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3
*
Departments of Nutritional Science and Dietetics and
Biochemistry, University of Nebraska at Lincoln, Lincoln, NE 68583-0806
3To whom correspondence should be addressed. E-mail: jzempleni2{at}unl.edu.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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KEY WORDS: • biotin • carboxylases • histones • humans • interleukin-2
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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). These carboxylases catalyze essential steps in the metabolism of glucose, amino acids and fatty acids. The attachment of biotin to the 
-amino group of a lysine residue in the four apocarboxylases is catalyzed by holocarboxylase synthetase (EC 6.3.4.10). Degradation of holocarboxylases leads to the formation of biotinylated peptides. These peptides are further degraded by biotinidase (EC 3.5.1.12) to release free biotin, which can then be used for the synthesis of new holocarboxylases (2
). Recently, Hymes et al. (3
) proposed an enzymatic mechanism by which biotinidase may also catalyze binding of biotin to histones, suggesting a potential role of biotin in transcription, replication or repair of DNA. Based on these pioneering studies, our laboratory has demonstrated that human cells indeed biotinylate histones (4
).
Consistent with these essential roles of biotin in metabolism, biotin status may affect cellular growth, proliferation and differentiation. For example, biotin deficiency causes arrest of HeLa cells in the G1 phase of the cell cycle (5
). Potentially, abnormal cellular growth and differentiation is the underlying cause of fetal malformations (6
,7
) and impaired immune function (8
) observed in biotin-deficient animals.
Proliferating cells may have an increased demand for biotin compared with quiescent cells. Previous studies have provided evidence that human mononuclear cells respond to proliferation with increased biotin uptake (9
), mediated by increased expression of biotin transporters (10
). Proliferating cells utilize biotin to increase biotinylation of carboxylases (11
) and histones (4
) compared with quiescent cells.
Notwithstanding the essential role of biotin in cell proliferation, ingestion of pharmacologic doses of biotin by humans might reduce both proliferation rates of mononuclear cells and secretion of some interleukins by these cells (12
). In summary, ingestion of both deficient and pharmacologic amounts of biotin might adversely affect cell development and function. In this study, a human T cell line (Jurkat cells) was used to determine whether deficient and pharmacologic concentrations of biotin in culture medium affect cellular biotin status, as judged by carboxylase activities, biotinylation of carboxylases, and biotinylation of histones; expression of genes encoding biotinidase and PCC; expression of biotin transporter genes; and rates of cell proliferation and net secretion of interleukin-2 into the culture medium.
Jurkat cells were chosen for these studies based on the following lines of reasoning: Jurkat cells produce interleukin-2 in response to appropriate stimulation. The studies presented here required use of a cell line that can be cultured for several weeks to achieve cellular homeostasis with regard to biotin metabolism. In contrast to primary cell lines (such as freshly isolated lymphocytes), Jurkat cells can be cultured for extended periods.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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Jurkat cells (clone E6–1) were purchased from American Type Culture Collection (Manassas, VA). Medium for cell culturing contained the following components: 0.9 L of RPMI-1640 (Atlanta Biologicals, Norcross, GA) without biotin per liter of final medium; 0.1 L of fetal bovine serum without biotin (see below) per liter of final medium; and 100,000 IU/L penicillin and 100 mg/L streptomycin (final concentrations).
Before medium preparation, fetal bovine serum was depleted of biotin in our laboratory using a column packed with immobilized avidin (Immunopure; Pierce, Rockford, IL); 1.5 mL of immobilized avidin was used to deplete 70 mL of serum. After depletion, the biotin concentration in serum was determined by avidin-binding assay as described previously (13
) with the following modifications. For color development, 100 µL of TMB 1-component microwell substrate (Kirkegaard & Perry Laboratories, Gaithersburg, MD; catalogue number 50-76-04) were added to each sample on the 96-well plate; samples were incubated for 0.5 h at room temperature. Reactions were terminated by addition of 100 µL of stop solution (Kirkegaard & Perry Laboratories; catalogue number 50-85-04) and the absorbance was read at 450 nm in an Emax microwell plate reader (Molecular Devices, Sunnyvale, CA). Biotin in depleted serum was below detection limit (< 30 pmol/L) by assay, suggesting that biotin-depleted serum did not contain a meaningful amount of biotin.
