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,**2
*
Arkansas Children’s Hospital Research Institute and the Departments of
Biochemistry and
**
Pediatrics, University of Arkansas for Medical Sciences, Little Rock, AR 72205
2To whom correspondence should be addressed. E-mail: mockdonaldm{at}uams.edu
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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and interleukin-4 levels, IgG antibody responses and natural killer cell activity were not significantly different in mild to moderately biotin-deficient rats compared with biotin-sufficient controls. The absence of effects on immune function was not attributable to failure to induce biotin deficiency; the rats exhibited unequivocal evidence of biotin deficiency, including reduced hepatic biotin and impaired leucine metabolism resulting from deficiency of the biotin-dependent enzyme methylcrotonyl-CoA carboxylase. We conclude that the immune markers examined are not promising candidates as indicators of mild to moderate deficiency in humans.
KEY WORDS: • immune function • biotin deficiency • rats • marginal deficiency
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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), the specific mechanisms of single nutrient deficiencies and the role played by combined nutrient deficiencies have yet to be fully delineated.
Biotin, a water-soluble vitamin of the B complex, serves as a cofactor of four mammalian enzymes: propionyl-CoA carboxylase, pyruvate carboxylase, methylcrotonyl-CoA carboxylase and acetyl-CoA carboxylase (2
). These carboxylases play important roles in gluconeogenesis, Krebs cycle anapleurosis, branch-chain amino acid catabolism, odd-chain fatty acid catabolism and fatty acid elongation (2
). Because these processes are essential to cell viability, biotin deficiency can affect the immune response. Indeed, substantial effects of moderate to severe biotin deficiency have been documented in rodents (3
–8
). In humans, some clinical aspects of severe biotin deficiency, such as cutaneous fungal infections, suggest that biotin deficiency causes impaired immune function (2
). Moreover, in children suffering from an inborn error that leads to biotin deficiency or from deficiency of multiple biotin-dependent carboxylases, specific T cells and B cell defects have been demonstrated (9
).
Recent clinical studies indicate that marginal asymptomatic biotin deficiency may be a common occurrence in normal human gestation (10
–12
) and in individuals treated for extended periods with certain anticonvulsants (13
–18
). To assess the effects of marginal biotin deficiency on immune function and thereby evaluate immune function as a potential marker for impaired biotin status, we investigated immune function in a rat model during progression from biotin sufficiency to moderate biotin deficiency. As immune function indicators, we assessed the IgG response to a vaccine and the cytokine responses and lymphocyte subpopulations in the immunocytes from blood, spleen and thymus.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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Twenty-two-d-old male Sprague-Dawley rats (Harlan, Indianapolis, IN) were individually housed in stainless steel, wire bottom cages in a room maintained on a 12-h light, 12-h dark cycle. Rats were provided a standard diet (Prolab R-M-H 3000; Agway, Syracuse, NY) for 5 d to stabilize their nutritional status. Rats consumed water ad libitum. This research was approved by the University of Arkansas for Medical Sciences’ Animal Care and Use Committee.
On day 1 of the study, rats were randomly assigned to the biotin-deficient or biotin-sufficient group. The deficient group received a diet containing 30 g of egg white solids per 100 g diet (TD 87400; Harlan Teklad, Madison, WI). This quantity of egg white has been shown to reliably produce biotin deficiency in rats (19
,20
). The biotin-sufficient group received a diet containing 30 g of egg white solids per 100 g diet to which sufficient biotin had been added to prevent deficiency (TD 97197; Harlan). Biotin-sufficient rats were given enough food each day to ensure that they maintained the same weight gain as the deficient group. Adjustment of food intake of the control group was necessary because mild to moderate biotin deficiency can decrease food intake and weight gain and because in many animal species including the rat, a variety of immune functions are influenced by even moderate reductions in food intake.
Before killing on days 1, 7, 14, 28, and 40, urine was collected for 24 h while rats were deprived of food. After processing, urine was stored at -20°C until analysis. Heparinized blood from the inferior vena cava vein, liver, spleen and thymus were harvested. Because blood was the limiting factor in having sufficient material for all studies, enough rats were killed at each time point to pool the blood and to make five samples. This required two rats per blood sample at days 1, 7 and 14, and one rat per sample at day 28 and day 40. Livers were perfused with 154 mmol/L NaCl before being frozen in liquid nitrogen. Livers were stored at -70°C until assayed for biotin.
