite V2.46 software for Windows. Alternatively, a Leica DMI3000B connected to a DFC300 digital camera with LAS 3.3.1 software, or a Leica DMI 4000B or a Leica DMRD microscope connected to a Leica DC200 digital camera with IM1000 software were used. To visualize the size of the capsule, the cells were mixed with an India ink suspension. Digital Images were processed with Adobe Photoshop 7.0 software (San Jose, CA). For confocal microscopy, a SP5 confocal microscope (Leica Microsystems) was use. Macrophage-like cell lines and cell culture techniques The macrophage-like cell line RAW264.7 was maintained in DMEM medium supplemented with 10% heat-inactivated fetal bovine serum, 10% NTCT, and 1% of non-essential amino acids at 37°C in the presence of a 5% CO2 atmosphere. Live-imaging of the interaction between macrophages and fungal cells For phagocytosis experiments, 5×104 macrophages were placed on 96-well plates and incubated overnight at 37°C in the presence of 5% CO2, so that a total number of 105 macrophages was expected after this incubation given a phagocytic cell replication time of approximately 12 h. Fungal cells were added at a 1∶2 (macrophage:yeast cells) ratio in 200 µL of medium. Yeast cells of regular size were obtained by growing in Sabouraud medium overnight. To isolate giant cells, fungal cells were isolated from the lungs of infected mice as described above. Giant cells were separated from the rest of the fungal population by passing the sample through 11 µm filters. Although we defined giant cells as those larger than 30 µm (capsule included), we observed that the capsule did not contribute to retention on the filters, since the size of the cell delimited by the cell wall was the main factor associated with retention or passage. The filters containing the giant cells were incubated in PBS with shaking for 20 minutes, and the cells were concentrated by centrifugation. After filtration, we observed that the population was significantly enriched in giant cells, being more than 90% of the sample. Finally, the cells that transited or were retained by the filter were counted using a haemocytometer. Once regular and giant cells were obtained and exposed to the macrophages, the 96-wells plate was placed under a Leica DMI 4000B microscope using a 20× objective with a 5% CO2 environment and 37°C. Pictures were taken at different time intervals (see figure legend of the corresponding supporting videos). The videos generated by the Leica software were exported as .avi documents and processed with ImageJ (National Institutes of Health, USA, http://rsb.info.nih.gov/ij/index.html) and VidCrop 2.1.0.0 (GeoVid) softwares. The final videos were generated by merging 5 frames per second. Melanin detection by immunofluorescence Suspensions of cells isolated from mouse lung were air-dried on poly-L-lysine-coated slides (Sigma). The slides coated with the cells were washed in PBS, incubated in blocking buffer (Pierce, Rockford, IL) for 1 h at 37°C followed by incubation with 10 µg/ml of the IgM melanin-binding monoclonal antibody (mAb) 6D2 for 1 h at 37°C. MAb 6D2 was generated against melanin derived from C. neoformans [27]. After washing, the slides were incubated with a 1∶1000 dilution of tetramethyl rhodamine isothiocyanate (TRITC) -conjugated goat anti-mouse (GAM) IgM (Southern Biotechnologies Associates, Inc; Birmingham, AL) for 1 h at 37°C. The slides were washed, mounted using a 50% glycerol, 50% PBS, and 0.1 M N-propyl gallate solution, and viewed with an Olympus AX70 microscope equipped with fluorescent filters. Negative controls consisted of cells incubated with mAb 5C11, which binds mycobacterial lipoarabinomannan [65], as the primary Ab or with TRITC-labeled Ab alone. Scanning electron microscopy Yeast cells were washed in PBS and suspended in fixing solution (2% p-formaldahyde, 2.5% glutaraldehyde, 0.1 M sodium cacolydate). Cells were then serially dehydrated with ethanol, coated with gold palladium and visualized using a JEOL (Tokyo, Japan) JAM 6400 microscope. Transmission electron microscopy Cells grown in vitro or isolated from the lungs of infected mice (see above) were fixed with 2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer. The cells were treated with osmium tetraoxide and serially dehydrated. The samples were embedded in epoxy resin and ultrathin sections were obtained, stained with uranyl acetate and lead citrate, and observed in a CM12-Phillips transmission electron microscope. DMSO and γ-radiation treatment Yeast cells with enlarged capsule were exposed to varying amounts of γ-radiation from 137Cs to remove layers of the polysaccharide capsule as described [29], [66]. Briefly, giant and in vitro-grown cells were washed three times in PBS to remove shed capsular polysaccharides, suspended in 1 mL of distilled H2O, and irradiated using a Shepherd Mark I Irradiator at the dose rate of 1388 rads/min. For all experiments, cells were irradiated for 40 minutes. Irradiated cells were collected by centrifugation. In other experiments, the fungal cells were suspended in DMSO as described in [29]. The presence of capsule after the treatments (γ-irradiation or DMSO) was visually observed by suspending the cells in India Ink and regular microscopy. Complement labelling and detection Complement (C3; complement protein 3) deposition on the cryptococcal capsule was performed as in [20]. Briefly, C57BL/6J mice were bled from the retro-orbital cavity and serum was obtained by centrifugation. Approximately 2×107 cryptococcal cells were suspended in 700 µL freshly-obtained serum, and incubated at 37°C for 1 h. Cells were extensively washed and suspended in PBS. C3 was then detected using a fluorescein-isothiocyanate (FITC) conjugated GAM C3 antibody (4 µg/mL, Cappel, ICN, Aurora, OH). Yeast washed and not suspended in serum were used as controls. To delineate the capsular edge, mAb 18B7 (10 µg/mL) specific for GXM [67] was added, and detected using a TRITC conjugated GAM IgG1 antibody (10 µg/ml, Southern Biotechnology Associates, Inc). The cells were observed under fluorescent filters with the Olympus AX70 microscope, QCapture Suite V2.46 software for Windows, and Adobe Photoshop 7.0 for Macintosh. Imaging of