Biotin-depleted serum was mixed with biotin-free RPMI 1640 and antibiotics as indicated above. Next, biotin was added to the medium to produce the following final concentrations: 25 pmol/L of biotin (denoted deficient), 250 pmol/L of biotin (denoted physiologic), or 10,000 pmol/L of biotin (denoted pharmacologic). These concentrations were chosen based on the following lines of reasoning: 250 pmol/L of biotin equals the physiologic concentration of biotin in plasma from healthy adults (14
). Twenty-five pmol/L of biotin is > 2 SD below the mean physiologic concentration in normal plasma (14
); thus, 25 pmol/L equals a deficient concentration of biotin. Ingestion of a typical biotin supplement providing 25 times the adequate intake of biotin for adults (15
) is associated with plasma concentrations of 
10,000 pmol/L of biotin in healthy adults (12
); thus, this concentration represents a pharmacologic concentration of biotin in plasma.
In a prephase to this study, Jurkat cells from all treatment groups were cultured in medium containing 250 pmol/L biotin for 20 d to adjust the cells to physiologic biotin concentrations; medium was replaced with fresh medium 12 times during this period of time. After completion of this prephase, cells were split and transferred into media containing 25, 250 or 10,000 pmol/L of biotin (denoted d 0). Culturing was continued for 4 wk; culture medium was replaced with fresh medium every 48 h. At timed intervals, aliquots were collected and analyzed as described below.
Propionyl-CoA carboxylase activity.
This assay quantifies the binding rate of radioactive bicarbonate to propionyl-CoA, catalyzed by PCC in samples of lysed cells. Aliquots of cell suspension (8 x 106 cells) were centrifuged for 10 min at 2260 x g; supernatants were discarded and the cell pellets were suspended in 180 µL of homogenization buffer as described previously (16
). After vortexing, 50 µL of Triton X-100 (5 mL of Triton X-100/L water) were added to lyse the cells. Complete lysis of cells by this procedure was confirmed by visual inspection using a light microscope. PCC activity was quantified as described previously with minor modifications (16
). Briefly, lysed Jurkat cells were incubated with propionyl-CoA, [14C]bicarbonate, and cofactors to allow for covalent binding of [14C]bicarbonate to propionyl-CoA. After incubation, unbound [14C]bicarbonate was volatilized by addition of perchloric acid and samples were dried. Finally, samples were suspended in scintillation fluid and the bound [14C]bicarbonate was quantified by liquid scintillation counting.
Biotinylation of biotin-dependent carboxylases.
Holocarboxylases (as opposed to apocarboxylases) contain covalently bound biotin. Biotin in holocarboxylases can be probed with streptavidin peroxidase, using Western blots of cellular protein extracts. In this study, 6 x 106 Jurkat cells were lysed in 480 µL of detergent (BugBuster; Novagen, Madison, WI), immediately followed by the addition of 1.6 µL of protease inhibitor cocktail (Sigma, St. Louis, MO; catalogue number P-8340) and 0.5 µL of DNase (Benzonase; Novagen). Samples were incubated for 5 min at room temperature to allow for hydrolysis of DNA by DNase. Next, 65 µL of sample were mixed with 25 µL of loading buffer (Nupage 4x; Invitrogen, Carlsbad, CA), and 10 µL of 1 mol/L dithiothreitol; samples were heated for 10 min at 70°C and loaded onto a 3–8% Tris-Acetate gel (Invitrogen). After electrophoresis, proteins were transferred onto polyvinylidene fluoride membranes by electroblotting. Biotin in carboxylases was probed using streptavidin peroxidase in analogy to our previous studies of biotinylated histones (4
).
Human cells contain five classes of histones: H1, H2A, H2B, H3 and H4. Histones were extracted from Jurkat cell nuclei (4 x 106 cells) by using hydrochloric acid as described previously (4
). Equimolar amounts of histones (as judged by gel densitometry after staining with coomassie blue) were electrophoresed using 16% Tris glycine gels (Invitrogen); biotin in histones was probed using streptavidin peroxidase as described previously (4
).
Rates of biotin transport into Jurkat cells were determined using 475 pmol/L [3H]biotin as described previously (17
); the concentration of biotin that was used in transport studies is within the range of biotin concentrations in normal human plasma (14
). Biotin uptake into mammalian cells is mediated by the sodium-dependent multivitamin transporter (SMVT) (18
) and, perhaps, other yet unidentified biotin transporters (17
,19
). The mass of SMVT protein in cells was quantified by Western blotting, using a polyclonal antibody to human SMVT as described previously (20
).
