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© 2004 The American Society for Nutritional Sciences J. Nutr. 134:1970-1977, August 2004

Nutritional Immunology

Biotin Deficiency Blocks Thymocyte Maturation, Accelerates Thymus Involution, and Decreases Nose-Rump Length in Mice1

Armida Báez-Saldaña and Enrique Ortega2

Departamento de Inmunología, Instituto de Investigaciones Biomédicas, Universidad Nacional Autónoma de México, México

2To whom correspondence should be addressed. E-mail: ortsoto{at}servidor.unam.mx.


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 LITERATURE CITED
 
Biotin deficiency in experimental animals causes low body weight as well as several phenomena suggestive of an altered immune system. We reported previously that chronic biotin deficiency in mice decreases body weight and alters the number and proportion of lymphocyte subpopulations in the spleen. To further characterize the effects of biotin deficiency, we studied in detail the maturation of thymocytes and the status of biotin in the thymus, as well as the body length of biotin-deficient mice. Male Balb/cAnN mice were fed for up to 20 wk either standard control diet, a biotin-deficient diet, or a biotin-sufficient diet. At different times, nose-rump length, weight of the thymus, spleen and liver, total number of cells in the spleen and thymus, pyruvate carboxylase (PC) and propionyl CoA carboxylase (PCC) activity in thymus cells, and the proportion of distinct thymocyte subsets were determined. These variables did not differ between mice fed the control and biotin-sufficient diets. In contrast, biotin-deficient mice differed from biotin-sufficient mice in all of the analyzed variables. PC and PCC specific activities of thymocytes of mice fed the biotin-depleting diet decreased during the first 4 wk by 84.5%. The maturation of thymocytes in biotin-deficient mice was arrested at the double-negative stage. Our results suggest that biotin deficiency in mice causes an accelerated involution of the thymus and decreases nose-rump length, but these effects do not correlate in magnitude or in temporality with the sharp decrease in the activity of the biotin-dependent carboxylases. As such, the possibility that the aforementioned effects are not related directly to the prosthetic function of biotin should be considered.

KEY WORDS: • biotin deficiency • thymocyte maturation • propionyl CoA carboxylase • pyruvate carboxylase • thymus involution

Although many studies have been published about the direct relation between malnutrition and susceptibility to infections, there are fewer reports that address the role of a particular nutritional factor and its effect on the immune system.

There is evidence that micronutrients such as Zn (1,2) and vitamins such as A, B-6, pantothenic acid, thiamin, and biotin play important roles in immune processes (2,3). The vitamin biotin is a member of the B complex, and it acts as a prosthetic group of CO2-fixing enzymes (carboxylases) in cells from animals and plants. Inside the cell, the biotinylated enzymes acetyl-CoA carboxylase (ACC),3 methylcrotonyl CoA carboxylase (MCC), propionyl CoA carboxylase (PCC), and pyruvate carboxylase (PC) participate in the metabolism of fatty acids and amino acids, as well as in gluconeogenesis (4).

Gompertz (5) described for the first time an innate error of biotin metabolism known as multiple carboxylase deficiency. Individuals affected by this disease cannot recycle biotin due to genetic defects in either biotinidase or holocarboxylase synthetase. Clinically, such patients have a poor response to immunization with different antigens, and are prone to severe infections by diverse pathogens that cannot be controlled by chemotherapy and that can lead to death (6).

Biotin deficiency in rats decreases the weights of the body, spleen, thymus, and mesenteric lymph nodes (7). Biotin-deficient rats are also less likely to develop experimentally induced autoimmunity (8) and are prone to present lesions in the lungs with accumulation of eosinophils and foam cells (9). In guinea pigs, biotin deficiency changes the proportions of circulating leukocytes, with a reduced number of B and T lymphocytes and a higher percentage of neutrophils (10).