daughter cell emergence from the giant mother cells Yeast cells were isolated from infected mice as described above, and placed on Sabouraud agar plates for 18 h 30°C. Initially, we tried recording the budding of giant cells by basic microscopy techniques, such as taking pictures every few minutes or seconds. However, the separation of the bud through the capsule of the giant cell was too fast, so we developed a new approach to record the phenomenon. The surface agar plate was observed with an Olympus AX70 microscope to visualize and continuously record giant cells. To record real-time daughter cell emergence, the cells were observed in the computer screen with the “Preview” option, and the image of the screen was recorded with a Digital Handycam Sony Camcorder affixed to a tripod. The videos were converted into digital files using Windows Movie Maker software provided by Microsoft Windows and processed with the Quick Media Converter (V. 3.6.5) software. Although this method provided lower resolution than the regular CCD used in microscopy, it permitted a precise measurement of the phenomenon. XTT viability assay Giant cells were obtained by filtering the yeasts obtained from the lung of infected mice through 22 µm filters. Then, the yeast cells were separated from the filter by gently shaking the filters in 20 mL of water in 50 mL centrifuge tubes. After 20 minutes, the filters were removed, and the tubes centrifuged at 2000 r.p.m. Then, the cells were suspended in 2 mL of sterile water and the cell concentration was estimated using a haemocytometer. Approximately 105 giant cells were placed on 96 wells plates. In parallel, regular cells were obtained by overnight incubation in Sabouraud, washed with sterile water and counted with a haemocytometer. Then, the same number of cells (105) was placed in 96-wells plates. As negative controls, equal numbers of giant and regular cells were heat-inactivated (45 minutes at 60°C) and placed in different wells of the 96-wells plates. Viability measurement based on the reduction of 2,3-bis(2-methoxy-4-nitro-5-sulfophenyl)-2H-tetrazolium-5-carboxanilide inner salt (XTT) by living cells was performed as described in [21] with minor modifications, which involved the use of 1 mg/mL of XTT and 25 µM menadione. Optical density at 450 nm was recorded every 30 minutes for 18 hours in a iEMS Spectrophotometer (Thermofisher). Differences in metabolic activities were calculated by fold differences in the optical densities of the different wells. Indirect immunofluorescence To detect capsular features, we observed the immunofluorescence pattern after incubating the cells with the mAb 18B7 to the capsular polysaccharide as described above (see complement labelling and detection section), but using goat anti-mouse IgG1-FITC conjugated as the detection Ab. Yeast washed and incubated with the IgG1-FITC alone were used as controls. In some experiments, calcofluor (10 µg/mL) was included to visualize the cell wall. Wheat germ agglutinin staining To observe the presence of chitin-like structures, fungal cells with enlarged capsule (incubated in 10% Sabouradud in 50 mM MOPS buffer pH 7.3) were treated as in [33]. Briefly, the cells were washed with PBS and suspended in 4% p-formaldehyde cacodylate buffer (0.1 M, pH 7.2) and incubated for 30 min at room temperature. The fixed cells were washed in PBS and suspended in 100 µl of a 5 µg/mL of WGA conjugated to Alexa 594 (Molecular Probes, Invitrogen) for 1 hour at 37°C. Cell suspensions were mounted over glass slides and photographed with a Leica DMI 3000B fluorescence microscope. Vacuole staining and visualization by confocal microscopy To identify the vacuole in the yeast cells, we used the specific dye MDY-64 (Molecular Probes, Invitrogen, Eugene, Oregon) following the manufacturer's recommendations. Briefly, the cells were suspended in 10 mM HEPES buffer (pH 7.4) supplemented with 5% glucose. MDY-64 was dissolved in DMSO, and added to 106 cells at a final concentration of 10 µM. The cells were incubated for 5 minutes at room temperature, and washed twice with the same buffer. The cells were observed with a SP5 confocal microscope (Leica Microsystems). Cellular DNA content Cells were isolated as described above and fixed by heating the cells at 60°C for 45 minutes in PBS buffer. Then, the cells were separated in two parallel samples, and propidium iodide was added to one of them at a final concentration of 10 µg/mL. DNA content was analyzed using a FACSCalibur cytometer (Becton Dickinson). As a control, cells grown in vitro in Sabouraud medium were also analyzed. Nuclear staining To visualize the nucleus, the cells were treated with 3.7% formaldehyde for 30 minutes. Then, the cells were washed with PBS and DAPI was added at 0.3 µg/mL. The cells were incubated for 10 minutes at 37°C, and then washed twice with PBS. Finally, fluorescence was visualized in a Leica DM3000 microscope. Real-time PCR Giant cells were obtained by filtering the lung extracts through 22 µm filters as above. The filters were then placed in 50 mL tubes containing 20 mL of sterile water with moderate shaking, and after 20 minutes, the filters were removed. The tubes were centrifuged, and the pellet suspended in 0.5 mL of sterile water. Then, the cell concentration was determined using a haemocytometer. In parallel, cells obtained from a fresh liquid culture in Sabouraud were counted, and a cell suspension was prepared at the same concentration as that calculated for the giant cells. A real-time PCR using whole cells was then designed using equivalent numbers of the different cell types in the well. The reaction (final volume of 20 µL) contained 2.8×103 or 2.8×102 cells, 1.5 mM MgCl2, and 0.8 µM of ITS1 (5′TCCGTAGGTGAACCTGCGG3′) and ITS2 (5′GCTGCGTTCTTCATCGATGC3′) oligonucleotides, which amplify the ITS1 region from the ribosomal DNA. The real time was performed using the SensiMix Kit (Quantance) using the enzymes and SYBR green concentrations recommended by the manufacturer. The reaction mix was placed in a 96-wells plate and PCR was performed in a LC480 real-time PCR machine (Roche). We included wells with a known concentration of C. neoformans genomic DNA (20, 2, 0.2 and 0.02 ng) to