Mediators of Inflammation

Mediators of Inflammation / 2015 / Article

Research Article | Open Access

Volume 2015 |Article ID 208491 | https://doi.org/10.1155/2015/208491

Pei Li, Ting Wu, Xin Su, Yi Shi, "Activation of Vitamin D Regulates Response of Human Bronchial Epithelial Cells to Aspergillus fumigatus in an Autocrine Fashion", Mediators of Inflammation, vol. 2015, Article ID 208491, 14 pages, 2015. https://doi.org/10.1155/2015/208491

Activation of Vitamin D Regulates Response of Human Bronchial Epithelial Cells to Aspergillus fumigatus in an Autocrine Fashion

Academic Editor: Genhong Cheng
Received23 Nov 2014
Accepted10 Feb 2015
Published15 Apr 2015

Abstract

Aspergillus fumigatus (A. fumigatus) is one of the most common fungi to cause diseases in humans. Recent evidence has demonstrated that airway epithelial cells play an important role in combating A. fumigatus through inflammatory responses. Human airway epithelial cells have been proven to synthesize the active vitamin D, which plays a key role in regulating inflammation. The present study was conducted to investigate the impact of A. fumigatus infection on the activation of vitamin D and the role of vitamin D activation in A. fumigatus-elicited antifungal immunity in normal human airway epithelial cells. We found that A. fumigatus swollen conidia (SC) induced the expression of 1α-hydroxylase, the enzyme catalyzing the synthesis of active vitamin D, and vitamin D receptor (VDR) in 16HBE cells and led to increased local generation of active vitamin D. Locally activated vitamin D amplified SC-induced expression of antimicrobial peptides in 16HBE cells but attenuated SC-induced production of cytokines in an autocrine fashion. Furthermore, we identified β-glucan, the major A. fumigatus cell wall component, as the causative agent for upregulation of 1α-hydroxylase and VDR in 16HBE cells. Therefore, activation of vitamin D is inducible and provides a bidirectional regulation of the responses to A. fumigatus in 16HBE cells.

1. Introduction

Aspergillus fumigatus (A. fumigatus) is ubiquitous, saprophytic, and airborne mold that causes a range of lung diseases in humans [1, 2]. Airway epithelial cells serve as the first line of the host defense system against A. fumigatus. Recognition of A. fumigatus by airway epithelial cells results in gene transcription and secretion of a variety of effector molecules, including antimicrobials, type I interferons, and proinflammatory cytokines and chemokines, suggesting that these cells play an important role in combating inhaled A. fumigatus infection through inflammatory responses [38]. Meanwhile, airway epithelial cells have a key role in maintaining the inflammatory homeostasis in order to keep gas-exchange surface integrity and stretch the discriminatory powers of the immune system to its limits [9]. As exposure to inhaled A. fumigatus conidia is common and lifelong [10], and hosts with normal immunity rarely develop pulmonary aspergillosis, we were curious about how airway epithelial cells maintain the inflammatory balance when interacting with A. fumigatus in the lungs of healthy hosts.

Vitamin D, a pluripotent hormone whose functions extend beyond its classical role in calcium homeostasis, has recently been recognized as an important modulator of the pulmonary defense and inflammatory processes [1115]. The active form of vitamin D, 1,25-dihydroxyvitamin D3 (1,25D3), has been shown to decrease the production of respiratory syncytial virus-induced NF-kappaB-linked chemokines and cytokines [16] and to inhibit proinflammatory cytokine release from LPS stimulated cystic fibrosis (CF) respiratory epithelial cells [17]. Furthermore, 1,25D3 has been shown to induce antimicrobial actions in epithelial cells by virtue of its ability to upregulate expression and secretion of the antimicrobial peptide LL-37 [1721]. Thus, vitamin D may dampen the inflammatory response to pathogens without negatively affecting pathogens clearance. The biological effects of vitamin D are achieved through the regulation of gene expression mediated by the vitamin D receptor (VDR), which is expressed in virtually all cells in the body [11, 22]. An increasing variety of tissues and cell types have been found to express 1α-hydroxylase, the enzyme responsible for the final and rate-limiting step of active vitamin D synthesis, in which 25-hydroxyvitamin D3 (25D3), the primary circulating or storage form of vitamin D, is converted to 1,25D3 [19, 2328]. Local synthesis of tissue-specific active vitamin D rather than systemic production is thought to be responsible for the immunomodulatory effects of vitamin D. Human airway epithelial cells have also been proven to express 1α-hydroxylase [19], indicating that active vitamin D can be produced locally within the airway epithelium, which may act in an autocrine or paracrine fashion to modulate airway immune function.

Vitamin D deficiency is highly prevalent and has been associated with increased susceptibility to respiratory infections, including ABPA [2940]. In our previous study, we found that vitamin D deficiency caused an aggravated inflammatory response and an impaired host defense to pulmonary challenge with A. fumigatus in immunocompetent mice [41]. Based on the fact that local synthesis of active vitamin D occurs in airway epithelium, we investigated the role of A. fumigatus in modulating the expression of 1α-hydroxylase and VDR in normal human airway epithelial cells and how this affects antifungal immunity in the airway epithelium.

2. Materials and Methods

2.1. A. fumigatus Strain and Preparation of Conidia

The A. fumigatus strain was obtained from a fatal case of pulmonary aspergillosis at the Department of Respiratory and Critical Care Medicine, Jinling Hospital, Nanjing University School of Medicine. Conidia were harvested by washing a 7-day-old slant culture on Sabouraud dextrose agar (10 g/L peptone, 40 g/L glucose, and 15 g/L agar) with phosphate-buffered saline (PBS) supplemented with 0.1% Tween 20. The suspension was filtered through a 40 μm cell strainer (Falcon) to separate conidia from the contaminating mycelia, and the absence of mycelia in the filtrate was verified microscopically. Inactivated resting or swollen conidia were obtained as described previously [42, 43]. Resting conidia (RC) were washed with PBS three times and counted. For experiments with swollen conidia (SC), conidia were incubated for 3.5 h in RPMI 1640 (Invitrogen Gibco, Carlsbad, CA, USA) at 37°C. Since human cells have to be exposed to the different forms of A. fumigatus for various periods of time (including 24 hours to allow the RC to swell, germinate, and grow into hyphae), all A. fumigatus morphotypes were heat-inactivated at 90°C for 60 min.