Indices of biotin status.
Biotin status in the rats was assessed by measurement of hepatic biotin content and urinary 3-hydroxyisovaleric acid (3HIA)3
excretion. The mass of an aliquot of the frozen liver was determined gravimetrically. The aliquot was immediately homogenized in water using a sintered-glass homogenizer as described previously (21
). Biotin was then released from covalent binding to proteins by acid hydrolysis using 1.5 mol/L HCl as described previously (21
). This method releases >95% of the bound biotin with destruction of <10% of the biotin (22
). Biotin was then separated by HPLC from biotin analogs and other substances that interfere with the avidin-binding assay (23
). The HPLC fractions containing biotin were then assayed directly using an avidin-binding assay as described previously (23
).
Increased 3HIA excretion reflects decreased activity of methylcrotonyl-CoA carboxylase, a biotin-dependent enzyme. Urinary excretion of 3HIA has been shown to be an early and sensitive indicator of biotin deficiency in rats (24
) and humans (25
). Urinary 3HIA was quantitated by GC/MS using unlabeled 3HIA as the external standard and deuterated 3HIA as the internal standard (26
).
Biotin-free culture of blood mononuclear cells (BMC).
BMC suspensions were obtained from heparinized blood by density gradient separation. Blood was overlaid onto Fico/Lite-LymphoH, density 1.077 (Atlanta Biologicals, Norcross, GA) and centrifuged at 650 x g for 30 min. The buffy coat layer containing the BMC was removed and sedimented at 200 x g for 10 min at room temperature. The cell pellet containing primarily mononuclear cells was washed three times in RPMI medium. The RPMI medium was specially formulated to omit added biotin (Gibco, Grand Island, NY) with 10 mL/L biotin-free fetal calf serum. Because up to 41 nmol/L of biotin is present in normal fetal calf serum (27
), we removed biotin from the fetal calf serum before mixing with media; we used avidin (Monomeric)-agarose (Sigma, St. Louis, MO) according to manufacturer’s instructions as follows. A 1.0-mL column of avidin-agarose beads was equilibrated with phosphate buffered saline (PBS), pH 7.0, and 10-mL aliquots were passed over the column bed. Biotin was not detectable in the biotin-depleted serum (<35 pmol/L); concentration of biotin in plasma from nonpurified diet-fed rats is 
5000 pmol/L.
Immunocyte harvest.
Splenocytes and thymocytes were harvested from their respective organs by perfusion and mincing in heparinized RPMI medium. Contaminating erythrocytes were lysed with 1.0 mL of distilled water for 15 s. Cells were washed with the medium and pelleted by centrifugation at 650 x g for 10 min. Cell pellets were washed two times in biotin-free medium. Cells in the final suspension were enumerated using the Coulter Counter ZM (Coulter Corp., Miami, FL). Cell viability was assayed by Trypan blue exclusion; cell viability was >95% for all preparations. Cell concentrations were adjusted to 4.0 x 109/L before assessment of immune function as described below.
Characterization of lymphocyte subpopulations.
Fluorescein-labeled monoclonal mouse antibodies to rat lymphocyte subpopulations (PharMingen, San Diego, CA) were used to quantitate lymphocyte subpopulations; subpopulations included Pan T cells (CD3), T suppressor cells (CD8), T helper cells (CD4) and B lymphocytes (CD45R). Fluorescein-labeled mouse immunoglobulins G1, G2a and G2b were included as nonspecific immunoglobulin binding controls. Thymocytes, splenocytes and BMC subpopulations were incubated with each antibody and suspended at a concentration of 3–6 x 109 cells/L in biotin-free RPMI 1640 media containing 5 mL/100 mL heat-inactivated, biotin-free fetal calf serum with 100 mg/L of streptomycin and 1 x 105 U/L of penicillin. This cell suspension (100 µL) and monoclonal antibody (20 µL) were mixed and incubated for 20 min in the dark. The reaction was terminated by centrifugation. The supernatant was removed; the pellet was washed twice in PBS and fixed in 0.5 mL of 10 mL/L formaldehyde solution in PBS. The lymphocyte subpopulations were enumerated using a Coulter Epics Profile II flow cytometer (Coulter Corp., Hialeah, FL).