quantify the results. The PCR was performed according to the following protocol: initial step of 10 minutes at 95°C and 45 amplification cycles (10 seconds at 95°C, 5 seconds at 54°C and 30 seconds at 72°C). Once the PCR was finished, a standard curve was calculated using the wells of the known genomic DNA concentration, and this curve was used to calculate the estimate of DNA present in each of the samples. Classification and Regression Trees (CART) analysis The CART system was proposed by Breiman et al. [68], and is characterized by binary-split searches, automatic self-validation procedures and surrogate splitters. This analysis is used to find associations between events with statistical support. CART analysis (CART 6.0 Salford Systems, Ca., USA) was used to find associations between giant cell formation and inflammation in the lungs. This analysis was performed with the following methodological conditions, Gini method, minimum cost tree regardless of the size for selecting the best tree, 10 v-fold-cross-validation, equal priors, no costs, and no penalties. Relative error of 0 means no error or perfect fit, whereas 1 represents the performance of random guessing. The statistical support for this association is given by the ROC curve. In this graph, specificity (false positive rate) vs. sensitivity (true positives rate) is calculated, and the area under the curve is analysed. When this area is 1 (100% of sensitivity and 0% false positives), a total agreement for the prediction is obtained. An area of 0.5 or below is indicative of random guess. Oxidative stress susceptibility Yeast cells (regular and giant) were obtained as described above. The cells were incubated in PBS with or without 1 mM H2O2 at a cell density of 104 cells/mL. After two hours of incubation at 37°C, 100 µL of each sample was plated on Sabouraud agar medium. In addition, a 1/10 dilution was done in PBS, and 100 µL of this dilution was also plated. The plates were incubated at 30°C for 48 hours and the colonies were enumerated. The survival was expressed as the percentage of colonies counted in the samples incubated with H2O2 compared to colonies of control samples not exposed to the oxidative agent. Statistical analysis Normal distribution in group samples were assessed using the Shapiro-Wilk and Kolmogorov-Smirnov tests using Unistat 5.0 (Unistat Ltd, London, England) and Analyse-it (Analyse-it Ltd, Leeds, England) softwares for Excel. Statistical differences between groups were tested using Student's t-Test (normal distributions) or Kruskal-Wallis test (non-parametric test for non-normally distributed samples). Differences were considered significant when p value was below 0.05. Ethics statement All the experiments involving the use of animals have been performed following the guidelines of the Bioethical and Animal Welfare Committee of the Instituto de Salud Carlos III (approved protocol PA-349, to be performed at the National Centre for Microbiology). Supporting Information Video S1 Live imaging of giant cells budding in vitro. Giant cells were obtained from infected mice as described in Materials and Methods and in Figure 4 legend. Live imaging of budding was recorded and processed as described in Materials and Methods. (2.39 MB AVI) Click here for additional data file. Video S2 Live imaging of giant cells budding in vitro. Giant cells were obtained from infected mice as described in Materials and Methods and in Figure 4 legend. Live imaging of budding was recorded and processed as described in Materials and Methods. (0.18 MB WMV) Click here for additional data file. Video S3 Live imaging of giant cells budding in vitro. Giant cells were obtained from infected mice as described in Materials and Methods and in Figure 4 legend. Live imaging of budding was recorded and processed as described in Materials and Methods. (1.71 MB AVI) Click here for additional data file. Video S4 Live imaging of giant cells budding in vitro. Giant cells were obtained from infected mice as described in Materials and Methods and in Figure 4 legend. Live imaging of budding was recorded and processed as described in Materials and Methods. (2.94 MB AVI) Click here for additional data file. Video S5 Phagocytosis of C. neoformans by murine-like macrophages. The video shows the interaction of RAW264.7 macrophage cell lines exposed to C. neoformans H99 strain at a ratio 1∶2. Videos were performed as described in Materials and Methods. Pictures were taken every 5 minutes, and 5 frames per second are shown in the video (1 second of the video is equivalent to 25 minutes of real time). A field of 100 µm width is shown. (5.46 MB AVI) Click here for additional data file. Video S6 Macrophage fusion after C. neoformans phagocytosis and cell division. RAW264.7 macrophage cell lines were exposed to C. neoformans H99 strain at a ratio 1∶2. Videos were performed as described in Materials and Methods. Pictures were taken every 5 minutes, and 5 frames per second are shown in the video (1 second of the video is equivalent to 25 minutes of real time). A field of 100 µm width is shown. (6.16 MB AVI) Click here for additional data file. Video S7 C. neoformans intracellular replication. Macrophages and C. neoformans grown in Sabouraud were mixed as described in supplemental Videos S5 and S6. The pictures were taken every 2 minutes, and 5 frames per seconds are shown in the video (1 second of the video is equivalent to 10 minutes of real time). (6.16 MB AVI) Click here for additional data file. Video S8 Interaction between giant cells and macrophages. Macrophages and C. neoformans giant cells were mixed at 1∶2 ratio as described in Materials and Methods. Pictures were taken every 3 minutes, and 5 frames per second are shown in the video (1 second of the video is equivalent to 15 minutes of real time). A field of 100 µm width is shown. (5.09 MB AVI) Click here for additional data file. Video S9 Interaction between giant cells and macrophages. Macrophages and C. neoformans giant cells were mixed at 1∶2 ratio as described in Materials and Methods. Pictures and video were taken and assemble as described in Supporting Video S8. Scale bar denotes 50 µm. (2.04 MB WMV) Click here for additional data file.

