Vitamin D3 down-regulates monocyte TLR expression and triggers hyporesponsiveness to pathogen-associated molecular patterns



Toll-like receptors (TLR) represent an ancient front-line defence system that enables the host organism to sense the presence of microbial components within minutes. As inducers of inflammation, TLR act as important triggers of distinct entities such as sepsis or autoimmune disease exacerbation. We report here that vitamin D3 [1α,25-dihydroxycholecalciferol, 1,25(OH)2D3] suppresses the expression of TLR2 and TLR4 protein and mRNA in human monocytes in a time- and dose-dependent fashion. Despite 1,25(OH)2D3-induced up-regulation of CD14, challenge of human monocytes with either LPS or lipoteichoic acid resulted in impaired TNF-α and procoagulatory tissue factor (CD142) production, emphasizing the critical role of TLR in the induction of inflammation. Moreover, reduced TLR levels in 1,25(OH)2D3-treated phagocytes were accompanied by impaired NF-κB/RelA translocation to the nucleus and by reduced p38 and p42/44 (extracellular signal-regulated kinase 1/2) phosphorylation upon TLR-ligand engagement. Both TLR down-regulation and CD14 up-regulation were substantially inhibited by the vitamin D receptor (VDR) antagonist ZK 159222, indicating that the immunomodulatory effect of 1,25(OH)2D3 on innate immunity receptors requires VDR transcription factor activation. Our data provide strong evidence that 1,25(OH)2D3 primes monocytes to respond less effectively to bacterial cell wall components in a VDR-dependent mechanism, most likely due to decreased levels of TLR2 and TLR4.


disseminated intravascular coagulation


extracellular signal-regulated kinases


Hanks’ balanced salt solution


lipoteichoic acid


mean fluorescence intensity




pathogen-associated molecular patterns


propidium iodide


stress-activated protein kinase


tissue factor


vitamin D receptor


Toll-like receptors (TLR) are innate immune pattern recognition receptors that enable vertebrates to deal quickly and efficiently with invading foreign microorganisms such as bacteria, virus or fungi 1, 2. At present, ten human germ-line-encoded members of the TLR family have been identified 3. As members of an evolutionary ancient first-line defence system, TLR detect the presence of conserved pathogen-associated molecular patterns (PAMP), which are produced by microorganisms. While lipopolysaccharide (LPS) signals mainly through TLR4 4, TLR2 binds peptidoglycan, lipoteichoic acid (LTA), yeast-particle zymosan and the glycosylphosphatidylinositol anchor of Trypanosoma cruzi 5, 6. Moreover, binding of LPS is facilitated through the lipid-binding, glycosylphosphatidylinositol-anchored membrane protein CD14 7, 8.

When TLR sense the presence of intruders, they engage an intracellular downstream signaling pathway that is very similar in all members of the TLR/IL-1R superfamily, including the proximal adapter molecule myeloid differentiation factor 88 (MyD88) 9 and the activation of various downstream MAPK [p38, p42/p44 (extracellular signal-regulated kinases 1/2, ERK1/2), c-Jun N-terminal kinases], and most notably leading to nuclear translocation of several transcription factors of the NF-κB/Rel and activator protein-1 families 10. Binding of these transcriptional factors to specific DNA binding sites culminates in transcriptional activation of pro-inflammatory genes such as TNF-α, IL-8, tissue factor (TF) and numerous other effectors of the innate immune response 11.

TLR play a major role in the initiation of protective immune responses; however, the extensive release of TLR-triggered pro-inflammatory mediators may harm the host organism as comes clinically overt in cases of sepsis or autoimmune disease disorders. LPS-induced expression of TF (CD142) in activated monocytes triggers thrombosis and may culminate in disseminated intravascular coagulation (DIC) 12. Moreover, TNF-α per se is known to be a potent inducer of TF expression 13. The occurrence of DIC as a secondary complication of sepsis with gram-negative bacteria is associated with poor prognosis and is an independent predictor of mortality 14. Monocytes and macrophages constitutively express high levels of TLR2 and TLR4, and they are important participants in the first strike against intruders of either gram-positive or gram-negative origin 15.