2.2. Cell Line and Growth Conditions

16HBE cells were originally isolated from human bronchial epithelial cells and transformed with the origin-defective simian virus 40 genome. The 16HBE cells used in this study were kindly provided by Professor Laiyu Liu (Department of Respiratory Diseases, Nanfang Hospital, Southern Medical University, Guangzhou, China). 16HBE cultures were maintained in high glucose Dulbecco’s Modified Eagle’s Medium (DMEM; Invitrogen Gibco) containing 10% fetal bovine serum (FBS; Invitrogen Gibco), 2 nM L-glutamine, 100 U/mL penicillin, and 100 μg/mL streptomycin at 37°C in a humidified 5% CO2 atmosphere.

2.3. Cell Treatments

16HBE cells were treated with RC or SC (multiplicity of infection (MOI) = 1-2), or β-glucan (Sigma-Aldrich, St. Louis, MO, USA) in the absence or presence of inactive vitamin D (25D3; Sigma-Aldrich) or active vitamin D (1,25D3; Sigma-Aldrich) for appropriate durations. The concentrations of β-glucan, 25D3, and 1,25D3 used as well as the treatment durations are detailed in the figure legends.

2.4. Western Blot Analysis

Cells were lysed with a 1% NP-40 hypotonic lysis buffer containing 1 mM phenylmethanesulfonyl fluoride, 1% aprotinin, and 1 mM sodium vanadate (Sigma-Aldrich). Cell lysates were mixed with loading buffer, boiled for 5 min, resolved by 12% sodium dodecyl sulfate-polyacrylamide gel electrophoresis, and transferred onto a polyvinylidene fluoride membrane (Millipore, Bedford, MA, USA). This membrane was subsequently blocked with 5% nonfat milk. Western blot analysis was then performed in accordance with standard protocols using antibodies (Santa Cruz Biotechnology, Santa Cruz, CA, USA) against 1α-hydroxylase, VDR, LL-37, or β-defensin-2 (HBD2), followed by incubation with relevant horseradish peroxidase-conjugated secondary antibodies (Santa Cruz Biotechnology). Relative changes in protein expression were estimated from the mean pixel density using Quantity One software 4.6.2 (Bio-Rad, Hercules, CA, USA), normalized to glyceraldehyde-3-phosphate dehydrogenase (GAPDH), and presented as relative density units.

2.5. Quantitative Real-Time Polymerase Chain Reaction (PCR)

Total RNA was isolated from cultured cells, and cDNA synthesis was performed on the RevertAid First Strand cDNA Synthesis Kit (Thermo Scientific, Waltham, MA, USA) according to the manufacturer’s instructions. Data were analyzed using the delta-delta Ct method and normalized to the housekeeping gene GAPDH. Each PCR run included a no-template control. Results are expressed as fold change from control. Specific primer sets used are as follows (5′ to 3′): 1α-hydroxylase, AAC CCT GAA CAA CGT AGT CTG CGA (forward) and ATG GTC AAC AGC GTG GAC ACA AA (reverse); VDR, CTG CTG AAG TCA AGT GCC AT (forward) and ACA AGT ACC GCG TCA GTG AC (reverse); tumor necrosis factor-α (TNF-α), CTG GGA TTC AGG AAT GTG TG (forward) and TTG ATC CCT GAC ATC TGG AA (reverse); interleukin (IL)-1β, CCA TGC AAT TTG TGT CTT CC (forward) and AGC CTG GAC TTT CCT GTT GT (reverse); IL-6, CTG CGC AGC TTT AAG GAG TT (forward) and TCC ACT GGG CAC AGA ACT TA (reverse); IL-8, ATC TGG CAA CCC TAG TCT GC (forward) and GTG AGG ACA TGT GGA AGC AC (reverse). All primers were synthesized by GenScript (Nanjing, China).

2.6. Quantitative Determination of 1,25D3

1,25D3 was quantified using an enzyme immunoassay kit for 1,25D3 (Immunodiagnostic Systems, Boldon, UK) according to the manufacturer’s instructions.

2.7. Enzyme Linked Immunosorbent Assay (ELISA)

Cell-free supernatants were collected and aliquots were stored at −70°C until use. The protein levels of TNF-α and IL-8 in medium were measured with commercially available cytokine specific ELISA kits (R&D Systems, Minneapolis, MN, USA) according to the manufacturers’ recommendations.

2.8. RNA Interference Experiments

16HBE cells were transfected with small interfering RNAs (siRNAs; GenePharma, Shanghai, China) against 1α-hydroxylase or VDR when reaching 70% confluence in six-well plates. A nonspecific siRNA (control siRNA) was used in each experiment as a negative control. For each transfection, 100 pmol siRNA per well was diluted in 250 μL serum-free Opti-MEM I medium (Invitrogen Gibco) and gently mixed with 5 μL Lipofectamine 2000 (Invitrogen Gibco) diluted in 250 μL Opti-MEM I medium. After incubation for 25 min at room temperature, siRNA and Lipofectamine 2000 complexes were added to each well of six-well culture plates. The plates were gently mixed by rocking back and forth. Transfected cells were incubated for 24 h before being stimulated with A. fumigatus. Cell treatments are detailed in the figure legends.