Lymphocyte proliferation.
The T cell mitogen phytohemagglutinin (PHA; final concentration of 10 or 20 mg/L) was added to 100 µL of a BMC, thymocyte or splenocyte cell suspension containing 4.0 x 109 cells/L. After incubation for 3 d at 37°C in a 95% O2, 5% CO2, 3H-thymidine uptake was assessed by pulse-labeling with 37 kBq/well of 3H-thymidine for 18 h. Lysed cells were harvested with the Filtermate 196 cell harvester (Packard Instruments, Downers Grove, IL), and DNA was absorbed to a membrane. The 3H bound to DNA was counted with a Matrix 9600 direct ß-counter (Packard Instruments). The blastogenic index was calculated as the ratio of 3H cpm of the PHA-stimulated cells to 3H in the media alone. BMC not exposed to mitogen served as a control.
Cytokine production.
Immunocytes from each of the three sites were stimulated with mitogen for 24 h. The cells were pelleted at 200 x g for 10 min; supernatants were removed and stored at -70°C until assay. The media concentrations of the cytokine interleukin (IL)-4 and interferon- were measured using standardized cytokine assay kits (Biosource International, Camarillo, CA).
Response to active Haemophilus influenzae b conjugate vaccine (ActHIB).
Rats were injected with 100 µL of ActHIB vaccine (2 µg capsular polysaccharide, 5 µg tetanus toxoid; Lederle Laboratories, Pearl River, NY). Immunoglobulin G was assessed using an ELISA-based immunoassay developed in our laboratory. To determine relative levels of antigen-specific IgG, 100 µL of Act HIB vaccine (1 mg/L) in a pH 9.6 carbonate/bicarbonate buffer was adsorbed to wells of a 96-tissue culture plate and incubated at room temperature for 4 h. The tissue culture plates were washed with blocking buffer (PBS, 5.0 g/L Tween 20; pH 7.0) and 100 µL of plasma was added to the ActHIB vaccine coated wells for 1 h at room temperature. The plates were washed in blocking buffer and 100 µL of rabbit alkaline phosphate-labeled anti-rat IgG (Sigma) diluted 1:1000 (v/v) in blocking buffer was added to the wells. After 1 h at room temperature, the plates were washed and alkaline phosphatase substrate (paranitrophenyl phosphate) was added according to ELISA protocol provided by Sigma. Color development was monitored and read at 405 nm using a MicroPlate reader (BioRad, Hercules, CA). To determine the antigen-specific IgG, plasma from preimmunized rats (day 1) was used to establish baseline absorbency. To determine the effect of biotin deficiency on IgG production, the plasma from biotin-sufficient and biotin-deficient rats was analyzed at days 7, 14, 28 and 40. The mean absorbency values from triplicate wells was recorded and compared to determine an in vivo functional response to immunization.
Natural killer cell function.
The ability of effector lymphocytes from blood, spleen and thymus cell suspensions to cause lysis of target tumor cells was measured by release of 51Cr from 51Cr-loaded tumor cells. Effector cells were isolated by density gradient as described above and resuspended in RPMI 1640 media with 10 mL/L fetal calf serum media. The cell suspensions were mixed with equal volumes of dimethyl sulfoxide and stored in liquid nitrogen until immunocytes from both diet groups at all time points were available. Cells were thawed at 37°C and washed three times in media before incubation with the target cells, which had been previously loaded with 51Cr as follows.
YAC-1 cells (TIB 160; American Type Culture Collection, Manassas, VA) from a natural killer-sensitive line were suspended in media at 5 x 109 cells/L; 1 mL of this suspension was incubated with 3.7 MBq Na251Cr04 for 1 h at 37°C. The 51Cr-loaded target cells were washed three times and resuspended at 1 x 109 cells/L.