It is well known that the most active metabolite of vitamin D3, 1α,25-dihydroxycholecalciferol [1,25(OH)2D3] or calcitriol, influences the expression of several genes whose products are involved in calcium homeostasis, cell differentiation and proliferation. Expression of these genes is mediated by the nuclear vitamin D receptor (VDR) 16, which belongs to the steroid/retinoid receptor superfamily and acts as a zinc-finger nuclear transcription factor. Upon ligand activation VDR undergoes a conformational change and binds specific nucleotide sequences (vitamin D response elements) in vitamin D target genes, resulting in activation or repression of gene expression. Since VDR is expressed in several cells of the immune system, including T lymphocytes and antigen-presenting cells (APC), there is growing evidence that 1,25(OH)2D3 is a potent modulator of the immune system 17. Previously, it has been shown in an animal model that administration of 1,25(OH)2D3 prevents the development of LPS-induced DIC 18. Based upon this observation and taking into consideration that the impact of 1,25(OH)2D3 on the system of bacterial pathogen recognition is poorly understood, we investigated the effects of 1,25(OH)2D3 on the expression of TLR2 and TLR4 at the protein and mRNA level in human monocytes, and analyzed the functional consequences.


Vitamin D3 decreases TLR2 and TLR4 protein and mRNA in human monocytes in a time- and dose-dependent fashion

To assess the influence of 1,25(OH)2D3 on TLR2, TLR4 and CD14 protein expression, human monocytes were exposed to 10–7 M 1,25(OH)2D3 in a time course of 12, 24, 48 and 72 h. Cell surface expression of TLR2, TLR4 and CD14 was determined by flow cytometry using specific mAb (My4-FITC, HTA125-PE, TL2.1-PE). In Fig. 1A, baseline levels of TLR and CD14 in cells incubated with medium alone were set to 1 at each time point and fold changes followed after 1,25(OH)2D3 treatment were calculated. As can be seen, 1,25(OH)2D3 markedly increased CD14 levels in a time-dependent manner peaking at a 3.9±0.1-fold up-regulation after 72 h (p<0.001). In contrast to CD14, however, the expression of TLR2 and TLR4 protein was diminished. A significant effect was initially observed at 12 h and reached a maximum after 72 h (2.6±0.07-fold reduction for TLR2; 4.8±0.03-fold reduction for TLR4). While TLR2 baseline expression of untreated cells remained stable during the 72-h time period, we observed up-regulation of TLR4. In control experiments 0.0075% ethanol (representing the ethanol concentration of 10–7 M 1,25(OH)2D3 did not influence TLR or CD14 antigen expression (not shown).

Figure 1.

1,25(OH)2D3 down-regulates TLR2 and TLR4 in a time- and dose-dependent manner. (A) Monocytes were cultured in presence or absence of 10–7 M 1,25(OH)2D3 in a time course from 12, 24, 48 to 72 h. Positive percentages of cells expressing TLR2, TLR4 and CD14 were determined by flow cytometry. Changes over time were calculated as compared to TLR expression of untreated cells, which was set to 1. Data represent means ± SD; *p<0.05, ***p<0.001. (B) Down-regulation of TLR mRNA after incubation with 1,25(OH)2D3. Quantitative RT-PCR for human TLR2 and TLR4 was conducted with human monocytes and normalized to β-actin gene expression. Cells were incubated with 10–7 M 1,25(OH)2D3 for 3, 6 and 9 h or with medium alone. Data show results from four independent experiments and are expressed as fold changes. Error bars show means ± SD; statistically significant changes of 1,25(OH)2D3-treated monocytes vs. untreated cells: *p<0.05, **p<0.01 as calculated by one-way ANOVA. (C) Effect of 1,25(OH)2D3 on TLR2, TLR4 and CD14 expression is dose-dependent. Human monocytes were incubated in absence or in presence of increasing amounts of 1,25(OH)2D3 ranging from 10–11–10–7 M for 48 h. The MFI of monocyte TLR expression was measured by flow cytometry. Data from four individual experiments represent means ± SD; *p<0.05, **p<0.01 vs. medium alone. (D) Representative two-dimensional dot plot histograms of one from three individually analyzed apparently healthy volunteers show both reduced TLR and increased CD14 expression on monocytes after single administration of 10–7 M 1,25(OH)2D3 which peaked after 72 h. We further observed an increase of TLR4 expression in untreated cells after 72 h.