2.9. Statistical Analysis

Data were expressed as either a representative experiment or the mean ± standard error (SE) of three independent experiments. Except where otherwise indicated, Student’s paired -test was used to compare the difference between groups, with values < 0.05 considered statistically significant.

3. Results

3.1. A. fumigatus Induces the Expression of 1α-Hydroxylase and VDR and the Conversion of 25D3 to 1,25D3 in 16HBE Cells

To explore whether A. fumigatus has any effect on the expression of 1α-hydroxylase and VDR in 16HBE cells, we stimulated 16HBE cells with RC and SC. We observed basal expression of 1α-hydroxylase and VDR in nontreated 16HBE cells at both the protein and mRNA levels (Figure 1(a)). In cells stimulated with RC, protein and mRNA expression of 1α-hydroxylase and VDR remained at basal levels (Figure 1(a)). When challenged with SC, protein and mRNA expression of 1α-hydroxylase and VDR significantly increased in a time-dependent manner, peaking at 24 h after stimulation (Figure 1(a)).

We then examined whether A. fumigatus affects the conversion of inactive vitamin D to its active form. 16HBE cells stimulated with A. fumigatus were treated with increasing concentrations of inactive vitamin D (25D3), and the levels of active vitamin D (1,25D3) in supernatants were measured 24 h later (Figure 1(b)). We found that 16HBE cells could convert the inactive vitamin D to the active form in a dose-dependent manner without other stimuli. Consistent with the increased expression of 1α-hydroxylase, 16HBE cells stimulated with SC generated greater amounts of 1,25D3 than cells without any treatment when exposed to inactive vitamin D. No differences were found in the amounts of 1,25D3 in supernatants between nontreated cells and cells stimulated with RC in the presence of inactive vitamin D. These data were in agreement with the unchanged expression of 1α-hydroxylase in 16HBE cells challenged with RC. When inactive vitamin D was added to medium without cells, no active vitamin D was detected.

3.2. Locally Activated Vitamin D Synergistically Increases the Expression of LL-37 and HBD2 in 16HBE Cells Infected with A. fumigatus but Attenuates A. fumigatus-Induced Production of Chemokines and Cytokines

Vitamin D plays an important role in innate immunity. The active form of vitamin D has been proven to prevent infections by bacteria (e.g., M. tuberculosis) or viruses by inducing the expression of the cathelicidin antimicrobial peptide and β-defensins [44, 45] and decreasing the inflammatory response to microbial infections [16, 4649]. Exposure of respiratory epithelial cells to A. fumigatus increases the release of LL-37 and β-defensins and the expression of chemokines and cytokines that initiate an inflammatory reaction [3, 57]. Having established that 16HBE cells generate more active vitamin D and have upregulated VDR expression after stimulation with A. fumigatus, we hypothesized that when exposed to the inactive form of vitamin D, 16HBE cells stimulated with A. fumigatus would convert it to 1,25D3, which would alter the expression of inflammatory mediators and antimicrobial peptides induced by A. fumigatus in an autocrine fashion. To test this hypothesis, we initially treated 16HBE cells with 10−7 M of either the active or inactive form of vitamin D (1,25D3 or 25D3) and examined the protein expression of LL-37 and HBD2 in 16HBE cells stimulated with A. fumigatus for 24 h (Figure 2(a)). We found that both forms of vitamin D enhanced the basal expression of LL-37 and HBD2 in 16HBE cells to a similar extent. Regardless of treatment with either form of vitamin D, stimulation with RC did not change the expression of LL-37 or HBD2 in 16HBE cells. When stimulated with SC, the expression of LL-37 and HBD2 increased significantly. The presence of either 1,25D3 or 25D3 significantly augmented SC-induced expression of LL-37 and HBD2 to a similar extent in 16HBE cells.

Subsequently, we sought to examine whether locally activated vitamin D affects A. fumigatus-induced production of TNF-α, IL-1β, IL-6, and IL-8 in 16HBE cells. We found significant increases in the mRNA expression of TNF-α, IL-1β, IL-6, and IL-8 after stimulation with both RC and SC for 24 h (Figure 2(b)), an effect that was more profound with SC stimulation. Treatment with either 1,25D3 or 25D3 significantly inhibited SC- and RC-induced upregulation of all these inflammatory mediators in 16HBE cells to a similar extent (Figure 2(b)). We did not find any changes in the mRNA expression of TNF-α, IL-1β, IL-6, and IL-8 when 16HBE cells were exposed to either form of vitamin D alone (Figure 2(b)). We also measured the concentrations of TNF-α and IL-8 in culture supernatants of 16HBE cells using the appropriate ELISA (Figure 2(c)). Results showed that the protein expression of TNF-α and IL-8 induced by A. fumigatus was significantly attenuated in the presence of both forms of vitamin D.

Taken together, these data show that 1,25D3 generated by 16HBE cells is indeed biologically active and regulates the antifungal immune responses of 16HBE cells to A. fumigatus to a similar extent as exogenous exposure to 1,25D3. Higher expression of LL-37 and HBD2 enhanced by vitamin D may have the potential of locally enhancing the innate immunity within the lungs. The inhibited expression of inflammatory mediators may avoid tissue damage caused by excessive inflammation.

