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Originally published online as doi:10.1189/jlb.0607428 on January 3, 2008

Published online before print January 3, 2008
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(Journal of Leukocyte Biology. 2008;83:912-920.)
© 2008 by Society for Leukocyte Biology

Biotin deficiency up-regulates TNF-{alpha} production in murine macrophages

Toshinobu Kuroishi*,1, Yasuo Endo*, Koji Muramoto{dagger} and Shunji Sugawara*

* Division of Oral Immunology, Department of Oral Biology, Tohoku University Graduate School of Dentistry, and
{dagger} Laboratory of Biomolecular Function, Tohoku University Graduate School of Life Sciences, Sendai, Japan

1 Correspondence: Division of Oral Immunology, Department of Oral Biology, Tohoku University Graduate School of Dentistry, 4-1 Seiryo-machi, Aoba-ku, Sendai 980-8575, Japan. E-mail: kuroishi{at}mail.tains.tohoku.ac.jp


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ABSTRACT
 
Biotin, a water-soluble vitamin of the B complex, functions as a cofactor of carboxylases that catalyze an indispensable cellular metabolism. Although significant decreases in serum biotin levels have been reported in patients with chronic inflammatory diseases, the biological roles of biotin in inflammatory responses are unclear. In this study, we investigated the effects of biotin deficiency on TNF-{alpha} production. Mice were fed a basal diet or a biotin-deficient diet for 8 weeks. Serum biotin levels were significantly lower in biotin-deficient mice than biotin-sufficient mice. After i.v. administration of LPS, serum TNF-{alpha} levels were significantly higher in biotin-deficient mice than biotin-sufficient mice. A murine macrophage-like cell line, J774.1, was cultured in a biotin-sufficient or -deficient medium for 4 weeks. Cell proliferation and biotinylation of intracellular proteins were decreased significantly in biotin-deficient cells compared with biotin-sufficient cells. Significantly higher production and mRNA expression of TNF-{alpha} were detected in biotin-deficient J774.1 cells than biotin-sufficient cells in response to LPS and even without LPS stimulation. Intracellular TNF-{alpha} expression was inhibited by actinomycin D, indicating that biotin deficiency up-regulates TNF-{alpha} production at the transcriptional level. However, the expression levels of TNF receptors, CD14, and TLR4/myeloid differentiation protein 2 complex were similar between biotin-sufficient and -deficient cells. No differences were detected in the activities of the NF-{kappa}B family or AP-1. The TNF-{alpha} induction by biotin deficiency was down-regulated by biotin supplementation in vitro and in vivo. These results indicate that biotin deficiency may up-regulate TNF-{alpha} production or that biotin excess down-regulates TNF-{alpha} production, suggesting that biotin status may influence inflammatory diseases.

Key Words: cytokine • inflammation • nutrition


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INTRODUCTION
 
Biotin is a water-soluble vitamin of the B complex found in all organisms [1 ]. Biotin functions as a cofactor of five carboxylases—pyruvate carboxylase (EC 6.4.1.1), two forms of acetyl-CoA carboxylase (ACC1 and ACC2; EC 6.4.1.2), propionyl-CoA carboxylase (PCC; EC 6.4.1.3), and methylcrotonoyl-CoA carboxylase (EC 6.4.1.4)—enzymes that catalyze the metabolism of glucose, amino acids, and fatty acids. As these biotin-dependent enzymes are indispensable for cellular metabolism, biotin starvation or deficiency is potentially lethal [2 ]. Biotin is covalently attached to specific lysine residues of carboxylases by the enzyme holocarboxylase synthetase (HCS; EC 6.3.4.10) [2 ]. During the turnover of carboxylases, biotin is released from biotinylated peptides by biotinidase (EC 3.5.1.12) and recycled in the biotinylation of new carboxylases [2 ]. The defect of this biotin cycle causes a neonatal form of life-threatening ketoacidosis and organic acidemia, known as multiple carboxylase deficiency [2 ].

In addition to this classical function as a cofactor of carboxylases, biotin is involved in various cellular events. It was reported that biotin regulates the mRNA expression of HCS and biotin-dependent carboxylases via a cGMP-dependent pathway [3 ]. Biotin regulates transcription factors, such as NF-{kappa}B, specificity protein 1 (Sp1), and Sp3, in human T cell line Jurkat cells [4 , 5 ]. Moreover, biotinylation of histones in human cells was also reported [6 ]. In a human hepatoblastoma cell line, biotin regulates the expressions of asialoglycoprotein receptor and insulin receptor at the post-transcriptional level [7 ]. These reports clearly indicated that biotin regulates the various cellular events at transcriptional and post-transcriptional levels.

Immunological effects of biotin have also been studied. In vitro biotin supplementation induces IL-2R{gamma} expression and decreases the net secretion of IL-2 from Jurkat cells [8 , 9 ]. In vivo supplementation of a pharmacologic dose of biotin decreases the proliferation rate of PBMC and the release of IL-1β and IL-2 [10 ]. Moreover, biotin deficiency changes the number and subpopulations of spleen lymphocytes and blocks thymocyte maturation in mice [11 , 12 ]. On the other hand, contradictory results have been reported, namely that lymphocyte subpopulations, mitogen-induced cytokine production, IgG responses, and NK cell activity do not differ significantly between mild to moderately biotin-deficient and biotin-sufficient rats [13 ]. Therefore, the immunological effects of biotin have not been clarified.