Quantification of mRNA levels.
Levels of mRNA encoding PCC and biotinidase were quantified by polymerase chain reaction (PCR); mRNA encoding the housekeeping gene glyceraldehyde-3-phosphate dehydrogenase was quantified as a control. Total RNA was extracted from Jurkat cells by the RNeasy mini kit according to the manufacturer’s instructions (Qiagen, Valencia, CA). The amount of RNA was quantified spectrophotometrically at 260 nm. One microgram of RNA from each sample was reverse transcribed using the Smart PCR cDNA synthesis kit (Clontech, Palo Alto, CA) and Reverse Transcriptase Superscript II RT (Life Technologies, Gaithersburg, MD). The ssDNA was amplified by PCR using the following oligonucleotide primers (Integrated DNA Technologies, Coralville, IA): 5'-ATG GCG CAT GCG CAT ATT CAG GGC G-3' and 5'-GCC AAG CTG GTC AGG GAC TTC TAG G-3' for human biotinidase (21
); 5'-TCC GTG AGT GCC ACG ATC CCA GTG ACC-3' and 5'-GAG CTC ATG ACA TCA TAG GCA CCT CCA-3' for human PCC (22
); and 5'-ACC ACA GTC CAT GCC ATC ACT GCC ACC-3' and 5'-TCC ACC ACC CTG TTG CTG TAG CCA AAT-3' for human glyceraldehyde-3-phosphate dehydrogenase (23
).
PCR was performed using the following temperatures and times per cycle: 94°C for 1 min (denaturating), 55°C for 1 min (annealing), and 72°C for 2 min (extending). The mass of DNA produced by PCR parallels the mass of mRNA (template) that was present in cells by time of RNA purification. The PCR-amplified DNA was collected at timed intervals for up to 60 PCR cycles. Only values from within the linear phase of PCR amplification (typically < 30 PCR cycles) were considered for data analysis. Equal volumes (10 µL) of each sample were chromatographed using agarose gels (15 g agarose/L). DNA on gels was stained with ethidium bromide and analyzed using the Kodak EDAS 290 Documentation and Analysis System (Rochester, NY).
Cellular uptake of purine or pyrimidine bases (e.g., thymidine), or analogs thereof (e.g., 5-bromo-2'-deoxyuridine), increases in response to cell proliferation due to synthesis of new DNA. In this study, proliferation rates of Jurkat cells were quantified by measuring the cellular uptake of 5-bromo-2'-deoxyuridine (10 µmol/L final concentration), using a commercially available enzyme-linked immunosorbent assay (Cell proliferation ELISA system 2, Amersham Pharmacia Biotech, Piscataway, NJ). Jurkat cells were cultured with 5-bromo-2'-deoxyuridine for 1 h at a cell density of 50,000 cells/well in a 96-well plate. In pilot studies we determined that uptake of 5-bromo-2'-deoxyuridine increased linearly with incubation time for up to 1.5 h under the experimental conditions (data not shown).
In addition, proliferation rates were determined by measuring the uptake of [3H]thymidine (specific radioactivity 1.3 TBq/mmol; ICN, Irvine, CA) into Jurkat cells; [3H]thymidine uptake was measured as described previously (9
).
Theoretically, net rates of cell proliferation could be affected by changes in rates of cell death, apoptosis. Thus, cells were monitored for an early marker for apoptosis: activity of caspase-3. Enzyme activities were measured using a commercially available kit (ApoAlert Caspase-3 Colorimetric Assay Kit; Clontech).
Concentrations of interleukin-2 in culture media.
In pilot studies we quantified the secretion of interleukin-2 by Jurkat cells in response to various inducers of interleukin-2 secretion: phorbol-12-myristate-13-acetate, phytohemagglutinin, ionomycin, concanavalin A, and mouse anti-human CD28 cell surface marker; inducers were used in various combinations. Also, we established time-response curves of interleukin-2 secretion. Rates of interleukin-2 secretion were maximal within 24 h of addition of phorbol-12-myristate-13-acetate and phytohemagglutinin to the culture medium (data not shown). Thus, for this study we chose the following conditions to induce secretion of interleukin-2 by Jurkat cells: 1 x 106 cells were incubated with 50 µg/L of phorbol-12-myristate-13-acetate and 2 mg/L of phytohemagglutinin in a volume of 260 µL for 24 h (24
). Then, cell suspensions were centrifuged (400 x g for 10 min) and the supernatants were collected. Concentrations of interleukin-2 in medium were quantified by using a commercially available enzyme-linked immunosorbent assay (hIL-2 EAISA; Biosource, Camarillo, CA). Please note that some immune cells are capable of both secreting and internalizing interleukin-2 (25
); thus, concentrations of interleukin-2 in culture medium are the net result of both processes.