Effector cells were diluted to 1.25 x 106, 2.5 x 106, and 5 x 109 cells /L; 100 µL/well were added in triplicate to 96-well tissue culture plates to produce effector:target ratios of 12.5:1, 25:1 and 50:1. Loaded target cells (100 µL/well) were added to the effector cells. The cell mixture was incubated in a humidified, 5% CO2 atmosphere at 37°C for 4 h. The released 51Cr (EXP) was separated from the cells and debris by centrifugation of the plate at 50 x g for 5 min with no brake. A 100-µL aliquot of the supernatant was transferred to a 96-well solid scintillant plate and the radioactivity measured using Packard Lumaplates and TopCount microplate scintillation counter (Packard Instruments). Maximum 51Cr release (MR) was measured by lysing 100 µL of loaded target cells with 100 µL of 1 mL/L Triton X100 in RPMI 1640 media. Background radioactivity from spontaneous 51Cr release (SR) was measured in supernatant from 100 µL of loaded target cells alone which were incubated for 4 h at 37°C. The percentage of lysis was calculated as follows:
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Statistics.
For urinary excretion of 3HIA, the significance of differences between deficient and sufficient rats and of trends over time were evaluated by two-way ANOVA (time x treatment) using Statview, version 5.0 (28
). To determine the point at which 3HIA excretion had increased significantly compared with baseline, one-way ANOVA with Dunnett’s posthoc test was performed.
For hepatic biotin, significance of differences between means at day 40 in deficient rats and day 40 in sufficient rats were tested by Student’s t test. The significance of differences between mean hepatic biotin at day 40 in deficient rats and day 1 rats was tested by Student’s t test. A Bonferroni’s correction was applied for a second t test.
For immune function, significance of differences between deficient and sufficient rats were tested using the Student’s t test for pair-wise comparisons. With small sample size, there is a higher chance of type II error (accepting the null hypothesis when it is false); therefore, only P values < 0.1 were accepted as significant.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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Over the 40 d of the study, body weight did not diverge significantly between the biotin-deficient and the biotin-sufficient rats (two-way ANOVA). Likewise, spleen and thymus organ weights did not differ between the biotin-deficient and biotin-sufficient rats (two-way ANOVA). Cutaneous signs of biotin deficiency began to appear by day 28; hair roughened and began to thin. By day 40, partial alopecia was present in 80% of the deficient rats. Neurologic signs of biotin deficiency were apparent by day 40; kangaroo gait was noted in one-third of the deficient rats, and irritability was noted in all deficient rats. Sufficient rats showed no signs of deficiency.
Quantitative indicators of biotin status.
For the deficient rats, hepatic biotin decreased substantially from day 1 by day 28 (3.90 ± 0.24 vs. 0.74 ± 0.11 nmol/g liver; P < 0.0001) and was significantly less than that of the sufficient group (8.17 ± 0.56 nmol/g liver; P < 0.0001). We also observed a progressive increase in the urinary 3HIA excretion (Fig. 1
). This is the result of a progressive deficiency of the biotin-dependent enzyme methylcrotonyl-CoA carboxylase (21
). The differences in 3HIA excretion were significant for treatment (P < 0.0001), time (P < 0.0001) and for the interaction of treatment and time (P < 0.0001). As early as day 14, the mean urinary excretion of 3HIA was already significantly increased compared with day 1 (P < 0.01, one-way ANOVA with Dunnett’s posthoc test). In biotin-sufficient controls, urinary excretion of 3HIA did not change significantly.
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For Pan T cells (Fig. 2
) and CD4 lymphocytes (Fig. 3
), the subpopulation distribution differed significantly between biotin-sufficient and biotin-deficient rats during the course of the experimental protocol in the blood and thymus, and there was a trend for significant differences in the Pan T cells and CD4 lymphocytes in the spleen cell population at days 28 and 40 (Pan T: P = 0.06 and 0.01, respectively; CD4: P = 0.02 and 0.01, respectively). In the CD8 lymphocyte subpopulations (Fig. 4
;F4>) in the blood and thymus, biotin-sufficient and biotin-deficient rats tended to differ at day 40 (P = 0.05 and P = 0.07, respectively). In the splenic CD8 subpopulation, diet groups did not differ. B cells also did not differ (P > 0.20) between groups during the 40-d trial (data not shown). Thus, there did not seem to be an important redistribution of the lymphocyte cell population among the tissues during this mild to moderate biotin-deficient state over the 40-d trial.
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In BMC, splenocytes and thymocytes, the T cell proliferation response to PHA was not significantly different between the biotin-sufficient and -deficient groups (Fig. 5
; P > 0.25) in all of the cell populations tested with a single exception. Proliferation of the spleen cell population tended to differ (P = 0.08) between the biotin-sufficient and biotin-deficient rats at day 28.