Moderately severe biotin deficiency causes alopecia and scaly erythematous dermatitis [11 , 14 , 15 ]. In addition to applications as a dietary supplement, biotin is prescribed for chronic dermatitis. It was reported that biotin has a therapeutic effect on pustulosis palmaris et plantaris, a type of chronic dermatitis that is restricted to the palms and soles and is related to metal allergy [16 , 17 ]. Moreover, Makino et al. [18 ] reported that serum biotin levels are significantly lower in atopic dermatitis patients than in healthy subjects. These reports suggest that biotin deficiency is involved in inflammatory diseases. However, few reports are available about the biological roles of biotin in inflammatory responses.

In this study, we investigate the in vivo effects of biotin deficiency using a mouse model of LPS-induced TNF-{alpha} production. We also investigated the in vitro effects of biotin deficiency on the production of TNF-{alpha} by the murine macrophage cell line J774.1. We showed that biotin status affects the production of TNF-{alpha}, and biotin-supplementation down-regulated it in vivo and in vitro.


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MATERIALS AND METHODS
 
Reagents
LPS from Escherichia coli O55:B5, prepared by Westphal’s method, was purchased from Difco Laboratories (Detroit, MI, USA). All other reagents were purchased from Sigma-Aldrich (St. Louis, MO, USA), unless otherwise indicated.

Mice
Female BALB/c mice (4 weeks old), obtained from the Institute for Experimental Animals of the Tohoku University Graduate School of Medicine (Sendai, Japan), were used for the experiments. Mice were divided into two groups: biotin-sufficient and -deficient groups, which received a basal diet (AIN-76, containing 0.8 mg d-biotin per kg) or a biotin-deficient (biotin-free) AIN-76 diet (Nosan Corp., Yokohama, Japan), respectively. Preliminary experiments showed that 8-week feeding led to a significant decrease in the serum biotin level in mice. For in vivo biotin supplementation, biotin-deficient mice were provided with biotin-supplemented drinking water (15 µM) for 2 weeks. The dose of biotin was based on the dose of human biotin therapy (10–40 mg/day) and drinking volume of mice (5 ml/day) [19 , 20 ]. The Ethical Board for Nonhuman Species of the Tohoku University Graduate School of Medicine approved the experimental procedure followed in this study.

Measurement of biotin in serum
Biotin in serum was purified by HPLC and measured by ELISA, as originally described by Mock [21 ]. Briefly, serum (1 ml; the serum was pooled from five mice) was ultrafiltrated using a centrifugal ultrafiltration unit (a MW cutoff of 30 kDa) at 1500 g for 90 min at 4°C. The filtrates were adjusted to pH 2.5 by 6 M HCl and then separated by reversed-phase HPLC on a Capcelpak AG120A C18 (5 µm, 4.6x250 mm, Shiseido, Tokyo, Japan) column at a flow rate of 1.0 ml/min at 40°C. Elution was carried out with solution A [0.1% trifluoroacetic acid (TFA)] and solution B (80% acetonitrile in 0.8% TFA). The linear gradient was started at solution A 100% and solution B 0% and was reached at solution A 81% and solution B 19% at 30 min and then solution A 0% and solution B 100% at 60 min. The column eluent was collected every minute in test tubes and dried by a centrifugal concentrator. After drying, each fraction was dissolved in 500 µl H2O. From preliminary experiments with commercial biotin standard, we confirmed that biotin elutes at 32 min. For ELISA, 96-well, flat-bottomed plates (Nunc-Immuno modules, MaxiSorp, Nalge-Nunc International, Rochester, NY, USA) were coated with biotinyl-BSA, synthesized with BSA and biotin N-hydroxysuccinimide ester (Pierce Biotechnology, Inc., Rockford, IL, USA), and then blocked with 0.01% (wt/vol) BSA. Samples (100 µl) and 50 µl HRP-conjugated avidin (Calbiochem, Darmstadt, Germany) were mixed and incubated for 2 h at room temperature. Then, 100 µl of the mixture was transferred to a well of a biotinyl-BSA-coated plate and incubated for 4 h at room temperature. The ELISA was developed with 3,3',5,5' tetramethylbenzidine (substrate reagent set, BD Biosciences, San Diego, CA, USA) as a substrate. The detection range of biotin was 3–10,000 nM.

Measurement of PCC activity
PCC activity was measured as originally described by Zempleni et al. [22 ]. Briefly, the liver was homogenized on ice in buffer containing 50 mM Tris-HCl (pH 7.4), 1 mM EDTA, 10 mM 2-ME, and 0.25 M sucrose and centrifuged at 14,000 g for 30 min at 4°C. The supernatant was used as a crude enzyme fraction. For PCC assay, a 5-µl tenfold-diluted crude enzyme fraction was incubated with a 95-µl reaction mixture for 5 min at room temperature. The reaction mixture consisted of 100 mM Tris-HCl (pH 8.0), 0.75 mM DTT, 6 mM MgCl2, 3.14 mM Na2-ATP, 100 mM KCl, 1% Triton X-100, 1 mM propionyl-CoA, and 3.5 mM NaH[14C]O3 (2.11 kBq/nmol). The reaction was stopped by the addition of 30 µl 1 M HClO4. After centrifugation at 1000 g for 15 min, 100 µl aliquot was transferred into a vial and dried. Finally, the sample was resolved in a scintillation cocktail, and the bound [14C]bicarbonate was quantified by a liquid scintillation counter. PCC activity was expressed in units: 1 unit = 1 nmol HCO3– fixed/min/mg protein.