Homogeneity of variances among groups was tested using Bartlett’s test (26
). Variances were homogenous; therefore, data were not transformed before further statistical testing. Significance of differences among groups was tested by one-way ANOVA. Fisher’s Protected Least Significant Difference procedure was used for post hoc testing (26
). StatView 5.0.1 (SAS Institute, Cary, NC) was used to perform all calculations. Differences were considered significant at P < 0.05. Data are expressed as mean ± 1 SD.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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First, we determined whether biotin concentrations in culture medium affected intracellular biotin status, as judged by activities of PCC. When cells were cultured in biotin-deficient medium for 4 wk, activities of PCC were 19 ± 0.7 pmol bicarbonate/(106 cells x min) compared with 1.9 ± 1.2 pmol bicarbonate/(106 cells x min) in cells that were cultured in medium containing physiologic concentrations of biotin (P < 0.01; n = 6); activity of PCC was 40 ± 2.0 pmol bicarbonate/(106 cells x min) if cells were cultured in medium containing pharmacologic concentrations of biotin (P < 0.01 vs. other treatment groups).
Next, we determined whether PCC activities paralleled biotinylation of biotin-dependent carboxylases: PCC, 3-methylcrotonyl-CoA carboxylase, pyruvate carboxylase, and acetyl-CoA carboxylase. Biotin in holocarboxylases was probed with streptavidin peroxidase, using Western blots of Jurkat cell extracts. Please note that the biotin-containing 
-chains of PCC (molecular mass = 80 kDa) and 3-methylcrotonyl-CoA carboxylase (molecular mass = 83 kDa) migrate as one single band on polyacrylamide gels.
Cellular content of holo-PCC, holo-3-methylcrotonyl-CoA carboxylase and holo-pyruvate carboxylase paralleled the levels of biotin in culture media and PCC activities. After 4 wk of culturing in biotin-deficient medium, holocarboxylases were barely detectable in cell extracts (Fig. 1
). Levels of holocarboxylases were greater in cells that were cultured in medium containing physiologic concentrations of biotin; levels of holocarboxylases were greatest in cells that were cultured in medium containing pharmacologic concentrations of biotin. Acetyl-CoA carboxylase was not detectable in Jurkat cells in any of the treatment groups. Thus, PCC activities and levels of holocarboxylases are consistent with the hypothesis that biotin supply affects activities of biotin-dependent metabolic pathways in cells.
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Theoretically, cells might respond to biotin deficiency with increased expression of biotin transporter genes to increase cellular accumulation of biotin. Thus, transport rates of biotin were quantified in Jurkat cells that had been cultured in media containing either deficient, physiologic or pharmacologic amounts of biotin for 4 wk. When cells were cultured in biotin-deficient medium, transport rates of biotin increased to 297 ± 79% of controls (physiologic biotin medium = 100%; Fig. 2
). Transport rates of biotin were similar in cells that were cultured in media containing physiologic and pharmacologic concentrations of biotin, respectively. These data are consistent with the hypothesis that Jurkat cells respond to biotin deficiency with increased rates of biotin uptake.
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); human lymphocytes also express this transporter (10
). In this study, levels of SMVT protein in Jurkat cells were quantified by Western blotting, using an antibody to human SMVT. Levels of SMVT protein were similar among treatment groups (data not shown). These findings suggest that increased rates of biotin uptake into biotin-deficient cells (Fig. 2)
were not mediated by increased expression of SMVT but by increased expression of other biotin transporters.
Biotinylation of histones H1, H2A, H2B, H3 and H4 was similar in the treatment groups after 4 wk of culturing in media containing various concentrations of biotin (Fig. 3
). Please note that equimolar amounts of histones from the three treatment groups were probed for biotin (see the Materials and Methods section). These data suggest that, compared with biotinylation of carboxylases, biotinylation of histones is less susceptible to variations in cellular biotin status.