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levels.
Interferon- levels were determined from a standard curve ranging from 24 to 1500 ng/L. Interferon- levels in PHA-stimulated peripheral blood cultures of the biotin-sufficient rats were 288 ± 224 ng/L, 655 ± 418 ng/L, 81 ± 68 ng/L and 22 ± 10 ng/L during days 7, 14, 28, and 40 of the experimental design. By comparison, in biotin-deficient rats, the interferon- levels were 156 ± 126 ng/L, 992 ± 422 ng/L, 166 ± 100 ng/L and 32 ± 21 ng/L. Spleen interferon- levels were 2580 ± 745, 174 ± 147, 306 ± 248 and 144 ± 267 ng/L in biotin-sufficient rats and 1087 ± 256, 156 ± 225, 491 ± 258 and 34 ± 26 ng/L in biotin-deficient rats during the respective periods. None of the differences between the two groups was significant.
IL-4 levels.
IL-4 levels in supernatants from the PHA-stimulated peripheral blood and spleen cell cultures were determined at days 7, 14, 28, and 40 from a standard curve ranging from 22 to 1400 ng/L. In the peripheral blood, IL-4 was uniformly 94 ng/L for biotin-deficient and -sufficient groups. In the spleen cultures, IL-4 was uniformly 24 ng/L in sufficient and deficient groups with one exception; at day 28 in the biotin-deficient rats, the IL-4 level reached 70 ng/L in spleen cultures.
Vaccine response.
Immunization with the ActHIB vaccine at day 1 of the study resulted in a limited antigen-specific IgG production. Optical density measurements of IgG levels did not surpass a two-fold increase over background in the biotin-deficient rats throughout the study. At day 14, the biotin-sufficient rats reached a 2.8-fold increase above background. There was no difference in the optical density for IgG production during the study, and levels were only moderately elevated above background.
Natural killer cell activity.
When assessing the percentage of lysis at the different effector target ratios, there was a two-fold to three-fold increase in natural killer cell activity with a modest increase in killer cell activity over the experimental time frame (days 7, 14, 28, and 40). For example, in the peripheral blood, at days 7 and 14; at 12.5:1 the percentage of lysis was 11%; at 25:1, 19%; and at 50:1, 31% in the biotin-sufficient group and 11%, 18% and 34%, respectively, in the biotin-deficient group. By day 40, the levels increased to 16%, 19% and 34%, respectively, in the biotin-sufficient group, and 12%, 22% and 31% in the biotin-deficient group. Similar increases, although not identical, for the percentage of lysis determinations in the spleen and thymus were in evidence. Difference were not significant at any time point or effector:target ratio for natural killer cell activity in peripheral blood, spleen or thymus cultures.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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These minor changes in total cell populations and subcell populations in organs did not have an important effect on T cell lymphocyte function as assessed by lymphocyte proliferation and lymphocyte cytokine response to the T cell mitogen, PHA. Likewise, antigen-specific IgG response to Haemophilus B polysaccharide conjugate was not different between biotin-deficient and biotin-sufficient rats at any point in the study. Activities of natural killer cells from blood, thymus and spleen against the YAC-1 cell line were not affected by mild to moderate biotin deficiency.
The absence of deficiencies in immune function cannot be attributed to failure to induce biotin deficiency. The rats in the biotin-deficient group did indeed develop biotin deficiency. The deficient rats were mildly deficient for the first 3 wk of the study based on the observation that disturbances in leucine metabolism, due to deficiency of methylcrotonyl CoA carboxylase, were present, but no signs of biotin deficiency had yet appeared. In this regard, the onset of symptoms and increase of 3HIA excretion parallels the first 3 wk of egg white feeding in a study in which biotin deficiency was experimentally induced in healthy adults (25
). As judged both by hepatic biotin and by the presence of cutaneous and neurologic symptoms, by day 40 the rats were at least moderately biotin deficient. The discordance between the presence of cutaneous symptoms and the absence of overt immune difference in this study at day 40 suggests that immune dysfunction probably does not play an initial role in the cutaneous manisfestations of biotin deficiency. This inference is consistent with the results of a dietary interaction experiment (21
), which demonstrated that intraperitoneal supplementation of (n-6) fatty acids almost completely prevented the cutaneous manifestations of biotin deficiency during 13 wk of egg white feeding in rats. Notwithstanding, when biotin deficiency progresses to a more severe form, immune function is impaired (2
,9
) with secondary Candida infection playing a prominent role in the cutaneous manifestations.