3.3. Silencing of VDR or 1α-Hydroxylase Attenuates the Effects of Locally Activated Vitamin D on the Response of 16HBE Cells to A. fumigatus

To definitively link 1,25D3 generation to SC-induced 1α-hydroxylase expression in 16HBE cells and to confirm that the local activation of vitamin D directly mediates the responses of 16HBE cells to stimulation with A. fumigatus in an autocrine manner, we next explored whether silencing of 1α-hydroxylase or VDR would reverse the effects of locally activated vitamin D on the response of 16HBE cells to A. fumigatus. We performed knockdown experiments using siRNAs against 1α-hydroxylase or VDR. Knockdown of the 1α-hydroxylase or VDR gene was confirmed by Western blot and quantitative real-time analyses (Figures 3(a) and 3(b)). When cells were transfected with the control siRNA, both forms of vitamin D synergistically enhanced SC-induced upregulation of LL-37 and HBD2 expression and reduced SC-induced expression of TNF-α, IL-1β, IL-6, and IL-8, as shown previously in nontransfected cells (Figures 3(c)3(f)). When VDR expression was silenced, both forms of vitamin D no longer had any modulatory effect on LL-37 and HBD2 (Figures 3(c) and 3(e)), and their inhibition of chemokines and cytokines induced by SC was moderately, but significantly, impaired (Figure 3(d)). However, when 1α-hydroxylase expression was silenced, only the effect of 25D3 was eliminated (Figures 3(e) and 3(f)). Moreover, silencing of 1α-hydroxylase significantly reduced its ability to convert 25D3 to 1,25D3 in 16HBE cells challenged with SC (Figure 3(g)). This finding further supports our primary hypothesis that when exposed to the inactive form of vitamin D, 16HBE cells stimulated with A. fumigatus would convert it to 1,25D3, further altering the expression of inflammatory mediators and antimicrobial peptides in an autocrine fashion.

3.4. β-Glucan, an A. fumigatus Wall Component, Increases the Expression of 1α-Hydroxylase and VDR, and Vitamin D Synergistically Increases β-Glucan-Induced Expression of Antimicrobial Peptides but Attenuates β-Glucan-Induced Expression of Chemokines and Cytokines in 16HBE Cells

The cell walls of fungi, including A. fumigatus, are in a large part made up of β-glucan. We thus examined whether β-glucan has any effect on the expression of 1α-hydroxylase and VDR in 16HBE cells. We stimulated 16HBE cells with β-glucan for 24 h and examined the expression of 1α-hydroxylase and VDR at both protein and mRNA level. Results showed that β-glucan significantly increased the expression of 1α-hydroxylase and VDR in a dose-dependent manner (Figure 4(a)). Furthermore, 16HBE cells converted more 25D3 to 1,25D3 when β-glucan was present (Figure 4(b)). We then investigated the protein expression of LL-37 and HBD2 and found that β-glucan could induce LL-37 and HBD2 expression by itself. 16HBE cells exposed to β-glucan in the presence of inactive or active vitamin D had significantly greater amplification of LL-37 and HBD2 expression than by β-glucan alone (Figure 4(c)). In contrast to the synergistic effect of vitamin D exposure and β-glucan on LL-37 and HBD2 expression, both forms of vitamin D decreased β-glucan-induced expression of TNF-α, IL-1β, IL-6, and IL-8 at the mRNA level (Figure 4(d)). These data suggest that β-glucan induces vitamin D conversion and recognition in airway epithelial cells and that vitamin D synergizes with β-glucan to induce expression of antimicrobial proteins but attenuates β-glucan-induced expression of chemokines and cytokines in airway epithelial cells.

4. Discussion

Our results showed that A. fumigatus SC induced 1α-hydroxylase and VDR expression in 16HBE cells and led to increased local activation of vitamin D. Locally activated vitamin D could synergize with SC to further amplify the expression of LL-37 and HBD2 in 16HBE cells but attenuate SC-induced production of chemokines and cytokines in an autocrine fashion. Furthermore, we identified β-glucan, the major component of the A. fumigatus cell wall, as the causative agent responsible for upregulation of 1α-hydroxylase and VDR expression in 16HBE cells.

In this study, A. fumigatus RC were also used for the stimulation of 16HBE cells, but no induction of 1α-hydroxylase or VDR was observed. RC are surrounded by a hydrophobic layer of rodlet proteins [42]. The rodlet layer could protect conidia from recognition and phagocytosis by the host defense system. Conidial swelling releases the protective rodlet layer and exposes β-glucan [42]. Therefore, exposed β-glucan might be responsible for mediating the effect that A. fumigatus impacts vitamin D activation and signal transduction and thus affects the antifungal immunity in the airway epithelium. Our finding that β-glucan increased the expression of 1α-hydroxylase and VDR in 16HBE cells further confirmed this presumption. In human macrophages, toll-like receptor 2/1 (TLR2/1) and TLR8 activation upregulates the expression of VDR and 1α-hydroxylase [50, 51]. In the human skin, 1α-hydroxylase expression was increased after being wounded [25]. TLR2/6 ligands were also found to increase 1α-hydroxylase expression in cultured keratinocytes [25]. In human respiratory epithelial cells, respiratory syncytial virus (RSV) and Poly(IC), a synthetic analog of dsRNA, but not bacterial cell wall components (TLR2 ligands), were observed to increase the expression of 1α-hydroxylase [19]. The factors associated with regulation of 1α-hydroxylase and VDR expression in barrier tissues are less clear. Whether this phenomenon is common when pathogenic microorganisms are recognized by cells at the barrier sites is still unknown, awaiting further exploration.

Our results are different from those reported by Coughlan et al. who found that A. fumigatus downregulated VDR expression in airway epithelial cells and identified gliotoxin as the causative agent responsible for mediating this effect [52]. This obvious discrepancy between the two studies could be explained by the use of A. fumigatus at different growth stages. During asexual growth, the morphological form of A. fumigatus changes from RC to SC which then forms germ tubes that continue growing in hyphal form. The surfaces of various A. fumigatus morphotypes differ from each other and, consequently, the reaction of host cells may vary towards different A. fumigatus growth forms [53, 54]. Our previous study has shown that in the lungs of immunocompetent hosts A. fumigatus could hardly grow into hyphae [40]. So in the present study, we used SC for stimulation, in which β-glucan was one of the major antigens to be recognized by the airway epithelium. In order to find out whether Aspergillus colonization in the lungs of ABPA patients affect VDR expression in airway epithelial cells, Coughlan et al. stimulated these cells with culture filtrates from A. fumigatus grown for 4 days, when the hyphae of A. fumigatus form [52]. Gliotoxin is secreted by the hyphal form of A. fumigatus [52, 55], so hypha formation might compromise the positive impact conferred by SC-induced vitamin D synthesis and VDR expression in the airway epithelium. Our findings are an important supplement to the findings reported by Coughlan et al. These two completely different results also indicate host morphotype’s specific reactions to A. fumigatus.