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). This study provided evidence that expression of the biotinidase gene was not affected by cellular biotin supply (see the Biotin-dependent carboxylases section). The mechanism by which biotin-deficient Jurkat cells maintained biotinylation of histones without increasing expression of the biotinidase gene remains uncertain (see the Discussion section).
We determined whether biotin supply affects the rates of cell proliferation and cell death in Jurkat cells. Rates of cell proliferation (as judged by cellular uptake of 5-bromo-2'-deoxyuridine) did not differ among treatment groups after 4 wk of culturing in media containing various concentrations of biotin [units = absorbance at 450 nm/(50,000 cells x h)]: 1.3 ± 0.2 (at 25 pmol/L of biotin) vs. 1.0 ± 0.2 (at 250 pmol/L of biotin) vs. 1.3 ± 0.1 (at 10,000 pmol/L of biotin; P > 0.05). Similarly, proliferation rates were not significantly different among treatment groups when assessed by cellular uptake of [3H]thymidine [units = pmol of thymidine/(200,000 cells x5 h)]: 2.8 ± 0.9 (at 25 pmol/L of biotin) vs. 3.0 ± 0.5 (at 250 pmol/L of biotin) vs. 3.1 ± 0.5 (at 10,000 pmol/L of biotin; P > 0.05). Cells that were cultured in medium containing a pharmacologic concentration of biotin showed a transient increase of proliferation rates during the course of this 4-wk study (Fig. 4
). It is uncertain whether this transient increase of proliferation rates is physiologically meaningful.
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0.5 nmol p-nitroaniline cleaved from DEVD (aspartate, glutamate, valine, aspartate)-p-nitroaniline] (data not shown). As positive control, Jurkat cells were incubated with 50 µmol/L of actinomycin D for 18 h to induce apoptosis. Caspase-3 activity in positive controls was 1.6 nmol p-nitroaniline/(3 x 106 cells x h). These findings are consistent with the hypothesis that the culture conditions used in this study did not affect cell proliferation and apoptosis. Net secretion of interleukin-2.
We determined whether biotin supply affects net secretion of interleukin-2 into the culture medium. After 4 wk of culturing in biotin-defined media, Jurkat cells were stimulated with phytohemagglutinin and phorbol-12-myristate-13-acetate for 24 h to induce secretion of interleukin-2. Secretion of interleukin-2 was inversely associated with the biotin concentration in culture media (Fig. 5
). When cells were cultured in biotin-deficient medium, secretion of interleukin-2 increased to 151 ± 51% of control values (physiologic biotin medium = 100%); when cells were cultured in medium containing pharmacologic concentrations of biotin, secretion of interleukin-2 decreased to 41 ± 7.0% of control values. The mechanism by which biotin affects net secretion of interleukin-2 is uncertain.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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). Moreover, Lewis et al. (28
) reported that biotinylation of carboxylases in rat liver increased when dietary biotin was increased from 0 to 3.69 to 36.9 nmol/d. They also noted that biotinylation of some carboxylases decreased when an excessive amount of biotin was administered to rats (6,150 nmol of biotin/d). Notwithstanding the potential importance of this finding, an amount of 6,150 nmol of biotin per day in rats exceeds doses that are reasonably consumed by humans (if adjusted by body weight). In summary, this study and previous studies suggest that activities of biotin-dependent metabolic pathways may be affected by dietary intake of biotin.
In this study, culture conditions (biotin supply) did not affect biotinylation of histones. Expression of the enzyme that catalyzes biotinylation of histones (biotinidase) was similar among treatment groups; this finding suggests that biotin-deficient cells do not increase expression of the biotinidase gene to maintain normal levels of histone biotinylation. We speculate that cells might use the following mechanisms to maintain biotinylation of histones when biotin supply is changing: cells might respond to biotin deficiency with increased transfer of biotinidase from cytoplasm into the cell nucleus, which harbors the vast majority of cellular histones. In normal cells, 26% of cellular biotinidase activity is located in the cell nucleus (29
). Biotinylation of histones might be mediated not only by biotinidase but also by other, yet unidentified, enzymes. Our previous studies in human mononuclear cells have provided circumstantial evidence that enzymes other than biotinidase may mediate biotinylation of histones (4
). Biotinylation status of histones might be regulated by the rate of histone debiotinylation rather than by the rate of biotinylation.