Measurement of TNF-{alpha} in serum
LPS (1 µg/kg) was injected i.v. into mice. After 90 min, blood was collected and coagulated on ice for 1 h, and sera were recovered by centrifugation. The amounts of TNF-{alpha} in sera were measured with an OptEIA mouse TNF-{alpha} (Mono/Mono) set (BD Biosciences).

Biotin depletion from FCS
Biotin in FCS was depleted with immobilized avidin-agarose (Pierce Biotechnology, Inc.). Briefly, FCS (Tissue Culture Biologicals, Tulare, CA, USA) was mixed with avidin-agarose and gently stirred overnight at 4°C. After centrifugation, FCS was sterilized by filtration (0.22 µm pore size) and stored at –30°C until use. Biotin depletion was confirmed by ELISA as described above. The biotin concentration in FCS was 8.6 ± 0.25 nM, and that in biotin-deficient FCS was not detectable (less than 3 nM).

Cells and cell culture
J774.1, a murine macrophage-like cell line, was obtained from the Cell Resource Center for Biomedical Research, Institute of Development, Aging and Cancer, Tohoku University. J774.1 cells were grown in biotin-sufficient and -deficient medium. The biotin-sufficient medium was RPMI 1640 (Invitrogen Corp., Carlsbad, CA, USA) containing d-biotin (0.2 µg/ml) supplemented with 10% FCS. The biotin-deficient medium was biotin-free RPMI 1640 supplemented with 10% biotin-deficient FCS.

Western blotting
J774.1 cells were lysed in SDS-PAGE sample buffer. Cell lysates were separated by SDS-PAGE under reducing conditions. After SDS-PAGE, gel proteins were electrophoretically transferred to a polyvinylidene difluoride membrane (Bio-Rad Laboratories, Hercules, CA, USA). The blot was blocked for 1 h with 3% (wt/vol) skim milk and 0.05% Tween 20 in PBS and incubated with HRP-conjugated avidin (Calbiochem). After washing, the blot was analyzed with SuperSignal West Femto maximum sensitivity substrate (Pierce Biotechnology, Inc.) and a Chemi Imager (Alpha Innotech Corp., San Leadro, CA, USA).

[3H] TdR incorporation assay
J774.1 cells were suspended in RPMI 1640 with 10% FCS and seeded in a 96-well, flat-bottomed plate (Nalge-Nunc International) at the indicated cell numbers. The cells were incubated at 37°C for 3 h, and each well was pulsed with 1 kBq [3H] TdR (MP Biomedicals, Irvine, CA, USA) for 2 h. After the pulse, 100 µl 3% Triton X-100 was added to each well, and the cells were harvested onto a glass fiber filter. The counts of [3H] TdR per minute (cpm) were measured with a liquid scintillation counter (Packard Instrument Company, Meriden, CT, USA).

Measurement of TNF-{alpha} in culture supernatants of J774.1 cells
J774.1 cells were suspended in RPMI 1640 with 10% FCS and seeded in a 96-well flat-bottomed plate at 2 x 105 cells/200 µl/well. The cells were stimulated with LPS for 24 h at 37°C. The amount of TNF-{alpha} in the culture supernatant was measured with an OptEIA mouse TNF-{alpha} (Mono/Mono) set (BD Biosciences).

Flow cytometry
For TNF-{alpha} staining, J774.1 cells were seeded in a six-well, flat-bottomed plate (Nalge-Nunc International) at 1 x 106 cells/2 ml/well with RPMI 1640 with 10% FCS. The cells were cultured for 6 h at 37°C in the presence or absence of 1 µg/ml actinomycin D (Act-D). Then, the cells were stained with FITC-conjugated anti-mouse TNF mAb (clone MP6-XT22, BD Biosciences) for cell surface cytokine staining. For intracellular cytokine staining, the cells were incubated with a protein transport inhibitor (GolgiPlug, BD Biosciences). After fixation and permeabilization with Cytofix/Cytoperm (BD Biosciences), the cells were stained with FITC-conjugated anti-mouse TNF mAb. For cell surface receptor staining, J774.1 cells were stained with PE-conjugated anti-mouse CD120a (TNFR type I) mAb (clone 55R-286, BioLegend, San Diego, CA, USA), PE-conjugated anti-mouse CD120b (TNFR type II) mAb (clone TR75-89, AbD Serotec, Oxford, UK), PE-conjugated anti-mouse CD14 mAb (clone rmC5-3, BD Biosciences), or FITC-conjugated anti-TLR4/myeloid differentiation protein 2 (MD2) complex mAb (clone MTS510, Stressgen Bioreagents, Ann Arbor, MI, USA). Expression of each molecule was measured using a FACSCalibur flow cytometer and CellQuest software (BD Biosciences).