To assess TLR2 and TLR4 gene expression profiles in response to 1,25(OH)2D3 we further performed quantitative RT-PCR (Fig. 1B). Levels of TLR2 and TLR4 mRNA were significantly (p<0.05) down-regulated in monocytes treated for 3, 6 or 9 h with 10–7 M 1,25(OH)2D3 when compared to cells cultured in medium alone.

We next treated cells with increasing concentrations of 1,25(OH)2D3 ranging 10–11–10–7 M for 48 h. Our results show that the effect of 1,25(OH)2D3 on TLR2 and TLR4 is concentration-dependent (Fig. 1C). Down-regulation of TLR2 and TLR4 was barely detectable at doses of 10–11 M, and reached the lowest levels at a concentration of 10–7 M. In contrast to TLR2 and TLR4, CD14 was inversely regulated with highest fluorescence intensities at a concentration of 10–7 M (data upon request). High concentrations of 1,25(OH)2D3 (10–7 M) did not influence cell viability at any time point as confirmed by annexin V/propidium iodide (PI) staining (Fig. 2B, C).

Figure 2.

(A) Reduced SAPK phosphorylation upon TLR-ligand engagement in monocytes treated with 1,25(OH)2D3. Negatively depleted CD14+ cells pre-incubated with or without 1,25(OH)2D3 were stimulated for 10 min with either 10 ng LPS or 10 µg LTA. Cells were fixed, permeabilized and phospho-specific mAb were added to determine p38 and p42/44 activation. Colored histograms show cells treated with medium only while open histograms show activated cells stimulated with LPS or LTA. Incubation of 1,25(OH)2D3 prior to TLR engagement (10–7 M, shaded histograms) resulted in reduced phosphorylation of p38 and ERK1/2. Histograms shown are representative of three individually performed experiments. (B, C) Influence of 1,25(OH)2D3 on cell viability. Monocytes were treated with or without 10–7 M 1,25(OH)2D3, and annexin V-FITC/PI staining was performed at each time point (n=3). No difference was observed between untreated cells and cells that were incubated with 1,25(OH)2D3. Representative dot plot diagram depicted shows the impact of 1,25(OH)2D3 on monocytes viability after 48 h.

TLR-induced p38 and p42/44 phosphorylation is reduced in 1,25(OH)2D3-treated monocytes

Being that there is a divergent regulation of CD14 and TLR2 and TLR4, it is of interest to investigate TLR downstream signaling. Using phospho-specific antibodies, we analyzed p38 and p42/44 (ERK1/2) stress-activated protein kinase (SAPK) signaling in purified monocytes stimulated with ligands for TLR2 and TLR4, respectively. Monocytes pretreated with 1,25(OH)2D3 that were stimulated either with LPS or LTA exhibited lower levels of phosporylated p38 and p42/42 (ERK1/2) than untreated cells (Fig. 2A), suggesting that TLR down-regulation by vitamin D similarly impacts subsequent SAPK phosphorylation.

Decreased cytokine production and TF expression in 1,25(OH)2D3-treated monocytes upon TLR ligand induction

Since both CD14 and TLR4 are involved in LPS recognition but inversely regulated by 1,25(OH)2D3 (Fig. 1A, D), we enquired for the net effect as far as synthesis of inflammatory TNF-α and procoagulatory TF in monocytes is concerned. Monocytes were pretreated for 48 h with increasing concentrations of 1,25(OH)2D3 (10–11–10–7 M) and afterwards incubated with either 10 ng LPS or 10 µg LTA for 4 h. TNF-α was determined using two methods, intracellular cytokine staining of CD14+ cells (Fig. 3A) and measurement of cytokine release in supernatants (Fig. 3B). In accordance with TLR2 and TLR4 down-regulation, TNF-α synthesis upon LPS and LTA challenge was decreased in 1,25(OH)2D3-treated monocytes in a dose-dependent manner with lowest TNF-α levels at the highest 1,25(OH)2D3 concentration (p<0.01). No significant change was observed at a concentration of 10–11 M.