We reported previously that during the first 4 wk of consuming a biotin-depleting diet, experimental mice do not reach the same corporal weight as that of age- and gender-matched control mice; in fact, they lose weight after 6 wk of consuming such a diet (11). Moreover, biotin deficiency changes the subpopulations of spleen lymphocytes and diminishes the proliferative response of splenocytes to Concanavalin A (11). The mechanisms by which biotin deficiency causes these alterations are still unknown. However, these effects are not directly correlated either in time of appearance or in magnitude with the decrease in the activity of PC and PCC. In the work reported herein, we further investigated the effect of biotin deficiency on immune system homeostasis by studying the effect of chronic biotin deficiency on the thymus, a primary lymphoid organ.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 LITERATURE CITED
 
    Mice. The handling of mice at all stages of experimentation was performed in compliance with the standard procedures established by the Ethical Committee for Experimentation with Animals of the Biomedical Research Institute of the National University of Mexico.

Male Balb/cAnN Hsd mice were obtained from the breeding colony of our Institute. Mice were 4 wk old at the beginning of the experiment. They were maintained in barrier conditions with 12-h light:dark cycles, and were allowed to consume water and food ad libitum. Five lots of 50 to 80 mice were each divided into 3 experimental groups (control, sufficient, and deficient) of 15 to 30 mice. Each group was fed 1 of the 3 alternative diets previously described (11,12). Accordingly, 1 group (designated as the control group) was fed a commercial standard diet for experimental rodents (LM-485, Cat. T.7012.15, Harlan Teklad). Mice in the biotin-deficient group were fed a biotin-depleting diet lacking biotin and containing 30% dried egg white as the protein source (TD-01363, Harlan Teklad) (Table 1). Egg white contains avidin, a glycoprotein that binds biotin, forming a noncovalent complex that is not absorbed into the blood (4). The quantity of egg white contained in this diet was shown to reliably produce biotin deficiency in mice and rats (11–15). Finally, the biotin-sufficient group was fed a diet with the same composition as the biotin-depleting diet, but supplemented with 16.4 µmol biotin/kg (TD-01362, Harlan Teklad). This amount of biotin is sufficient to occupy all of the biotin-binding sites on avidin, and still provide enough biotin to meet the metabolic requirements (11,12,16).


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  		TABLE 1 Diets

 
Three to 5 mice from each group were analyzed at the beginning of the experimentation (time 0), and at wk 2, 4, 8, 12, 16, and 20 after initiation of the experiment. Mice were weighed and bled from the axillaries’ plexus under diethyl ether anesthesia. The nose-rump length was measured according to Hughes and Tanner (17) with slight modifications. Each mouse was placed over a metallic ruler, with the 4 extremities extended to the respective sides at approximately right angles, and it was measured from the distal point of the nose to the end of the buttocks. After the length of each mouse was determined, they were killed by cervical dislocation, and the liver, spleen, and thymus were obtained. Each organ was weighed, and livers were frozen at –70°C until they were used to measure the enzymatic activity of PCC and PC. Spleens and thymuses were employed for the determinations indicated in the next sections. To determine whether the 3 organs were equally affected by the corporal weight loss induced by the deficiency of bioti (11), we calculated the ratios of the weight of each organ over the respective corporal weight for each mouse.

    Cellularity of spleens and thymuses. The spleen and thymus obtained from each mouse killed were gently dispersed to single-cell suspensions in PBS containing 5% FBS (Gibco BRL). An aliquot of each cell suspension was used to determine the number of leukocytes per liter using a Beckman Coulter T890 automatic cell counter. From this point onward, only thymus cell suspensions were used. The thymocyte suspension was treated with hemolyzing solution (18), and flow cytometry was used to determine the percentage of cells expressing different cell-surface antigens in an aliquot of the suspension. The remaining thymocyte suspension was centrifuged at 400 x g for 5 min at 4°C, the supernatant was decanted, and the pelleted cells were stored at –70°C until used for determination of the enzymatic activity of PC and PCC.

    Determination of specific activity of PC and PCC in thymocytes. The thymocytes, which had been kept at –70°C, were thawed immediately before the assay. The specific activity of PC and PCC was determined by a radioenzymatic method using NaH14CO3 as substrate, as previously described (11,19). The same method was used to monitor the biotin status in livers of some mice at different times during the experiments by measuring the activities of PC and PCC.