Measurement of lactate dehydrogenase (LDH) activity
For the quantification of plasma membrane damage, LDH activity in the culture supernatants of J774.1 cells was measured with a cytotoxicity detection kit (Roche Diagnostics, Indianapolis, IN, USA), according to the manufacturer’s instructions.

Quantitative RT-PCR (qRT-PCR)
Cells were lysed in 1 ml Isogen (Nippon Gene, Toyama, Japan), and total RNA was extracted as described in the instruction manual. Total RNA was dissolved in 30 µl diethyl pyrocarbonate-treated water (Nippon Gene) and incubated at 65°C for 10 min. cDNA synthesis was carried out with a first-strand cDNA synthesis kit (GE Healthcare Bio-Science Corp., Piscataway, NJ, USA). Real-time PCR was performed with a LightCycler FastStart DNA Master SYBR Green I and a LightCycler 1.5 system (Roche Diagnostics). The primers used for PCR were as follows: TNF-{alpha}, forward 5'-AGCCTCTTCTCATTCCTGC-3' and reverse 5'-GGAGGCCATTTGGGAACT-3'; β-actin, forward 5'-CGTTGACATCCGTAAAGACCTC-3' and reverse 5'-AGCCACCGATCCACACAGA-3' (Nihon Gene Research Labs Inc., Sendai, Japan). The PCR conditions were 35 cycles at 95°C for 10 s, 60°C for 10 s, and 72°C for 10 s. The product sizes for TNF-{alpha} and β-actin were 106 bp and 173 bp, respectively. The mRNA expression levels were expressed as relative units after normalization by the β-actin level. The specificity of the PCR was confirmed by the molecular weight of the products and melting curve analysis for each data point. PCR products were electrophoresed using 3% agarose (Nusieve 3:1 agarose, BMA, Rockland, ME, USA). After staining with ethidium bromide, amplified DNA bands were analyzed with a Chemi Imager.

Measurement of NF-{kappa}B and AP-1 activity
Activities of NF-{kappa}B family and AP-1 in nuclear extracts were measured with a NF-{kappa}B assay kit specific for the p65, p50, and RelB, p52 subunits, and an AP-1 assay kit specific for the phospho c-Jun, according to the manufacturer’s instructions (Active Motif, Carlsbad, CA), respectively. The nuclear extractions were performed with a nuclear extract kit (Active Motif), according to the manufacturer’s instructions. Protein concentrations of each fraction were determined with a bicinchoninic acid protein assay kit (Pierce Biotechnology, Inc.). Each activity was expressed as absorbance at 450 nm (A450 nm) per 100 µg protein.

Data analysis
All of the experiments in this study were performed at least three times to confirm the reproducibility of the results. The data shown are representative results. Experimental values are given as the mean ± SD of triplicate assays. Statistical analysis was performed with the unpaired t-test or one-way ANOVA using Dunnett’s method, and P < 0.05 was considered significant.


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RESULTS
 
Augmentation of LPS-induced serum TNF-{alpha} levels in biotin-deficient mice
As it is well known that LPS induces the elevation of the serum concentration of TNF-{alpha} in mice, we first examined the in vivo effects of biotin deficiency on LPS-induced TNF-{alpha} production. Mice were fed biotin-sufficient or -deficient diets. After 8 weeks of feeding, the serum concentrations of biotin were significantly (P<0.001) lower in biotin-deficient mice than biotin-sufficient mice (Fig. 1A ). However, PCC activities in the liver were comparable between biotin-sufficient and -deficient mice (sufficiency: 2.99±0.49 nmol HCO3– fixed/min/mg protein; deficiency: 2.80±0.34 nmol HCO3– fixed/min/mg protein; n=5; P=0.517). No clinical symptoms were detected in the biotin-deficient mice, and no significant differences of body weights were detected between biotin-sufficient and -deficient mice (data not shown). A significant (P<0.01) increase of the serum TNF-{alpha} level was induced 90 min after i.v. injection of LPS (1 µg/kg) in both groups (Fig. 1B) . In biotin-deficient mice, the concentration of TNF-{alpha} was significantly (P<0.05) higher than that in biotin-sufficient mice. These results indicated that biotin deficiency augments TNF-{alpha} production in vivo.


Figure 1
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  		Figure 1. Serum levels of biotin and TNF-{alpha} in biotin-sufficient and -deficient mice. Female BALB/c mice (4 weeks old) were fed a biotin-sufficient or -deficient diet for 8 weeks. (A) Concentrations of biotin in sera were measured by ELISA after prior separation by HPLC. The results were expressed as mean ± SD of triplicate assay. ***, P < 0.001, compared with biotin sufficiency. (B) The mice were challenged i.v. with LPS (1 µg/kg) or saline alone, and blood was taken at 90 min after injection. Concentrations of TNF-{alpha} in sera were measured by ELISA. The results were expressed as mean ± SD for four mice. ND, Not detected; **, P < 0.01, compared with saline; #, P < 0.05, compared with biotin sufficiency.