Figure 3.

(A) Decreased intracellular TNF-α production in 1,25(OH)2D3-treated CD14+ cells upon TLR ligand induction. Monocytes were incubated for 48 h in absence or presence of various 1,25(OH)2D3 concentrations ranging from 10–7–10–11 M and afterwards stimulated for 4 h with 10 ng/mL LPS or 10 µg/mL LTA, respectively. Cells were double-stained with CD14-FITC, TNF-α-PE or with corresponding isotype antibodies and analyzed by flow cytometry. Two-color dot plots are from one representative experiment of three. Values in gate refer to the percentage of TNF-α+CD14+ cells. (B) TLR ligand triggered TNF-α secretion in the presence or absence of vitamin D3 in supernatants. Negatively selected monocytes were incubated with 1,25(OH)2D3, and stimulated with either 10 ng/mL LPS or 10 µg/mL LTA for 4 h. Supernatants were collected and analyzed for TNF-α concentrations by chemiluminescence. (C) Diminished TNF-α expression in monocytes pre-incubated with either anti-TLR2 or anti-TLR4 antibodies. Human monocytes (1×106/mL) were treated with either 10 ng/mL LPS or 10 µg/mL LTA for 4 h. Cells in the anti-TLR4- or anti-TLR2-treated sample were pre-incubated with 20 µg/mL HTA125 or 20 µg/mL TL2.1 for 1 h prior to addition of LPS or LTA. Intracellular TNF-α production (MFI) was measured as described in Materials and methods. Both clones HTA125 (TLR4) and TL2.1 (TLR2) substantially inhibited TNF-α expression. Bars show means ± SD of three independent experiments. Data represent the means ± SD (n=3); *p<0.05, **p<0.01 vs. control as determined by Tukey-Kramer.

In order to test the TLR specificity of LPS and LTA, we pre-incubated monocytes with either anti-TLR2 or anti-TLR4 sterile functional-grade antibodies. As shown in Fig. 3C, both clones HTA125 (TLR4) and TL2.1 (TLR2) substantially inhibited TNF-α expression, whereas incubation with the mAb alone had no stimulatory effects.

Similar to the results observed for TNF-α analysis, TF protein expression (Fig. 4A) and functional activity (Fig. 4B) were decreased in a dose-dependent fashion as determined by flow cytometry (percentage of CD142+CD14+ cells) and by one-stage clotting assays showing most prominent effects at a concentration of 10–7 M (46.1±6.1 pM vs. 119.2±46.1 pM; p<0.01, n=5) and remaining at baseline levels at 10–11 M.

Figure 4.

(A) Counteracting effects of 1,25(OH)2D3 on up-regulation of TF by LPS and LTA in phagocytes. Monocytes were incubated with increasing concentrations of 1,25(OH)2D3 and afterwards stimulated with 10 ng LPS or 10 µg LTA for 4 h. Levels of TF on CD14+ cells were measured by flow cytometry. Data from three independent experiments represent the means ± SD; *p<0.05, **p<0.01 vs. control as determined by Tukey-Kramer. (B) Reduced TF activity on human monocytes activated with 10 ng/mL E. coli LPS for 4 h. Prior to stimulation, cells were cultured for 48 h in absence or presence of various concentrations of 1,25(OH)2D3. Total TF activity of cell lysates was quantified using a standard curve generated with recombinant human TF. Results from five independent experiments represent the means ± SD; **p<0.01 vs. cells stimulated with LPS alone.