Vitamin D synergizes with both SC and β-glucan to further amplify the expression of the two antimicrobial proteins, LL-37 and HBD2, which have antifungal activity [56, 57]. And defensins have additional activities such as the chemoattraction of immature dendritic cells, T cells, and monocytes as well as activation of the professional antigen-presenting cells [5860]. Conversely, vitamin D attenuated the induction of proinflammatory cytokines and the chemokine IL-8 by A. fumigatus and β-glucan. Microbial infection causes cytokine-associated inflammation to remove pathogens. However, excessive or uncontrolled release of proinflammatory cytokines and chemokines can damage the integrity of gas-exchange surfaces, even cause immunodeficiency, septic shock, or induction of autoimmunity, and eventually impair disease eradication [9, 6163]. It has been proposed that in fungal infections disease pathology may be attributable to an aggravated or dysregulated host inflammatory response that results in extensive tissue damage. In mice with chronic granulomatous disease, the intrinsic, genetically determined failure to control inflammation after exposure to sterile fungal components determined the animals’ inability to resolve an infection with A. fumigatus [64]. In neutropenic patients with aspergillosis, the clinical and radiologic pulmonary deterioration during neutrophil recovery may be aspergillosis-related immune reconstitution inflammatory syndrome [65]. Thus, local synthesis of active vitamin D might be one of the ways by which the airway epithelium enhances its antifungal capability while limiting inflammation-induced tissue damage in challenge with A. fumigatus, suggesting the potential beneficial effects of proper 25D3 levels in blood circulation on pathogen-driven inflammation in respiratory epithelium.

In conclusion, we have shown that A. fumigatus led to more significant local activation of vitamin D, which resulted in enhanced antimicrobial peptide production and attenuating cytokine release in respiratory epithelial cells. We confirmed locally enhanced innate immunity and mitigated inflammation after exposure of airway epithelial cells stimulated with A. fumigatus to the inactive vitamin D. Hence, vitamin D might provide a novel treatment option that may reduce lung inflammation and disease severity in Aspergillus infection, without negatively affecting Aspergillus clearance. Given that low serum 25D3 levels are prevalent and have been associated with increased incidences of acute respiratory symptoms and lower respiratory infection [29, 36, 37], supplemental administration of vitamin D might be advocated for its potential to prevent A. fumigatus infection. It will be important to investigate whether disturbed vitamin D metabolism contributes to the pathopoiesis of A. fumigatus and whether local activation of vitamin D could enhance the barrier function of airway epithelium and delay the hyphal formation of A. fumigatus. Our findings may provide valuable insight into the role of vitamin D in susceptibility to pulmonary aspergillosis in immunosuppressed patients.

Conflict of Interests

The authors declare that there is no conflict of interests regarding the publication of this paper.

Authors’ Contribution

Xin Su and Yi Shi contributed equally to this work.

Acknowledgments

The authors would like to thank Ming Fan and Mei Huang for providing highly purified Aspergillus fumigatus.