In vitro effects of biotin deficiency on murine macrophages
Macrophages are the major cells that produce TNF-{alpha} in response to LPS. Therefore, we next examined the in vitro effects of biotin deficiency using J774.1 cells. In biotin-sufficient J774.1 cells, 130- and 80-kDa polypeptides were detected by Western blotting with HRP-conjugated avidin. On the other hand, these polypeptides were not detected after 2 weeks cultivation in biotin-deficient medium (Fig. 2A ). On the basis of molecular weight, it is likely that the 130-kDa polypeptide is pyruvate carboxylase, and the 80-kDa polypeptide is propionyl-CoA carboxylase and/or methylcrotonyl-CoA carboxylase. Moreover, [3H] TdR incorporation was significantly (P<0.01) lower in biotin-deficient cells than biotin-sufficient cells (Fig. 2B) . These results clearly indicated that biotinylation of cellular proteins and cell proliferation was reduced by biotin deficiency.


Figure 2
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  		Figure 2. In vitro effects of biotin deficiency on J774.1 cells, which were (A) cultured with biotin-sufficient or -deficient medium for the time indicated. Cells were lysed in SDS-PAGE sample buffer. Cell lysate (1x104 cells each) was subjected to Western blotting with HRP-conjugated avidin. (B) J774.1 cells were cultured with biotin-sufficient or -deficient medium for 4 weeks. Biotin-sufficient and -deficient cells in the medium with and without biotin, respectively, were seeded in 96-well flat-bottomed plates and incubated at 37°C for 3 h. Cells were pulsed with 1 kBq/well [3H] TdR for 2 h. After the pulse, 100 µl 3% Triton X-100 was added to each well, and cells were harvested onto a glass fiber filter. **, P < 0.01, compared with biotin sufficiency.

Augmentation of LPS-induced TNF-{alpha} production in biotin-deficient J774.1 cells
Next, we analyzed the production of TNF-{alpha} from biotin-sufficient and -deficient J774.1 cells. As shown in Figure 3A , both types of cells produced TNF-{alpha} in response to LPS in a dose-dependent manner. The concentration of TNF-{alpha} in the culture supernatant of biotin-deficient cells was significantly (P<0.01) higher than that of biotin-sufficient cells, even without LPS stimulation. A similar pattern of TNF-{alpha} production was observed when the type of medium was replaced with the opposite type during LPS stimulation for 24 h, which excluded the possibility of contamination with stimulatory or inhibitory factors in either type of medium. LDH activity was not detected in the culture supernatant of biotin-deficient cells (data not shown), indicating that the augmentation of TNF-{alpha} production in biotin-deficient cells was not a result of plasma membrane damage. The levels of TNF-{alpha} mRNA were significantly (P<0.01) elevated in biotin-deficient cells compared with those in biotin-sufficient cells with or without LPS stimulation (Fig. 3B) . Moreover, flow cytometric analysis revealed that the expression levels of cell surface and intracellular TNF-{alpha} were also higher in biotin-deficient cells than in biotin-sufficient cells (Fig. 3C and 3D) .


Figure 3
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  		Figure 3. TNF-{alpha} production of biotin-sufficient and -deficient J774.1 cells, which were cultured with biotin-sufficient or -deficient medium for 4 weeks. (A) Cells were seeded in 96-well flat-bottomed plates at 2 x 105 cells/200 µl/well with biotin-sufficient and -deficient medium and then stimulated with LPS at 37°C for 24 h. Concentrations of TNF-{alpha} in culture supernatants were measured by ELISA. **, P < 0.01, compared with medium alone (0 ng/ml LPS); ##, P < 0.01, compared with biotin sufficiency. (B) Biotin-sufficient and -deficient cells in medium with and without biotin, respectively, were seeded in 24-well flat-bottomed plates at 5 x 105 cells/500 µl/well and then stimulated with LPS (10 ng/ml) at 37°C for 4 h. TNF-{alpha} mRNA expression levels were determined by qRT-PCR. The results were expressed as relative units after normalization by the β-actin level. **, P < 0.01; ***, P < 0.001, compared with medium alone; ##, P < 0.01, compared with biotin sufficiency. (C and D) Cell surface (C) and intracellular TNF-{alpha} (D) of biotin-sufficient and -deficient cells were analyzed by flow cytometry. Dotted line, Unstained J774.1 cells (control); thin line, biotin sufficiency; bold line, biotin deficiency. Numbers in histograms indicate mean fluorescence intensity.

We also analyzed the effects of Act-D, an inhibitor of RNA polymerase II. In the presence of Act-D, intracellular TNF-{alpha} was decreased slightly in biotin-sufficient cells (Fig. 4A ). On the other hand, it was markedly decreased in biotin-deficient cells (Fig. 4B) and in LPS-stimulated, biotin-sufficient cells (Fig. 4C) . These results clearly indicated that biotin deficiency induces the augmentation of TNF-{alpha} production in vitro at the transcriptional level.