Vitamin D3 blocks LPS/LTA-induced NF-κB/RelA nuclear translocation in human monocytes

We analyzed whether 1,25(OH)2D3 influences RelA/p65 translocation to the nucleus upon stimulation with LPS or LTA. Human monocytes were incubated with vitamin D3 (10–7 M) for 48 h before stimulation with 10 ng LPS or 10 µg LTA for 30 minutes. Both TLR ligands strongly induced nuclear translocation of p65 (Fig. 5A). In 1,25(OH)2D3-treated monocytes, however, p65 remained largely localized in the cytoplasm, indicating that reduced TLR levels are accompanied by decreased p65 activation.

Figure 5.

(A) Diminished RelA translocation in 1,25(OH)2D3-treated monocytes after TLR challenge. Negatively selected CD14+ monocytes were incubated for 48 h in absence or presence of 10–7 M 1,25(OH)2D3 and stimulated with either 10 ng/mL LPS or 10 µg/mL LTA for 30 min. Cells were attached onto microscope slides, fixed and NF-κB/RelA (p65) and PI immunofluorescence staining was performed. Individual and merged pictures out of two independent experiments are shown. No background staining was observed in slides incubated with corresponding isotype antibodies alone (not shown). Original magnifications, ×100. (B, C) TLR down-regulation is blocked by the VDR antagonist ZK 159222. Two- and one-dimensional overlay plots of two separately performed experiments are presented. While 1,25(OH)2D3-treated monocytes (red cells) show divergent expression levels of TLR and CD14, cells incubated with both ZK 159222 and 1,25(OH)2D3 (orange) and untreated cells (green) are similar in CD14 and TLR expression. Concentrations for 1,25(OH)2D3 and ZK 159222 were 10–7 M and 10–6 M, respectively. ZK 159222 was added 1 h before 1,25(OH)2D3 administration and overall incubation time was 48 h.

Down-regulation of TLR2 and TLR4 is VDR-dependent

In order to get a mechanistic picture of the effects of 1,25(OH)2D3, we went on to investigate whether 1,25(OH)2D3-induced down-regulation of TLR2 and TLR4 requires involvement of the VDR transcription factor. Indeed, the VDR antagonist ZK 159222 reversed both 1,25(OH)2D3-induced TLR down-regulation and CD14 up-regulation (Fig. 5B).


Our data demonstrate a novel effect of the secosteroid hormone vitamin D3 on innate immune receptors that sense microbial products. We show for the first time that 1,25(OH)2D3 down-regulates TLR2 and TLR4 expression on human monocytes at protein and mRNA level in a time- and concentration-dependent manner. Down-regulation induced by 1,25(OH)2D3 was most prominent after 72 h cell incubation with 10–7 M, and still significant at 10–9 M. Concentrations of 10–10–10–11 M, which are similar to those observed physiologically in plasma, neither influenced TLR2 and TLR4 expression nor changed expression profiles of TNF-α and TF after TLR-ligand interaction.

The results presented suggest that the impaired inflammatory response to bacterial PAMP in monocytes treated with 1,25(OH)2D3 at concentrations of 10–9 M and above is, at least in part, due to TLR down-regulation. Since TLR are key components in pathogen recognition and crucial mediators in the early inflammatory response to foreign microorganisms, down-regulation of TLR2 and TLR4 by 1,25(OH)2D3 clearly represents an important and novel immunomodulating effect. Our observation is in accordance with previous studies by us and others showing that 1,25(OH)2D3 exerts a number of immunosuppressive effects on monocytes/macrophages, such as diminished expression of MHC class II, CD40, CD64, CD32, CD89, CD23, CD11b, CD11c, CD71, and impaired synthesis of IL-1β, IL-6, IL-12, TNF-α and M-CSF 19, 20. Moreover, our data confirm the results of a number of previously published articles indicating that 1,25(OH)2D3 dose-dependently suppresses LPS-induced TNF-α production 2123.