    Cytofluorometric analyses. Our procedure was based on a previously published protocol (20). Thymocytes in single-cell suspensions were incubated with 1 or with a combination of 2–4 of the following mAbs: fluorescein isothiocyanate (FITC)-L3T4 antigen of differentiation expressed on T lymphocytes (CD4) (Gibco/BRL); FITC-CD4 (L3T4), FITC-Ly-2 antigen of differentiation expressed on the surface of T lymphocytes (CD8a); phycoerytrin (PE)-CD8a; peridinin chlorophyll protein (PerCP)-{epsilon} chain of the T-cell receptor-associated CD3 complex (CD3{epsilon}); PE-{alpha} chain of the interleukin 2 receptor (CD25); peridinin chlorophyll protein-cyanin 5.5 (CyChrome)-glycoprotein expressed on hematopoietic and nonhematopoietic cells (CD44) (the last 6 were obtained from Pharmingen). Thymocytes were incubated for 30 min at 4°C with the corresponding antibodies, and were then washed and fixed in paraformaldehyde (2% in PBS, pH 7.4). Fixed cells were kept at 4°C in the dark and analyzed within the next 48 h in a FACScan cytometer (Becton Dickinson) using CellQuest software. The cytometer sensitivity was adjusted using cells incubated with an unrelated antibody of the same isotype as the corresponding specific antibody. In experiments in which cells were stained simultaneously with 2 or more conjugated antibodies, the fluorescence values were compensated with the corresponding single-fluorophore staining (20). Analyses were concluded after recording 10,000 events for each sample.

To define more precisely the substage at which the maturation of thymocytes is halted in biotin-deficient mice, thymocytes from mice of the 3 groups at wk 16 and 20 were stained simultaneously with CD4-FITC, CD8-FITC, CD25-PE, and CD44-CyChr.

    Statistical analysis. Each set of data presented in the Results section is the combined data from mice in at least 2 different lots of mice studied. Data were analyzed by 2-way ANOVA (diet x time) and post-hoc Tukey tests. Differences were considered significant at P < 0.05. SIGMASTAT 2.03 software was used for this analysis (Jandel Scientific Software).


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 LITERATURE CITED
 
Despite the differences between the composition of the Harlan LM-485 commercial standard diet for mice (control diet) and the diet with egg white and supplemented with biotin, mice fed these diets did not differ in any of the variables measured. Nose-rump length, and cellularity and weight of thymus and spleen in mice fed these 2 diets were consistent with those reported for age-matching mice of different strains (21–26).

    Nose-rump length of mice. The increase in nose-rump length during the first 8 wk of the study did not differ among the 3 groups (Fig. 1). From wk 8 to 20, the nose-rump length of the control and of the biotin-sufficient mice did not change significantly. In contrast, the nose-rump length of the biotin-deficient mice decreased from wk 8 until the end of the study, suggesting that biotin deficiency might affect bone remodeling in a different way than it affects the overall body weight.



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  		FIGURE 1 Nose-rump length of mice fed a commercial standard diet (control group), biotin-sufficient, or biotin-deficient diets. Values are means ± SD, n = 6–15. Effects of diet (P < 0.001), time (P < 0.001), and their interaction were significant (P < 0.001). Letters indicate differences from the control and biotin sufficient groups: a, P < 0.01; b, P < 0.001.

 
    Weight of spleen, liver, and thymus. The weight of the spleen of mice fed control and biotin-sufficient diets remained constant throughout the study (Fig. 2A). However, from wk 8 until the end of the study, the weight of the spleen of the mice fed the biotin-depleting diet was significantly lower than that of the mice fed the control and biotin-sufficient diets (Fig. 2A). The weight of the spleen was not affected by time (P = 0.730), but it was affected by diet (P < 0.001), and by the interaction between diet and time (P < 0.001). From wk 0 to 8, the weight of the thymus decreased at the same rate in the 3 experimental groups (Fig. 2B). Nonetheless, from wk 12 onward, the weight of the thymus of the biotin-deficient mice was significantly lower than those of control and sufficient mice (Fig. 2B). The weight of the thymus was affected (P < 0.001) by time, diet, and their interaction. During the first 4 wk, the weight of the liver of control and biotin-sufficient mice increased and after wk 4, it remained constant (Fig. 2C). In contrast, the liver of the biotin-deficient mice did not grow at all, maintaining its initial weight until wk 16, and decreasing in weight from that point on until wk 20 (Fig. 2C). The weight of the liver was affected by time and by diet (P < 0.001), and by their interaction (P = 0.002).