Figure 4
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  		Figure 4. Effects of Act-D on intracellular TNF-{alpha} levels of biotin-sufficient and -deficient J774.1 cells, which were cultured with biotin-sufficient or -deficient medium for 4 weeks. Biotin-sufficient and -deficient cells in the medium with and without biotin, respectively, were seeded in six-well flat-bottomed plates at 1 x 106 cells/2 ml/well with (thin line) or without (bold line) Act-D (1 µg/ml) at 37°C for 6 h. Intracellular TNF-{alpha} was analyzed by flow cytometry. The dotted line shows unstained J774.1 cells (control). (A) Biotin sufficiency without LPS; (B) biotin deficiency without LPS; (C) biotin sufficiency with LPS (10 ng/ml). Numbers in histograms indicate mean fluorescence intensity.

Cell surface expression of TNFRs
It was reported that biotin supplementation induces IL-2R{gamma} expression and decreases the net secretion of IL-2 by endocytosis of the cytokine [9 ]. Therefore, we measured cell surface expressions of TNFR types I and II on biotin-sufficient and -deficient J774.1 cells using a flow cytometer. As shown in Figure 5A and 5B , the expression levels of TNF-R types I and II were similar between biotin-sufficient and -deficient cells. Mean fluorescence intensities of TNFR type I in biotin-sufficient and -deficient cells were 33.3 and 30.6, respectively; those of TNFR type II in biotin-sufficient and -deficient cells were 19.4 and 17.7, respectively. These results indicated that the augmentation of TNF-{alpha} production in biotin-deficient cells was not a result of down-regulation of the cell surface expression of TNFR and endocytosis of TNF-{alpha}.


Figure 5
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  		Figure 5. Expression of TNFR, CD14, and TLR4/MD2 complex on biotin-sufficient and -deficient J774.1 cells, which were cultured with biotin-sufficient or -deficient medium for 4 weeks. Cells were stained with PE-conjugated anti-mouse TNFR type I (A), PE-conjugated anti-mouse TNFR type II (B), PE-conjugated anti-mouse CD14 (C), or FITC-conjugated anti-mouse TLR4/MD2 complex (D) and analyzed by flow cytometry. Dotted line, unstained J774.1 cells (control); thin line, biotin sufficiency; bold line, biotin deficiency.

Cell surface expression of CD14 and TLR4
To examine the possibility that biotin deficiency up-regulates LPS responsiveness through augmentation of LPS receptors, we measured the cell surface expression of CD14 and the TLR4/MD2 complex on biotin-sufficient and -deficient J774.1 cells. No differences were detected in the expression of these LPS receptors between biotin-sufficient and -deficient cells. Mean fluorescence intensities of CD14 in biotin-sufficient and -deficient cells were 9.0 and 9.2, respectively (Fig. 5C) . Those of the TLR4/MD2 complex in biotin-sufficient and -deficient cells were 2.5 and 2.2, respectively (Fig. 5D) . These results indicated that up-regulation of TNF-{alpha} production in biotin-deficient cells was not a result of augmentation of LPS receptors.

Activities of transcriptional factors in biotin-sufficient and -deficient J774.1 cells
We also analyzed the activities of two major transcriptional factors, which are responsible for TNF-{alpha} mRNA transcription—the NF-{kappa}B family (p65, p50, RelB, and p52) and AP-1 (phospho c-Jun)—in nuclear fractions of biotin-sufficient and -deficient J774.1 cells. The activities of NF-{kappa}B p65, p50, and AP-1 were increased significantly with LPS stimulation. However, no significant differences were detected in NF-{kappa}B family and AP-1 activities between biotin-sufficient and -deficient cells (Fig. 6 ), indicating that other factors are involved in the augmentation of TNF-{alpha} mRNA transcription in biotin-deficient cells.


Figure 6
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  		Figure 6. NF-{kappa}B and AP-1 activities in biotin-sufficient and -deficient J774.1 cells, which were cultured with biotin-sufficient or -deficient medium for 4 weeks. Biotin-sufficient and -deficient cells in the medium with and without biotin, respectively, were cultured in 1.5-ml tubes at 1 x 106 cells/0.5 ml/tube with LPS (10 ng/ml) at 37°C for the indicated time. NF-{kappa}B and AP-1 (phospho c-Jun) activities in the nuclear fraction were determined with NF-{kappa}B and AP-1 assay kits, respectively. Each activity was expressed in relative units defined as A450 nm per 100 µg protein. (A) NF-{kappa}B p65; (B) NF-{kappa}B p50; (C) RelB; (D) NF-{kappa}B p52; (E) AP-1. *, P < 0.05; **, P < 0.01, compared with no LPS stimulation (0 min).

Regulation of TNF-{alpha} levels by biotin supplementation in biotin-deficient J774.1 cells and biotin-deficient mice
To further confirm the effects of biotin deficiency, J774.1 cells were cultured with biotin-deficient medium for 4 weeks and then further incubated with biotin-sufficient medium for 2 weeks (biotin-supplement cells). [3H] TdR incorporation in biotin-supplemented cells was still lower than that in biotin-sufficient cells but was significantly (P<0.01) higher than that in biotin-deficient cells (Fig. 7A ). The concentrations of TNF-{alpha} in the culture supernatants of biotin-supplemented cells with and without LPS stimulation were significantly (P<0.01) reduced to near the levels in the supernatants of biotin-sufficient cells (Fig. 7B) . The TNF-{alpha} production of biotin-deficient cells was significantly (P<0.01) decreased, even in the presence of 50 µM biotin during LPS stimulation (Fig. 7C) . When biotin-deficient mice were provided with biotin-supplemented drinking water, the LPS-induced serum TNF-{alpha} levels of the mice were significantly (P<0.05) decreased (Fig. 7D) . These results indicated that biotin supplementation restored the TNF-{alpha} production to the basal level in vitro and in vivo.