Proximal LPS signaling by TLR4 is facilitated by several accessory proteins including CD14, MD2 and LPS-binding protein. Interestingly, 1,25(OH)2D3 has divergent effects on CD14 and TLR4, both of which are involved in LPS recognition. Moreover, CD14 has been shown to be a coworker of TLR2 as well 24. While CD14 expression is strongly enhanced by 1,25(OH)2D3, a fact that is well established 25, TLR2 and TLR4 are, to our surprise, inversely regulated. Indeed, 1,25(OH)2D3 treatment prior to LPS exposure leads to reduced SAPK phosphorylation and release of pro-inflammatory mediators, suggesting a minor role of CD14 in innate immune response and emphasizing the unique role of TLR in LPS-mediated intracellular signaling and cytokine production. As CD14 lacks a trans-membrane domain, it is reasonable to hypothesize that TLR expression and function may ultimately predict the outcome of innate immunity response.

The MyD88-dependent TLR signal transduction pathway ultimately involves NF-κB translocation and binding to specific κB binding sites in respective target genes. We suggest that the impaired NF-κB/RelA activation in 1,25(OH)2D3-treated cells upon TLR-ligand binding is likely due to TLR down-regulation since IL-1β-induced p65 and p50 translocation is not inhibited by 1,25(OH)2D3 or dexamethasone or by the combination of both 26. Yet, other mechanisms may contribute to the ability of 1,25(OH)2D3 to inhibit NF-κB/RelA activity.

Most of the pleiotropic actions of 1,25(OH)2D3 are mediated through interaction with the VDR ligand-binding domain, thus allowing VDR for conformational change. Our data suggest that both CD14 up-regulation and TLR down-regulation require functional VDR activity, as the VDR antagonist ZK 159222 – a vitamin D analogue that keeps VDR with its bulky side chain in an antagonistic conformation – substantially abrogated the effects of 1,25(OH)2D3. Preferentially, VDR acts as a heterodimer in concert with retinoid X receptor, another nuclear receptor superfamily member and, of interest, all-trans retinoic acid has been shown to down-regulate TLR2, but not CD14, TLR1 and TLR4 27.

In the attempt to prevent infection, the innate immune response initiated by microbial sensors is, ironically, capable of harming the host. For example, the severe systemic disorder of septic shock caused by an extensive release of LPS in gram-negative infection could serve as potential indication for agents that dampen the TLR-mediated pro-inflammatory response. In the experimental setting of dilated autoimmune cardiomyopathy, Eriksson et al.2 demonstrated that induction of autoimmunity requires autoantigen presentation, CD40 costimulation and TLR activation, as TLR4-deficient mice fail to develop autoimmune heart disease in an endotoxin environment. We propose that the mechanism underlying the protective role of 1,25(OH)2D3 in Th1-mediated autoimmune disease involves down-regulation of antigen-presenting molecules, costimulatory CD40 and, most importantly, down-regulation of TLR on APC. Interestingly, Hausmann and colleagues 28 reported that TLR2 and TLR4 levels in intestinal macrophages are up-regulated in inflamed areas of patients suffering inflammatory bowel disease, suggesting TLR to contribute to the inflammatory process in intestinal macrophages. Since macrophages of the normal mucosa do not express any TLR, 1,25(OH)2D3 or vitamin D analogues that lack systemic uptake might be beneficial in the treatment of autoimmune enteritis 29.

Beside its attractive immunosuppressive characteristics, however, 1,25(OH)2D3 has dose-limiting hypercalcemic effects. Thus, the systemic clinical use of 1,25(OH)2D3 is limited due to its side effects associated with unbalanced calcium homeostasis. In the last years, therefore, great effort has been made to synthesize vitamin D3 analogues that share immunosuppressive and anticoagulant properties but lack hypercalcemic activity.

Our work sheds light on the effects of 1,25(OH)2D3 on the major bacterial pattern recognition receptors TLR2 and TLR4. Other TLR recognizing viral compounds might likely be influenced by 1,25(OH)2D3 as well. Additional studies will determine the degree of modulation among other TLR by 1,25(OH)2D3. In conclusion, our results provide evidence that 1,25(OH)2D3 primes mononuclear phagocytes to respond less effectively to bacterial PAMP, most likely via modulation of TLR.