References

  1. C. Kosmidis and D. W. Denning, “The clinical spectrum of pulmonary aspergillosis,” Thorax, vol. 70, no. 3, pp. 270–277, 2015. View at: Publisher Site | Google Scholar
  2. C. Godet, B. Philippe, F. Laurent, and J. Cadranel, “Chronic pulmonary aspergillosis: an update on diagnosis and treatment,” Respiration, vol. 88, no. 2, pp. 162–174, 2014. View at: Publisher Site | Google Scholar
  3. L. Alekseeva, D. Huet, F. Féménia et al., “Inducible expression of beta defensins by human respiratory epithelial cells exposed to Aspergillus fumigatus organisms,” BMC Microbiology, vol. 9, article 33, 2009. View at: Publisher Site | Google Scholar
  4. S. G. Filler and D. C. Sheppard, “Fungal invasion of normally non-phagocytic host cells,” PLoS pathogens, vol. 2, no. 12, article e129, 2006. View at: Publisher Site | Google Scholar
  5. Z. Zhang, R. Liu, J. A. Noordhoek, and H. F. Kauffman, “Interaction of airway epithelial cells (A549) with spores and mycelium of Aspergillus fumigatus,” Journal of Infection, vol. 51, no. 5, pp. 375–382, 2005. View at: Google Scholar
  6. C. Cunha, M. Di Ianni, S. Bozza et al., “Dectin-1 Y238X polymorphism associates with susceptibility to invasive aspergillosis in hematopoietic transplantation through impairment of both recipient- and donor-dependent mechanisms of antifungal immunity,” Blood, vol. 116, no. 24, pp. 5394–5402, 2010. View at: Publisher Site | Google Scholar
  7. W.-K. Sun, X. Lu, X. Li et al., “Dectin-1 is inducible and plays a crucial role in Aspergillus-induced innate immune responses in human bronchial epithelial cells,” European Journal of Clinical Microbiology and Infectious Diseases, vol. 31, no. 10, pp. 2755–2764, 2012. View at: Publisher Site | Google Scholar
  8. V. Balloy, J.-M. Sallenave, Y. Wu et al., “Aspergillus fumigatus-induced interleukin-8 synthesis by respiratory epithelial cells is controlled by the phosphatidylinositol 3-kinase, p38 MAPK, and ERK1/2 pathways and not by the toll-like receptor-MyD88 pathway,” Journal of Biological Chemistry, vol. 283, no. 45, pp. 30513–30521, 2008. View at: Publisher Site | Google Scholar
  9. P. G. Holt, D. H. Strickland, M. E. Wikström, and F. L. Jahnsen, “Regulation of immunological homeostasis in the respiratory tract,” Nature Reviews Immunology, vol. 8, no. 2, pp. 142–152, 2008. View at: Publisher Site | Google Scholar
  10. T. R. T. Dagenais and N. P. Keller, “Pathogenesis of Aspergillus fumigatus in invasive aspergillosis,” Clinical Microbiology Reviews, vol. 22, no. 3, pp. 447–465, 2009. View at: Publisher Site | Google Scholar
  11. C. J. Rosen, J. S. Adams, D. D. Bikle et al., “The nonskeletal effects of vitamin D: an endocrine society scientific statement,” Endocrine Reviews, vol. 33, no. 3, pp. 456–492, 2012. View at: Publisher Site | Google Scholar
  12. D. A. Hughes and R. Norton, “Vitamin D and respiratory health,” Clinical and Experimental Immunology, vol. 158, no. 1, pp. 20–25, 2009. View at: Publisher Site | Google Scholar
  13. S. Hansdottir and M. M. Monick, “Vitamin D effects on lung immunity and respiratory diseases,” Vitamins & Hormones, vol. 86, pp. 217–237, 2011. View at: Publisher Site | Google Scholar
  14. V. Lagishetty, N. Q. Liu, and M. Hewison, “Vitamin D metabolism and innate immunity,” Molecular and Cellular Endocrinology, vol. 347, no. 1-2, pp. 97–105, 2011. View at: Publisher Site | Google Scholar
  15. M. Hewison, “An update on vitamin D and human immunity,” Clinical Endocrinology, vol. 76, no. 3, pp. 315–325, 2012. View at: Publisher Site | Google Scholar
  16. S. Hansdottir, M. M. Monick, N. Lovan, L. Powers, A. Gerke, and G. W. Hunninghake, “Vitamin D decreases respiratory syncytial virus induction of NF-κB-linked chemokines and cytokines in airway epithelium while maintaining the antiviral state,” Journal of Immunology, vol. 184, no. 2, pp. 965–974, 2010. View at: Publisher Site | Google Scholar
  17. P. McNally, C. Coughlan, G. Bergsson et al., “Vitamin D receptor agonists inhibit pro-inflammatory cytokine production from the respiratory epithelium in cystic fibrosis,” Journal of Cystic Fibrosis, vol. 10, no. 6, pp. 428–434, 2011. View at: Publisher Site | Google Scholar
  18. S. Yim, P. Dhawan, C. Ragunath, S. Christakos, and G. Diamond, “Induction of cathelicidin in normal and CF bronchial epithelial cells by 1,25-dihydroxyvitamin D3,” Journal of Cystic Fibrosis, vol. 6, no. 6, pp. 403–410, 2007. View at: Publisher Site | Google Scholar
  19. S. Hansdottir, M. M. Monick, S. L. Hinde, N. Lovan, D. C. Look, and G. W. Hunninghake, “Respiratory epithelial cells convert inactive vitamin D to its active form: potential effects on host defense,” Journal of Immunology, vol. 181, no. 10, pp. 7090–7099, 2008. View at: Publisher Site | Google Scholar
  20. V. Lagishetty, R. F. Chun, N. Q. Liu, T. S. Lisse, J. S. Adams, and M. Hewison, “1α-Hydroxylase and innate immune responses to 25-hydroxyvitamin D in colonic cell lines,” Journal of Steroid Biochemistry and Molecular Biology, vol. 121, no. 1-2, pp. 228–233, 2010. View at: Publisher Site | Google Scholar
  21. L. McMahon, K. Schwartz, O. Yilmaz, E. Brown, L. K. Ryan, and G. Diamond, “Vitamin D-mediated induction of innate immunity in gingival epithelial cells,” Infection and Immunity, vol. 79, no. 6, pp. 2250–2256, 2011. View at: Publisher Site | Google Scholar
  22. Y. Wang, J. Zhu, and H. F. DeLuca, “Where is the vitamin D receptor?” Archives of Biochemistry and Biophysics, vol. 523, no. 1, pp. 123–133, 2012. View at: Publisher Site | Google Scholar