Striking immune effects have been reported in biotin-deficient rats (3
,4
,7
). These studies reported physical signs of biotin deficiency but did not quantitate status. As judged by the length of egg white feeding and the reported physical findings of deficiency, biotin deficiency was more severe in these studies; differences in reported effects on immune function likely arose from the differences in severity of biotin deficiency.
Our results are different than, but not inconsistent with, those of Báez-Saldaña et al. (8
) in biotin-deficient mice. These investigators concluded that biotin deficiency in mice can have a profound effect on lymphocyte maturation and responsiveness to stimulation. During 7–20 wk of egg white feedings, they produced severe biotin deficiency and weight loss. They observed important effects on the production and maturation of both T and B cells. CD3, CD4 and CD8 splenic lymphocytes were significantly increased beginning at wk 4, and B cells decreased beginning at wk 12. Although the percentages of T cells increased with biotin deficiency, the functional activity was decreased by 20–25% in spleen cell proliferative assays, consistent with improper development of lymphoid cells. Although differences in severity and perhaps differences in species are likely sources of the different results of these two studies, the fact that the mice in that study lost weight adds malnutrition as another factor potentially explaining the differences in results between the studies.
In summary, these studies did not identify any immune functions that are good indicators of mild to moderate deficiency in rats. Because biotin status was documented in studies reported here, comparison can be made to clinical circumstances in which biotin deficiency develops (e.g., pregnancy and anticonvulsant therapy) or is deliberately induced (egg white feeding). The proportional decrease of biotin excretion from the mean of the normal range and the proportional increase in 3HIA excretion from the mean of the normal range in these rats are similar to those reported in pregnancy (11
) and in egg white feeding (26
). To the extent that the effects on immune function can be extrapolated from rats to humans at a similar degree of biotin deficiency, immune function is not likely to be seriously impaired and does not seem to be a promising marker for decreased biotin status. Because mild biotin deficiency seems to be common in pregnancy, the apparent absence of immune impairment by mild deficiency is reassuring.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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3 Abbreviations used: ActHIB, active Haemophilus influenzae conjugate vaccine; BMC, blood mononuclear cell; 3HIA, 3-hydroxyisovaleric acid; IL, interleukin; PBS, phosphate buffered saline; PHA, phytohemagglutinin.
Manuscript received June 21, 2001. Initial review completed July 26, 2001. Revision accepted September 27, 2001.
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LITERATURE CITED |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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1. Long, K. Z. & Santos, J. I. (1999) Vitamins and the regulation of the immune response. Pediatr. Infect. Dis. J. 18:283-290.[Medline]
2. Mock, D. M. (1996) Biotin. Ziegler, E. E. Filer, L. J., Jr eds. Present Knowledge in Nutrition 1996:220-235 International Life Sciences Institutes Nutrition Foundation, Washington, DC. .
3. Pruzansky, J. & Axelrod, A. E. (1955) Antibody production to diphtheria toxoid in vitamin deficiency states. Proc. Soc. Exp. Biol. Med. 89:323-325.
4. Kumar, M. & Axelrod, A. E. (1978) Cellular antibody synthesis in thiamin, riboflavin, biotin and folic acid-deficient rats. Proc. Soc. Exp. Biol. Med. 157:421-423.[Medline]
5. Kung, J. T., MacKenzie, C. G. & Talmage, D. W. (1979) The requirement for biotin and fatty acids in the cytotoxic T-cell response. Cell Immunol 48:100-110.[Medline]
6. Petrelli, F., Moretti, P. & Campanati, G. (1981) Studies on the relationships between biotin and the behavior of B and T lymphocytes in the guinea-pig. Experientia 37:1204-1205.[Medline]
7. Rabin, B. S. (1983) Inhibition of experimentally induced autoimmunity in rats by biotin deficiency. J. Nutr. 113:2316-2322.
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. Cowan, M. J., Wara, D. W., Packman, S., Yoshino, M., Sweetman, L. & Nyhan, W. (1979) Multiple biotin-dependent carboxylase deficiencies associated with defects in T-cell and B-cell immunity. Lancet 2:115-118.[Medline]
10.