Materials and methods

Cell isolation and culture

Heparinized whole blood was drawn from apparently healthy adult volunteers. Human monocytes were separated by density gradient centrifugation and by means of magnetic cell sorting (Monocyte isolation kit II; Milteny Biotec, Bergisch Gladbach, Germany). Monocyte purity of negatively selected cells was usually ∼95% as determined by flow cytometry. Cells (1×106/mL) were seeded in Teflon-coated hydrophobic culture plates (PetriPerm hydrophobic; Vivascience, Vienna, Austria) and cultured in RPMI 1640 supplemented with 10% FCS and 2 mM glutamine in the presence or absence of indicated 1,25(OH)2D3 concentrations.

Vitamin D3 and innate immunity ligands

1,25(OH)2D3 was kindly provided by Roche (Basel, Switzerland). A stock solution of 10–3 M was prepared in 75% ethanol, overlaid with N2 gas and stored in small sterile aliquots at –80°C. Effects on cell viability by high concentrations of 1,25(OH)2D3 (10–7 M) was assessed by annexin V-FITC/PI (Trevigen, Gaithersburg, MD) staining. The 25-carboxylic ester vitamin D analog ZK 159222 30 was synthesized at the Medicinal Chemistry Department at Schering AG and a stock solution of 10–2 M was dissolved in ethanol. Further dilutions were made in RPMI 1640 shortly before administration to cell culture. LPS purified from Escherichia coli R515 was obtained from Alexis Cooperation (Lausen, Switzerland) and proven by the manufacturer not to activate other TLR as determined with splenocytes and macrophages from TLR4-deficient mice. Staphylococcus aureus-derived LTA (Sigma-Aldrich) was diluted in sterile distilled pyrogen-free water and was stored according to manufacturer's suggestions.

Flow cytometry

PE-labeled anti-human TLR4 (HTA125, mouse IgG2a), TLR2 (TL2.1, mouse IgG2a) as well as corresponding isotype antibodies were purchased from eBioscience (San Diego, CA). Anti-CD14-mAb (My4-FITC; Beckman Coulter, Fullerton, CA) was used to define the monocyte population. Anti-human CD142 (TF, HTF1-PE) was purchased from Pharmingen (BD Biosciences, San Diego, CA). Antibody incubation was performed on ice for 30 min. Monocytes were washed twice with Hanks’ balanced salt solution (HBSS; Bio Whittaker, Verviers, Belgium) containing 0.1% NaN3, 0.3% BSA, and a total of 2×104 CD14+ cells were analyzed by flow cytometry on a Coulter Epics XL equipped with EXPO32 software.

Intracellular TNF-α cytokine staining of CD14+ cells following after 4-h incubation with LPS or LTA was performed with a PE-conjugated anti-human TNF-α mAb (mAb11, mouse IgG1). All reagents needed for detection of intracellular cytokines were purchased from Pharmingen and intracellular staining was performed according to manufacturer's suggestions. TLR2 and TLR4 blocking experiments were performed using anti-TLR2 (clone TL2.1) or anti-TLR4 (clone HTA125) antibodies. Cells in the anti-TLR4- or anti-TLR2-treated sample were pre-incubated with 20 µg/mL HTA125 or 20 µg/mL TL2.1 (functional grade, endotoxin-tested; eBioscience) for 1 h prior to addition of LPS or LTA. Intracellular TNF-α production (mean fluorescence intensity, MFI) was measured as described above.

Intracellular phospho-specific analysis of SAPK was undertaken as described previously 31, 32. Briefly, negatively selected monocytes were either treated with medium (deficient RPMI 1640) or stimulated with LPS (10 ng/mL) or LTA (10 µg/mL) for 10 min at 37°C. Fixation of phospho-epitopes was done using BD Phosflow Fix Buffer for 10 min at 37°C, followed by permeabilization with ice-cold 90% methanol for 30 min on ice. Cells were washed twice with staining buffer and the following phospho-specific mAb were added for 30 min at room temperature: PE-p38 (pT180/pY182, mouse IgG1, clone 36), PE-ERK1/2 (pT202/pY204, mouse IgG1), both BD Pharmingen, and CD14-FITC (mouse IgG2a, clone UCH-M1; Santa Cruz Biotechnology, Santa Cruz, CA).