Which mechanisms are utilized by cells to maintain biotin homeostasis at various levels of biotin supply? We considered the following potential mechanisms: previous studies have shown that biotinidase plays an important role in the recycling of peptide-bound biotin in degradation products of carboxylases (2
). Theoretically, biotin-deficient cells might increase expression of the biotinidase gene to maximize recycling of biotin. However, this study provided evidence that expression of the biotinidase gene was not affected by biotin status, suggesting that cells do not increase recycling of biotin in response to biotin-deficient conditions.
Previous studies have suggested that cells utilize acetyl-CoA carboxylase as storage vehicle for biotin (30
). Theoretically, cells that are cultured in media containing pharmacologic concentrations of biotin might increase expression of carboxylase genes to increase the mass of biotin-binding proteins and, hence, biotin stores. This study and a previous study in rats (27
) provided evidence that biotin supply does not affect expression of genes encoding PCC and pyruvate carboxylase. Moreover, acetyl-CoA carboxylase was below detection limit of Western blot analysis in this study. These studies suggest that expression of genes encoding biotin-binding carboxylases is not increased in response to pharmacologic doses of biotin. Of course, biotinylation of existing carboxylases parallels biotin status.
Previous studies have suggested that biotin transport into human cells is mediated by SMVT (18
) and other, yet unidentified, biotin transporters (17
,19
). Theoretically, biotin-deficient cells might increase expression of genes encoding biotin transporters to increase rates of biotin uptake. Consistent with this hypothesis, transport rates of biotin in biotin-deficient Jurkat cells were 300% of biotin-sufficient controls in this study. This study also supported our previous conclusions that (some) biotin transport into immune cells is mediated by transporters other than SMVT (17
,19
): levels of SMVT protein were similar among treatment groups, despite a significant increase of biotin transport in biotin-deficient cells. We speculate that expression of biotin transporters other than SMVT is increased in response to biotin-deficient conditions. Currently, our laboratory seeks to identify these transporters in human cells.
Does biotin status affect metabolic indicators of immune function? In this study, we used Jurkat cells to determine whether biotin status affects the following variables: net proliferation (proliferation and apoptosis) and net secretion of interleukin-2. Rates of proliferation and apoptosis were not affected by biotin status, suggesting that immune cells maintained normal growth in the presence of deficient or pharmacologic concentrations of biotin. Cells other than immune cells may be more susceptible to changes in biotin status; for example, biotin-depleted HeLa cells arrest in G1 phase of the cell cycle (5
). The reader should note that in this study, the biotin-deficient medium contained 25 pmol biotin/L; effects of biotin-free medium were not investigated because biotin-free nutrition over an extended period is very unusual in the general population.
Cellular biotin status had a significant effect on net secretion of interleukin-2 by Jurkat cells: secretion of interleukin-2 was increased when cells were cultured in biotin-deficient medium and was decreased when cells were cultured in medium containing pharmacologic concentrations of biotin. This finding is consistent with a previous study in our laboratory; in this previous study we provided evidence that supplementation of healthy adults with 3.1 µmol/d of biotin for 14 d caused a 56% decrease of interleukin-2 secretion by mononuclear cells ex vitro (12
). Mechanisms leading to increased interleukin-2 secretion by biotin-deficient cells and decreased interleukin-2 secretion by biotin-supplemented cells remain uncertain. Theoretically, increased concentrations of interleukin-2 in culture medium could be caused either by increased secretion or by decreased internalization of interleukin-2 by Jurkat cells.
Interleukin-2 plays an important role in the immune system, e.g., interleukin-2 stimulates growth and differentiation of T cells, natural killer cells, B cells, and some myeloid cells (31
). Given the important role of interleukin-2 in the immune system, effects of biotin status on interleukin-2 metabolism by immune cells may be physiologically significant. Immune cells of individuals with abnormal intake or metabolism of biotin may secrete (or internalize) abnormal amounts of interleukin-2. Members of the following subgroups of the general population are at increased risk to have abnormal intake or metabolism of biotin: pregnant women may develop marginal biotin deficiency due to accumulation of biotin in the fetal circulation (32
) or accelerated catabolism of biotin to inactive metabolites in maternal tissues (33
). Patients treated with anticonvulsants may have an abnormal metabolism of biotin. For example, the anticonvulsants primidone and carbamazepine inhibit biotin uptake into brush border membrane vesicles from human intestine (34
,35
). Patients with inborn errors of biotin metabolism (e.g., biotinidase deficiency) are treated with large doses (up to 40 µmol/d) of oral biotin (2
,36
). Users of biotin supplements might consume large amounts of the vitamin. Use of biotin supplements is fairly common in the United States: 15–20% of the population reports to take biotin supplements (15
). It remains uncertain as to whether biotin supplementation affects immune competence in persons with viral infections such as HIV or patients with HIV/AIDS.