  23. J. S. Adams and M. A. Gacad, “Characterization of 1α-hydroxylation of vitamin D3 sterols by cultured alveolar macrophages from patients with sarcoidosis,” Journal of Experimental Medicine, vol. 161, no. 4, pp. 755–765, 1985. View at: Publisher Site | Google Scholar
  24. D. D. Bikle, M. K. Nemanic, E. Gee, and P. Elias, “1,25-Dihydroxyvitamin D3 production by human keratinocytes. Kinetics and regulation,” Journal of Clinical Investigation, vol. 78, no. 2, pp. 557–566, 1986. View at: Publisher Site | Google Scholar
  25. J. Schauber, R. A. Dorschner, A. B. Coda et al., “Injury enhances TLR2 function and antimicrobial peptide expression through a vitamin D-dependent mechanism,” Journal of Clinical Investigation, vol. 117, no. 3, pp. 803–811, 2007. View at: Publisher Site | Google Scholar
  26. V. Tangpricha, J. N. Flanagan, L. W. Whitlatch et al., “25-hydroxyvitamin D-1α-hydroxylase in normal and malignant colon tissue,” The Lancet, vol. 357, no. 9269, pp. 1673–1674, 2001. View at: Publisher Site | Google Scholar
  27. C. M. Kemmis, S. M. Salvador, K. M. Smith, and J. Welsh, “Human mammary epithelial cells express CYP27B1 and are growth inhibited by 25-hydroxyvitamin D-3, the major circulating form of vitamin D-3,” Journal of Nutrition, vol. 136, no. 4, pp. 887–892, 2006. View at: Google Scholar
  28. N. H. Bell, “Renal and nonrenal 25-hydroxyvitamin D-1alpha-hydroxylases and their clinical significance,” Journal of Bone and Mineral Research, vol. 13, no. 3, pp. 350–353, 1998. View at: Publisher Site | Google Scholar
  29. M. F. Holick, “Medical progress: vitamin D deficiency,” The New England Journal of Medicine, vol. 357, no. 3, pp. 266–281, 2007. View at: Publisher Site | Google Scholar
  30. T. D. Thacher and B. L. Clarke, “Vitamin D insufficiency,” Mayo Clinic Proceedings, vol. 86, no. 1, pp. 50–60, 2011. View at: Publisher Site | Google Scholar
  31. D. A. Jolliffe, C. J. Griffiths, and A. R. Martineau, “Vitamin D in the prevention of acute respiratory infection: systematic review of clinical studies,” Journal of Steroid Biochemistry and Molecular Biology, vol. 136, no. 1, pp. 321–329, 2013. View at: Publisher Site | Google Scholar
  32. K. E. Nnoaham and A. Clarke, “Low serum vitamin D levels and tuberculosis: a systematic review and meta-analysis,” International Journal of Epidemiology, vol. 37, no. 1, pp. 113–119, 2008. View at: Publisher Site | Google Scholar
  33. M. E. Belderbos, M. L. Houben, B. Wilbrink et al., “Cord blood vitamin D deficiency is associated with respiratory syncytial virus bronchiolitis,” Pediatrics, vol. 127, no. 6, pp. e1513–e1520, 2011. View at: Publisher Site | Google Scholar
  34. K. M. Kunisaki, D. E. Niewoehner, and J. E. Connett, “Vitamin D levels and risk of acute exacerbations of chronic obstructive pulmonary disease: A prospective cohort study,” American Journal of Respiratory and Critical Care Medicine, vol. 185, no. 3, pp. 286–290, 2012. View at: Publisher Site | Google Scholar
  35. A. Hossein-Nezhad and M. F. Holick, “Vitamin D for health: a global perspective,” Mayo Clinic Proceedings, vol. 88, no. 7, pp. 720–755, 2013. View at: Publisher Site | Google Scholar
  36. V. Wayse, A. Yousafzai, K. Mogale, and S. Filteau, “Association of subclinical vitamin D deficiency with severe acute lower respiratory infection in Indian children under 5 y,” European Journal of Clinical Nutrition, vol. 58, no. 4, pp. 563–567, 2004. View at: Publisher Site | Google Scholar
  37. I. Laaksi, J.-P. Ruohola, P. Tuohimaa et al., “An association of serum vitamin D concentrations < 40 nmol/L with acute respiratory tract infection in young Finnish men,” The American Journal of Clinical Nutrition, vol. 86, no. 3, pp. 714–717, 2007. View at: Google Scholar
  38. W. B. Hall, A. A. Sparks, and R. M. Aris, “Vitamin D deficiency in cystic fibrosis,” International Journal of Endocrinology, vol. 2010, Article ID 218691, 9 pages, 2010. View at: Publisher Site | Google Scholar
  39. L. A. McCauley, W. Thomas, T. A. Laguna, W. E. Regelmann, A. Moran, and L. E. Polgreen, “Vitamin D deficiency is associated with pulmonary exacerbations in children with cystic fibrosis,” Annals of the American Thoracic Society, vol. 11, no. 2, pp. 198–204, 2014. View at: Publisher Site | Google Scholar
  40. J. Xia, L. Shi, L. Zhao, and F. Xu, “Impact of vitamin D supplementation on the outcome of tuberculosis treatment: a systematic review and meta-analysis of randomized controlled trials,” Chinese Medical Journal, vol. 127, pp. 3127–3134, 2014. View at: Google Scholar
  41. P. Li, X. Xu, E. Cao et al., “Vitamin D deficiency causes defective resistance to Aspergillus fumigatus in mice via aggravated and sustained inflammation,” PLoS ONE, vol. 9, no. 6, Article ID e99805, 2014. View at: Publisher Site | Google Scholar
  42. V. Aimanianda, J. Bayry, S. Bozza et al., “Surface hydrophobin prevents immune recognition of airborne fungal spores,” Nature, vol. 460, no. 7259, pp. 1117–1121, 2009. View at: Publisher Site | Google Scholar
  43. C. Beisswenger, C. Hess, and R. Bals, “Aspergillus fumigatus conidia induce interferon-beta signalling in respiratory epithelial cells,” European Respiratory Journal, vol. 39, no. 2, pp. 411–418, 2012. View at: Publisher Site | Google Scholar
  44. A. F. Gombart, “The vitamin D-antimicrobial peptide pathway and its role in protection against infection,” Future Microbiology, vol. 4, no. 9, pp. 1151–1165, 2009. View at: Publisher Site | Google Scholar
  45. J. H. White, “Vitamin D as an inducer of cathelicidin antimicrobial peptide expression: past, present and future,” Journal of Steroid Biochemistry and Molecular Biology, vol. 121, no. 1-2, pp. 234–238, 2010. View at: Publisher Site | Google Scholar