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Greetje Vande Velde:

ORCID: http://orcid.org/0000-0002-5633-3993

greetje.vandevelde@kuleuven.be

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Journal ID (nlm-ta): Sci Rep

Journal ID (iso-abbrev): Sci Rep

Title: Scientific Reports

Publisher: Nature Publishing Group UK (London )

ISSN (Electronic): 2045-2322

Publication date (Electronic): 14 February 2018

Publication date PMC-release: 14 February 2018

Publication date Collection: 2018

Volume: 8

Electronic Location Identifier: 3009

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[1 ]ISNI 0000 0001 0668 7884, GRID grid.5596.f, Biomedical MRI unit/MoSAIC, Department of Imaging and Pathology, KU Leuven, Herestraat 49 O & N1 box 505, ; 3000 Leuven, Belgium

[2 ]ISNI 0000 0001 0668 7884, GRID grid.5596.f, Laboratory of Clinical Bacteriology and Mycology, Department of Microbiology and Immunology, KU Leuven, Herestraat 49 box 6711, ; 3000 Leuven, Belgium

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DOI: 10.1038/s41598-018-20545-4

PMC ID: 5813038

PubMed ID: 29445211

SO-VID: b0e00cd5-b0be-497e-b8cf-4fb6fdf0f28a

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Date received : 24 May 2017

Date accepted : 21 January 2018

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