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FOOTNOTES |
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2 This work was supported by National Institutes of Health Grant DK 60447 and the U.S. Department of Agriculture/National Research Initiative Competitive Grants Program Project Award 2001-35200-10187. A contribution of the University of Nebraska Agricultural Research Division, Lincoln, NE 68583, Journal Series No. 13562.
4 Abbreviations used: PCC, propionyl-CoA carboxylase; PCR, polymerase chain reaction; SMVT, sodium-dependent multivitamin transporter.
Manuscript received 28 November 2001. Initial review completed 10 January 2002. Revision accepted 14 February 2002.
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LITERATURE CITED |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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1. Zempleni, J. & Mock, D. M. (1999) Biotin biochemistry and human requirements. J. Nutr. Biochem. 10:128-138.
2. Wolf, B. & Heard, G. S. (1991) Biotinidase deficiency. Barness, L. Oski, F. eds. Advances in Pediatrics 1991:1-21 Medical Book Publishers Chicago, IL. .
3. Hymes, J., Fleischhauer, K. & Wolf, B. (1995) Biotinylation of histones by human serum biotinidase: assessment of biotinyl-transferase activity in sera from normal individuals and children with biotinidase deficiency. Biochem. Mol. Med. 56:76-83.[Medline]
4. Stanley, J. S., Griffin, J. B. & Zempleni, J. (2001) Biotinylation of histones in human cells: effects of cell proliferation. Eur. J. Biochem. 268:5424-5429.[Medline]
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6. Watanabe, T. (1983) Teratogenic effects of biotin deficiency in mice. J. Nutr. 113:574-581.
7. Watanabe, T., Dakshinamurti, K. & Persaud, T.V.N. (1995) Biotin influences palatal development of mouse embryos in organ culture. J. Nutr. 125:2114-2121.
8. Báez-Saldaña, A., Díaz, G., Espinoza, B. & Ortega, E. (1998) Biotin deficiency induces changes in subpopulations of spleen lymphocytes in mice. Am. J. Clin. Nutr. 67:431-437.[Abstract]
9.
Zempleni, J. & Mock, D. M. (1999) Mitogen-induced proliferation increases biotin uptake into human peripheral blood mononuclear cells. Am. J. Physiol. Cell Physiol. 276:C1079-C1084.
10. Zempleni, J., Stanley, J. S. & Mock, D. M. (2001) Proliferation of peripheral blood mononuclear cells causes increased expression of the sodium-dependent multivitamin transporter gene and increased uptake of pantothenic acid. J. Nutr. Biochem. 12:465-473.[Medline]
11. Zempleni, J. & Mock, D. M. (2000) Utilization of biotin in proliferating human lymphocytes. J. Nutr. 130:335S-337S.
12.
Zempleni, J., Helm, R. M. & Mock, D. M. (2001) In vivo biotin supplementation at a pharmacologic dose decreases proliferation rates of human peripheral blood mononuclear cells and cytokine release. J. Nutr. 131:1479-1484.
13. Mock, D. M. (1997) Determinations of biotin in biological fluids. McCormick, D. B. Suttie, J. W. Wagner, C. eds. Methods in Enzymology 1997:265-275 Academic Press New York, NY. .
14. Mock, D. M., Lankford, G. L. & Mock, N. I. (1995) Biotin accounts for only half of the total avidin-binding substances in human serum. J. Nutr. 125:941-946.
15. Institute of Medicine, Panel on Folate, other B vitamins, and Choline (1998) Dietary Reference Intakes for Thiamin, Riboflavin, Niacin, Vitamin B-6, Folate, Vitamin B-12, Pantothenic Acid, Biotin, and Choline 11th ed. 1998 National Academy Press Washington, DC. .
16.
Zempleni, J., Trusty, T. A. & Mock, D. M. (1997) Lipoic acid reduces the activities of biotin-dependent carboxylases in rat liver. J. Nutr. 127:1776-1781.
17.
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