  46. M.-L. Xue, H. Zhu, A. Thakur, and M. Willcox, “1α,25-Dihydroxyvitamin D3 inhibits pro-inflammatory cytokine and chemokine expression in human corneal epithelial cells colonized with Pseudomonas aeruginosa,” Immunology and Cell Biology, vol. 80, no. 4, pp. 340–345, 2002. View at: Publisher Site | Google Scholar
  47. Y. Zhang, D. Y. M. Leung, B. N. Richers et al., “Vitamin D inhibits monocyte/macrophage proinflammatory cytokine production by targeting MAPK phosphatase-1,” The Journal of Immunology, vol. 188, no. 5, pp. 2127–2135, 2012. View at: Publisher Site | Google Scholar
  48. A.-L. Khoo, L. Y. A. Chai, H. J. P. M. Koenen et al., “Vitamin D3 down-regulates proinflammatory cytokine response to Mycobacterium tuberculosis through pattern recognition receptors while inducing protective cathelicidin production,” Cytokine, vol. 55, no. 2, pp. 294–300, 2011. View at: Publisher Site | Google Scholar
  49. A.-L. Khoo, L. Y. A. Chai, H. J. P. M. Koenen et al., “1,25-Dihydroxyvitamin D3 modulates cytokine production induced by Candida albicans: impact of seasonal variation of immune responses,” Journal of Infectious Diseases, vol. 203, no. 1, pp. 122–130, 2011. View at: Publisher Site | Google Scholar
  50. P. T. Liu, S. Stenger, H. Li et al., “Toll-like receptor triggering of a vitamin D-mediated human antimicrobial response,” Science, vol. 311, no. 5768, pp. 1770–1773, 2006. View at: Publisher Site | Google Scholar
  51. G. R. Campbell and S. A. Spector, “Toll-like receptor 8 ligands activate a vitamin D mediated autophagic response that inhibits human immunodeficiency virus type 1,” PLoS Pathogens, vol. 8, no. 11, Article ID e1003017, 2012. View at: Publisher Site | Google Scholar
  52. C. A. Coughlan, S. H. Chotirmall, J. Renwick et al., “The effect of Aspergillus fumigatus infection on vitamin D receptor expression in cystic fibrosis,” American Journal of Respiratory and Critical Care Medicine, vol. 186, no. 10, pp. 999–1007, 2012. View at: Publisher Site | Google Scholar
  53. C. Steele, R. R. Rapaka, A. Metz et al., “The beta-glucan receptor dectin-1 recognizes specific morphologies of Aspergillus fumigatus,” PLoS pathogens, vol. 1, no. 4, p. e42, 2005. View at: Publisher Site | Google Scholar
  54. S. Bellocchio, S. Bozza, C. Montagnoli et al., “Immunity to Aspergillus fumigatus: the basis for immunotherapy and vaccination,” Medical Mycology, vol. 43, supplement 1, pp. S181–S188, 2005. View at: Publisher Site | Google Scholar
  55. R. Amitani, G. Taylor, E.-N. Elezis et al., “Purification and characterization of factors produced by Aspergillus fumigatus which affect human ciliated respiratory epithelium,” Infection and Immunity, vol. 63, no. 9, pp. 3266–3271, 1995. View at: Google Scholar
  56. J. M. Kahlenberg and M. J. Kaplan, “Little peptide, big effects: the role of LL-37 in inflammation and autoimmune disease,” The Journal of Immunology, vol. 191, no. 10, pp. 4895–4901, 2013. View at: Publisher Site | Google Scholar
  57. T. Tecle, S. Tripathi, and K. L. Hartshorn, “Defensins and cathelicidins in lung immunity,” Innate Immunity, vol. 16, no. 3, pp. 151–159, 2010. View at: Publisher Site | Google Scholar
  58. D. Yang, O. Chertov, S. N. Bykovskaia et al., “β-defensins: linking innate and adaptive immunity through dendritic and T cell CCR6,” Science, vol. 286, no. 5439, pp. 525–528, 1999. View at: Publisher Site | Google Scholar
  59. A. Soruri, J. Grigat, U. Forssmann, J. Riggert, and J. Zwirner, “beta-defensins chemoattract macrophages and mast cells but not lymphocytes and dendritic cells: CCR6 is not involved,” European Journal of Immunology, vol. 37, no. 9, pp. 2474–2486, 2007. View at: Publisher Site | Google Scholar
  60. N. Funderburg, M. M. Lederman, Z. Feng et al., “Human β-defensin-3 activates professional antigen-presenting cells via Toll-like receptors 1 and 2,” Proceedings of the National Academy of Sciences of the United States of America, vol. 104, no. 47, pp. 18631–18635, 2007. View at: Google Scholar
  61. J. R. Tisoncik, M. J. Korth, C. P. Simmons, J. Farrar, T. R. Martin, and M. G. Katze, “Into the eye of the cytokine storm,” Microbiology and Molecular Biology Reviews, vol. 76, no. 1, pp. 16–32, 2012. View at: Publisher Site | Google Scholar
  62. L. Romani, “Immunity to fungal infections,” Nature Reviews Immunology, vol. 11, no. 4, pp. 275–288, 2011. View at: Publisher Site | Google Scholar
  63. O. Takeuchi and S. Akira, “Pattern recognition receptors and inflammation,” Cell, vol. 140, no. 6, pp. 805–820, 2010. View at: Publisher Site | Google Scholar
  64. L. Romani, F. Fallarino, A. De Luca et al., “Defective tryptophan catabolism underlies inflammation in mouse chronic granulomatous disease,” Nature, vol. 451, no. 7175, pp. 211–215, 2008. View at: Publisher Site | Google Scholar
  65. M. H. Miceli, J. Maertens, K. Buvé et al., “Immune reconstitution inflammatory syndrome in cancer patients with pulmonary aspergillosis recovering from neutropenia: proof of principle, description, and clinical and research implications,” Cancer, vol. 110, no. 1, pp. 112–120, 2007. View at: Publisher Site | Google Scholar

Copyright © 2015 Pei Li et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.


More related articles

1279 Views | 430 Downloads | 2 Citations
 PDF Download Citation Citation
 Download other formatsMore
 Order printed copiesOrder

Related articles

We are committed to sharing findings related to COVID-19 as quickly as possible. We will be providing unlimited waivers of publication charges for accepted research articles as well as case reports and case series related to COVID-19. Review articles are excluded from this waiver policy. Sign up here as a reviewer to help fast-track new submissions.