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  		FIGURE 2 Weights of spleen (A), thymus (B), and liver (C) of mice fed a commercial standard diet (control group), biotin-sufficient, or biotin-deficient diets. Values are means ± SD, n = 6–15 (spleen and thymus) and 9–33 (liver). (A) Letters indicate differences between the biotin-deficient group and a, the control group, P < 0.05; b, the control and biotin-sufficient groups, P < 0.001. (B) Letters indicate differences from the control and the biotin-sufficient groups: a, P < 0.05; b, P < 0.001. (C) The letter "a" indicates differences between the biotin-deficient group and the control and biotin sufficient groups, P < 0.001.

 
From wk 8 onward, the ratios of spleen weight to body weight were significantly lower in mice fed the biotin-depleting diet than in mice fed the control and sufficient diets. Moreover, from wk 16 on, the ratios of thymus weight to body weight in mice fed the biotin-depleting diet were also significantly lower than the ratios in mice fed the control and biotin-sufficient diets. In contrast, the ratios of liver weight to body weight did not change significantly during the 20 wk of experimentation for any of the 3 groups and not differ among the 3 diet groups (data not shown). Thus, we conclude that the spleen and the thymus are more sensitive to biotin depletion than the liver, which maintained its relative weight throughout the 20 wk of biotin deficiency.

    Cellularity of spleen and thymus. Like the weight of the spleen, the total number of cells in the spleen of mice fed control and biotin-sufficient diets remained constant throughout the study (Fig. 3A). The number of splenocytes in the biotin-deficient mice decreased during the experiment and from wk 8 on, it was significantly lower than that in the control and biotin-sufficient mice. The number of splenocytes was affected (P < 0.001) by time and diet, and by their interaction. The number of thymocytes decreased asymptotically during the 20 wk of experimentation for all 3 experimental groups. Notwithstanding the above, the number of thymocytes in the biotin-deficient mice at the end of the 20 experimental weeks was only 10% of the number of thymocytes in control and sufficient mice (Fig. 3B). The number of thymocytes was affected (P < 0.001) by time, diet, and their interaction.



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  		FIGURE 3 Total number of cells in the spleen (A) and thymus (B) of mice fed a commercial standard diet (control group), biotin-sufficient, or biotin-deficient diets. Values are means ± SD, n = 3–9. Letters indicate differences from the control and the biotin-sufficient groups: a, P < 0.05; b, P < 0.001; c, P < 0.004.

 
Interestingly, the ratio of the number of leukocytes to the weight of the spleen in biotin-deficient mice had decreased significantly only at wk 16 and at 20 (data not shown), whereas the ratio of the number of thymocytes to thymus weight was significantly lower for biotin-deficient mice than for mice in the other 2 groups after only 2 weeks (data not shown). Hence, we conclude that this ratio is an early indicator of biotin deficiency.

    Specific activity of PC and PCC in thymus. The specific activities of PC and PCC in the liver of mice from the 3 experimental groups were similar to those we reported previously (data not shown) (11). The specific activity of PC and PCC in the thymus of control and biotin-sufficient mice remained constant during the 20 wk of experimentation (Fig. 4). Additionally, they were within the range that we previously reported in spleen cells (11). The specific activities of PC and PCC in thymocytes of biotin-deficient mice decreased significantly during the first 4 wk of the study (Fig. 4). For PC and PCC activity in the thymus, there was a significant effect of time (P < 0.001), diet (P < 0.002), and their interaction (P < 0.028).