Figure 7
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  		Figure 7. Effects of biotin supplementation in vitro and in vivo. (A) J774.1 cells were cultured with biotin-deficient medium for 4 weeks and then incubated further in medium without biotin (biotin deficiency) or with biotin (biotin supplementation) for 2 weeks. J774.1 cells were also cultured in biotin-sufficient medium for 6 weeks (biotin sufficiency). [3H] TdR incorporation is shown in Figure 2B . **, P < 0.01, compared with biotin sufficiency; ##, P < 0.01, compared with biotin deficiency. (B) Cells (2x105 cells/200 µl/well) in A were stimulated with LPS at 37°C for 24 h. Concentrations of TNF-{alpha} in culture supernatants were measured by ELISA. *, P < 0.05; **, P < 0.01; ***, P < 0.001, compared with biotin sufficiency; ###, P < 0.001, compared with biotin deficiency. (C) Cells (2x105 cells/200 µl/well) in A were stimulated with 10 ng/ml LPS in the presence of 50 µM biotin at 37°C for 24 h. Concentrations of TNF-{alpha} in culture supernatants were measured by ELISA. **, P < 0.01, compared with biotin deficiency (without biotin). (D) Mice were fed a biotin-deficient diet for 10 weeks (deficiency). Biotin-supplemented drinking water (15 µM) was administered to the mice for the last 2 weeks (biotin supplementation). The mice were then challenged i.v. with LPS (1 µg/kg) or saline alone, and blood was collected at 90 min after injection. Concentrations of TNF-{alpha} in sera were measured by ELISA. The results were expressed as mean ± SD for four mice. **, P < 0.01, compared with biotin sufficiency; #, P < 0.05, compared with biotin deficiency.


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DISCUSSION
 
In this study, we clearly demonstrated that TNF-{alpha} production is up-regulated under biotin-deficient conditions and that biotin supplementation down-regulates the TNF-{alpha} production to the basal level in vivo and in vitro. These results suggest that biotin contributes to the regulation of inflammatory responses.

As biotin is produced by intestinal bacteria, many studies analyzed biotin-deficient animals that were fed a biotin-deficient diet containing egg-white protein, which contains avidin, a glycoprotein that forms a noncovalent and nonabsorbed complex with biotin [11 12 13 , 23 ]. It was reported that biotin-deficient mice fed egg white for more than 7 weeks showed body weight loss and clinical symptoms, such as alopecia and squatting posture [11 ]. Biotin-deficient rats fed egg white for 40 days showed partial alopecia and neurologic signs, such as kangaroo gait and irritability [13 ]. Moreover, severe biotin deficiency causes alopecia and scaly erythematous dermatitis in humans [11 , 14 , 15 ]. In this study, biotin-deficient mice were fed a biotin-deficient AIN-76 basal diet for 8 weeks, and the biotin concentrations in sera from biotin-deficient mice were significantly (P<0.001) lower than those from biotin-sufficient mice (Fig. 1A) . However, PCC activities in the liver were comparable in biotin-sufficient and -deficient mice, and no body weight loss or clinical symptoms were detected. Therefore, we considered that our method causes mild biotin deficiency in mice.

Biotin-deficient J774.1 cells exhibited lower proliferation than biotin-sufficient cells (Fig. 2B) . It was reported that intracellular uptake of biotin increases with cell proliferation [24 ]. As mentioned in Introduction, biotin-dependent carboxylases are essential for cellular metabolism, indicating that biotin deficiency causes inactivation of biotin-dependent carboxylases followed by the down-regulation of cellular metabolism and proliferation.

TNF-{alpha} production was augmented in biotin-deficient mice (Fig. 1B) and cells (Fig. 3) . Biotin-deficient J774.1 cells produced a significantly higher amount of TNF-{alpha}, even without LPS stimulation. Moreover, the cell surface expression levels of the CD14 and TLR4/MD2 complex were not significantly different between biotin-sufficient and -deficient cells (Fig. 5C and 5D) . These results exclude the possibility that the augmentation of TNF-{alpha} production was mediated by up-regulation of LPS receptors. The TNF-{alpha} mRNA expression was significantly higher in biotin-deficient cells than -sufficient cells, and intracellular TNF-{alpha} was decreased markedly in the presence of Act-D (Fig. 4) . It was reported that biotin supplementation up-regulates the expression level of the IL-2R and decreases the net production of IL-2 by endocytosis of cytokines [9 ]. However, no differences were detected in the expression levels of TNFR types I and II between biotin-sufficient and -deficient cells (Fig. 5A and 5B) . Furthermore, no differences were observed in the TNF-{alpha}-converting enzyme activities or mRNA levels of the tissue inhibitor of metalloproteinase-3, an inhibitor of the TNF-{alpha}-converting enzyme, between biotin-sufficient and -deficient cells (data not shown). The expression of intracellular and cell surface TNF-{alpha} was higher in biotin-deficient than -sufficient cells (Fig. 3C and 3D) . These results indicate that the augmentation of TNF-{alpha} production was regulated at the transcriptional level.