Real-time PCR

Monocytes were incubated with or without 1,25(OH)2D3 and at indicated time points, cells were washed with HBSS, lysed in 1 mL Trizol® reagent (Gibco BRL, Life Technologies, Paisley, UK) and total RNA was prepared according to manufacturer's instruction. Purity and quantity of the extract was determined by ultraviolet absorption and gel electrophoresis. All reagents and devices used for real-time PCR were obtained from Applied Biosystems (Foster City, CA). A total of 200 ng RNA was reverse-transcribed to cDNA using TAQ-Man reverse transcription agents.

Quantitative RT-PCR of target cDNA was conducted for TLR2, TLR4 and normalized to β-actin gene expression. All primers and probes (TAQ-Man Gene Expression Assays®) were obtained from Applied Biosystems. Experiments were performed in 96-well plates in triplicates using TAQ-Man Universal Master Mix. Real-time PCR amplification was performed on a Gene Amp® 5700 Sequence Detection System. PCR conditions were 50°C for 2 min, 95°C for 10 min, then 45 cycles at 95°C for 15 s, and 60°C for 1 min.

Measurement of TNF-α in supernatants

Cell culture supernatants were collected after indicated time points, centrifuged to remove cellular components, and stored at –80°C. TNF-α concentrations were measured with an Immulite2000 system (LKNF1) from DPC Biermann (Bad Nauheim, Germany). The calibration range is 5–1000 pg/mL, with an analytical sensitivity of 1.7 pg/mL for TNF-α.

TF induction assay

One-stage clotting assays were performed with isolated human monocytes as described 33. Briefly, cells were treated with different concentrations of calcitriol for 48 h and stimulated with 10 ng LPS for 4 h. Cells were then harvested, washed twice with sterile HBSS and incubated with 1 mL of lysis buffer (25 mM Tris in distilled water, pH 8.0) for 10 min at 37°C, followed by freezing at –80°C for 30 min. Cell lysates were then thawed at 37°C and centrifuged at 2000×g for 5 min. Cell pellet was resuspended in 120 μL of clotting buffer (50 nM Tris, 100 nM NaCl, 0.1% BSA, pH 7.5) and incubated with 100 μL factor VIII-deficient plasma (George-King Biomedical, Overland Park, KS) for 110 s. Clotting reaction was catalyzed by adding 100 μl of CaCl2 (30 nM) and time of clot formation was measured. TF activity was quantified by reference to standard curves constructed with lipidated human recombinant TF (American Diagnostica, Stamford, CT).


For NF-κB/RelA staining, cells were incubated with or without 1,25(OH)2D3 and stimulated for 30 min with either 10 ng LPS or 10 µg LTA. Cells were spun down onto microscope slides and immediately fixed with 4% paraformaldehyde (20 min, room temperature). Monocytes were permeabilized with 0.4% Triton X for 5 min and primary antibody against p65 (polyclonal rabbit IgG, sc-109X; Santa Cruz Biotechnology) was added (1:200 in PBS with 1.5% BSA, 120 min, 4°C). Cells were washed in PBS and incubated with Alexa Fluor 488 goat anti-rabbit IgG (1:200; Molecular Probes, Eugene, OR) for 1 h at room temperature. Cells were washed again and nuclei were stained with PI (2.5 µg/mL, 10 min, room temperature. Samples were mounted with SlowFade Light (Molecular Probes) and sealed. Fluorescence images were taken with a Zeiss LSM 510 laser scanning confocal microscope (Zeiss, Oberkochen, Germany).

Statistical analysis

Data are presented as mean ± standard deviation. Multiple comparisons were adjusted according to Tukey-Kramer. p values are two-sided and p<0.05 was considered statistically significant.


This work was supported in part by the ‘Hygiene Fonds’ of the Medical University of Vienna. The authors are grateful to Brustoski Kim for careful reading of the manuscript and for helpful discussion. The authors would like to acknowledge Schering AG for kindly providing the substance ZK159222. The authors have no conflicting financial interests.


  1. 1


  2. 2


  3. 3


  4. 4


  5. 5