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  		FIGURE 4 Specific activity of PCC (A) and PC (B) in thymocytes of mice fed a commercial standard diet (control group), biotin-sufficient, or biotin-deficient diets. Values are means ± SD, n = 3–9. At each time, the biotin-deficient group differed from the other groups, P < 0.001.

 
    Subpopulations of thymocytes. From wk 2 to 12 of experimentation, there were no significant differences among the groups in the proportions of double-negative (CD4–CD8–) thymocytes (DN), double-positive (CD4+CD8+) thymocytes (DP), and single-positive (SP) cells in the thymus (Fig. 5). However, from wk 16 to 20, the proportions of the 4 subpopulations in the thymus of biotin-deficient mice differed from those in the control and biotin-sufficient groups (Fig. 5A–D). The percentages of DN, DP, and CD4-SP and CD8-SP cells were affected by time, diet, and their interaction (P < 0.001). The percentage of CD3+ thymocytes in mice of the 3 experimental groups did not change during the study and did not differ among the 3 groups (Fig. 5E).



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  		FIGURE 5 Lymphocyte subpopulations in the thymus of mice fed a commercial standard diet (control group), biotin-sufficient, or biotin-deficient diets. Values are means ± SD, n = 3–9. (A) CD4–CD8– (DN) thymocytes; (B) CD4+CD8+ (DP) thymocytes; (C) CD4+ SP thymocytes; and (D) CD8+ SP thymocytes. (E) CD3{epsilon}+ thymocytes. The letter "a" indicates differences from the control and biotin-sufficient groups, P < 0.001.

 
Representative dot plots of thymocytes analyzed for the expression of CD4 and CD8 from 1 mouse of each experimental group after 16 wk of experimentation showed that the cells of biotin-deficient mouse had a diminished percentage of DP cells, and a corresponding increase in the percentage of DN and SP (CD4 and CD8) cells compared with the control and the biotin-sufficient mice (Fig. 6). These data suggest that chronic biotin deficiency induces an arrest of thymocyte maturation at the DN stage.



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  		FIGURE 6 Dot-plots of CD4 vs. CD8 expression in thymocytes from mice fed a commercial standard diet (control group), biotin-sufficient, or biotin-deficient diets. Representative thymic T cell subset profile of cells obtained from 1 mouse from each experimental group at wk 16 of experimentation. Total thymocytes were stained with anti CD4-FITC and anti CD8a-PE and analyzed by flow cytometry as described in the Materials and Methods. The percentage of each cell subpopulation is indicated within each quadrant.

 
To define more precisely the substage at which the maturation of thymocytes is arrested in biotin-deficient mice, we determined the expression of CD25 and CD44 in the DN thymocytes. Representative dot-plots of FSC vs. Fl-1 (FITC fluorescence) of thymocytes of 1 mouse from each experimental group at wk 20, illustrate how we selected the cell population negative for the expression of CD4 and CD8 (R1 in Fig. 7A). The gating and further analysis of the expression of CD25 and CD44 on DN cells, allowed us to quantify the DN-1 to DN-4 subgroups. The dot plots of CD25 and CD44 expression in the DN population of thymocytes from each of the 3 mice tested showed that 75% of the DN subpopulation in biotin-deficient thymocytes was CD44–CD25– (substage DN4). Also, it is worth noting that expression of CD25 is very low in DN cells from biotin-deficient mice (Fig. 7B).



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  		FIGURE 7 Analysis of DN thymocytes from mice fed the commercial standard diet (control group), biotin-sufficient and biotin-deficient diets. (A) Dot plots of CD4 and/or CD8 expression (FITC labeled, FL1) vs. forward scatter (size) of thymocytes from 1 mouse from each of the experimental groups at wk 20 of experimentation. R1 is the region of CD4 and CD8 double-negative cells. (B) Cytograms of cells in the region R1 (double negative cells) stained with antibodies anti-CD25-PE and anti-CD44-CyChrome. The number on each quadrant indicates the percentage of cells expressing the possible combinations of the expression of these cell surface markers.