On the other hand, the biotin content in the AIN-76 diet (0.8 mg/kg) is much higher than the metabolic requirement of biotin. It was reported that the adequate intake of biotin for adult humans (~60 kg body weight) is 45 µg/day (0.75 µg/kg/day, Ministry of Health, Labor and Welfare, Japan). Based on the biotin content (0.8 mg/kg) and the total intake of diet (~5 g/day), biotin-sufficient mice (~25 g body weight) received 4 µg biotin per day (160 µg/kg/day). Moreover, the concentration of biotin in RPMI-1640 medium (0.2 µg/ml, ~800 nM) is much higher than biotin levels in normal mouse (40 nM) and human sera (0.2 nM) [11 , 25 ]. Therefore, it is possible that the excess of biotin is involved in the down-regulation of TNF-{alpha} production.

No differences were detected in the activities of two major transcriptional factors involved in regulating TNF-{alpha} expression, namely NF-{kappa}B family members and AP-1 (Fig. 6) . This result conflicts with the finding reported by Rodriguez-Melendez et al. [5 ] that the nuclear translocation and transcriptional activities of NF-{kappa}B were increased by biotin deficiency in Jurkat cells. The discrepancy may have been caused by the differences of cell line (murine macrophage cell line J774.1 and human T cell line Jurkat) or assay conditions. J774.1 cells were stimulated with LPS up to 60 min. On the other hand, Jurkat cells were stimulated with PMA and phytohemagglutinin for 3 h. Further studies are needed to clarify this point. It was reported that lysine residues in histones are modified by biotinylation, and the biotinylation of histones was enriched in transcriptionally inactive chromatin [6 , 26 , 27 ]. It is well known that some histone modifications, such as acetylation and methylation, correlate with transcriptional activation [28 ]. These observations suggest that the mechanism by which biotin deficiency contributes to up-regulation of TNF-{alpha} in J774.1 cells might be through biotin deficiency, producing reduced biotinylation of critical histones, leading to increased gene transcription.

Because of the uncertain immunological and pharmacological mechanisms, few studies have been reported about biotin treatment of inflammatory diseases [16 ]. On the other hand, it was reported that 10 nM biotin inhibited IL-2 production by Jurkat cells [8 , 9 ]. Moreover, Zempleni et al. [10 ] reported that in vivo supplementation of biotin in healthy human subjects (3.1 µmol/day) inhibited IL-1β and IL-2 production of PBMCs. In this study, we demonstrated clearly that biotin supplementation down-regulated the augmented TNF-{alpha} production induced by biotin deficiency in vivo and in vitro (Fig. 7) . Therefore, we speculate that a pharmacological dose of biotin might have some potentially therapeutic effects on inflammatory diseases, but this remains to be tested empirically.

TNF-{alpha} production of biotin-deficient J774.1 cells was drastically decreased by biotin supplementation in experiments in which biotin-deficient cells were cultured with biotin-sufficient medium (containing ~0.5 µM biotin) for 2 weeks (Fig. 7B) . On the other hand, TNF-{alpha} production was similar in biotin-deficient cells cultured with biotin-sufficient and -deficient medium (Fig. 3A) . A high dose of biotin (50 µM) slightly but significantly decreased TNF-{alpha} production by biotin-deficient cells (Fig. 7C) . These results indicate that long-term supplementation of biotin is more effective for the inhibition of TNF-{alpha} up-regulation than stimulation with a high dose of biotin. Therefore, we considered that TNF-{alpha} up-regulation in biotin-deficient cells is regulated via various metabolic pathways that are affected by biotin rather than via the direct effects of biotin about TNF-{alpha} production.

It is well known that biotin deficiency causes cutaneous abnormalities [11 , 14 , 15 ]. In addition, it was reported that the biotin concentration in serum correlates with inflammatory diseases [11 , 14 , 15 , 18 ]. Although several studies reported the contribution of abnormalities in lipid metabolism to cutaneous abnormalities (reviewed in ref. [14 ]), the pathological mechanisms of disease conditions caused by biotin deficiency remain to be clarified. In this study, we clearly demonstrated that biotin regulates TNF-{alpha} production in vivo and in vitro. TNF-{alpha} plays important roles in the pathogenesis of inflammatory diseases, which have been reported to be correlated with biotin [29 30 31 ]. Therefore, biotin status may be involved in the pathological mechanisms of dermatitis and other inflammatory diseases. Our results should encourage further investigations on biotin treatment for various inflammatory diseases.


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ACKNOWLEDGEMENTS
 
This study was supported in part by Grants-in-Aid for Scientific Research from the Japan Society for the Promotion of Science (18791379) and a 21st Century Center for Excellence Program Special Research Grant from the Ministry of Education, Culture, Sports, Science, and Technology, Japan.

Received June 25, 2007; revised December 5, 2007; accepted December 5, 2007.


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