Mock, D. M., Stadler, D., Stratton, S. & Mock, N. I. (1997) Biotin status assessed longitudinally in pregnant women. J. Nutr. 127:710-716.
11. Mock, D. M., Mock, N. I., Stratton, S. L., Heird, G. M. & Stadler, D. D. (1996) Increased excretion of 3-hydroxyisovaleric acid provides evidence of biotin deficiency during pregnancy. FASEB J 10:A464.
12. Mock, D. M. & Stadler, D. D. (1997) Conflicting indicators of biotin status from a cross-sectional study of normal pregnancy. J. Am. Coll. Nutr. 16:252-257.[Abstract]
13. Krause, K.-H., Berlit, P. & Bonjour, J.-P. (1982) Impaired biotin status in anticonvulsant therapy. Ann. Neurol. 12:485-486.[Medline]
14. Krause, K.-H., Berlit, P. & Bonjour, J.-P. (1982) Vitamin status in patients on chronic anticonvulsant therapy. Int. J. Vitam. Nutr. Res. 52:375-385.[Medline]
15. Krause, K.-H., Bonjour, J.-P., Berlit, P., Kynast, G., Schmidt-Gayk, H. & Schellenberg, B. (1988) Effect of long-term treatment with antiepileptic drugs on vitamin status. Drug Nutr. Interact. 5:317-343.[Medline]
16. Mock, D. M. & Dyken, M. E. (1995) Biotin deficiency results from long-term therapy with anticonvulsants. Gastroenterology 108:A740.
17.
Mock, D. M. & Dyken, M. E. (1997) Biotin catabolism is accelerated in adults receiving long-term therapy with anticonvulsants. Neurology 49:1444-1447.
18. Mock, D. M., Mock, N. I., Lombard, K. A. & Nelson, R. P. (1998) Disturbances in biotin metabolism in children undergoing long-term anticonvulsant therapy. J. Pediatr. Gastroenterol. Nutr. 26:245-250.[Medline]
19. Mock, D. M. (1988) Evidence for a pathogenetic role of fatty acid (FA) abnormalities in the cutaneous manifestations of biotin deficiency. FASEB J 2:A1204.
20. Kramer, T. R., Briske-Anderson, M., Johnson, S. B. & Holman, R. T. (1984) Effects of biotin deficiency on polyunsaturated fatty acid metabolism in rats. J. Nutr. 114:2047-2052.
21.
Mock, D. M. (1990) Evidence for a pathogenic role of 
6 polyunsaturated fatty acid in the cutaneous manifestations of biotin deficiency. J. Pediatr. Gastroenterol. Nutr. 10:222-229.[Medline]
22.
Mock, D. M. & Malik, M. I. (1992) Distribution of biotin in human plasma: most of the biotin is not bound to protein. Am. J. Clin. Nutr. 56:427-432.
23. Mock, D. M. (1997) Determinations of biotin in biological fluids. McCormick, D. B. Suttie, J. W. Wagner, C. eds. Methods in Enzymology 279:265-275 Academic Press New York, NY. part I.[Medline]
24. Mock, N. I. & Mock, D. M. (1989) Use of an improved method for determination of urinary 3-hydroxyisovaleric acid allows earlier detection of biotin deficiency in the rat. Clin. Res. 37:955(abs.).
25.
Mock, N., Malik, M., Stumbo, P., Bishop, W. & Mock, D. (1997) Increased urinary excretion of 3-hydroxyisovaleric acid and decreased urinary excretion of biotin are sensitive early indicators of decreased status in experimental biotin deficiency. Am. J. Clin. Nutr. 65:951-958.
26. Mock, D. M., Jackson, H., Lankford, G. L., Mock, N. I. & Weintraub, S. T. (1989) Quantitation of urinary 3-hydroxyisovaleric acid using deuterated 3-hydroxyisovaleric acid as internal standard. Biomed. Environ. Mass Spectrom. 18:652-656.[Medline]
27. Moskowitz, M., Cheng, D., Moscatello, D. K. & Otsuka, H. (1980) Growth stimulation of BHK cells in culture by free biotin in serum. J. Natl. Can. Inst. 3:639-643.
28. SAS Institutes (1999) Statview, Version 5 1999 Cary, NC SAS Institute .
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