 

   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
LITERATURE CITED
 
As previously described for humans (27–30), rats, and mice (11,12,14,15,31), during the first 4 wk of egg white feeding, a marginal asymptomatic biotin deficiency is produced; after a longer time, this develops into a frank and symptomatic biotin deficiency. Bearing this in mind, the lack of increase in the weight of the liver (Fig. 2C), and the marked decrease in the specific activity of PC and PCC in the thymus (Fig. 4), are early indicators of biotin deficiency in our experimental animal model. When the deficiency becomes chronic, we demonstrated 3 different effects that are present in biotin-deficient mice: 1) the overall growth of the mice is impaired, with reductions in the spleen, thymus, and liver weight, and in the nose-rump length; 2) the involution of the thymus is accelerated, as defined by loss of thymus weight, the decrease of cellularity, and the increase of the proportion of DN thymocytes; and 3) thymocyte maturation is arrested at the DN4 substage.

The conclusion that all the aforementioned alterations are caused by the deficiency of biotin is based on the observation that none of these effects were present in mice fed the biotin-sufficient diet. However, as was the case for other effects reported earlier by us (11), none of these effects had a time course that directly reflects the time course of the fall of the specific activity of the biotin-dependent carboxylases PC and PCC.

As indicators of the biotin status of the experimental mice, we used the specific activity of PC and PCC in the liver. Because PC activity in the liver is often the most dramatically reduced carboxylase in the biotin-deficient state, it is frequently used to confirm and estimate the extent of biotin deficiency (11–13). Lymphocyte PCC activity has also been used as an indicator of biotin status because it closely parallels the decrease in hepatic PCC activity (11,12). Moreover, the specific activity of these 2 enzymes is considered a better indicator of biotin status than the measurement of biotin metabolites in urine (28,30) or the concentration of biotin in serum (29,32).

Many studies reported that biotin deficiency produces weight loss, but it has not been determined whether it affects the length of the body or how it affects the weight of individual organs. In our mouse model, we observed that nose-rump length decreased from wk 8 of biotin deficiency (Fig. 1). It was reported that biotin deficiency in chickens and turkeys causes an abnormal maturation of the bones (33). By X-ray, we observed that the reduction in nose-rump length was not due to deformities in the spinal column (data not shown), suggesting that biotin deficiency in mice affects bone homeostasis. Additionally, our results show that biotin deficiency not only decreases the absolute weight of each of the studied organs (Fig. 2), but the deficiency affects differentially the ratio of the weight of the spleen, the thymus, and the liver, to the corporal weight of each mouse (data not shown). Thus, although free biotin and biotin-dependent carboxylases are ubiquitous, they could be subjected to different regulation mechanisms in different organs.

Growth hormone (GH) production by the pituitary is regulated by hormones produced in the hypothalamus. The physiologic actions of GH are pleiotropic and involve multiple organs and physiologic systems. Longitudinal bone growth and bone remodeling, skeletal muscle growth (fiber, strength), and liver growth, are some of the biological actions of GH (34). Our results suggest that biotin is essential for proper function of the GH neuroendocrine system.

T cell maturation in the thymus goes through distinct stages defined phenotypically by the expression of CD4 and CD8 coreceptors. On the basis of the expression of these 2 markers, thymocytes can be classified as double-negative (DN), double-positive (DP), or single-positive (SP) either CD4+CD8– or CD4–CD8+ cells (35). T-cell precursors that enter the thymus cortex are DN. As maturation proceeds, cells move toward the medulla and become DP thymocytes. Maturation from DN to DP goes through 4 sequential substages defined in terms of the expression of CD44 and CD25 molecules on the membrane; the 4 are as follows: CD44+CD25– (DN-1), CD44+CD25+ (DN-2), CD44–CD25+ (DN-3), and CD44–CD25– (DN-4). DP cells continue their maturation by reaching the medulla and becoming SP mature T cells; they then leave the thymus to travel through the bloodstream and reach secondary lymphoid organs (35). We showed that biotin deficiency not only results in a reduction in the number of thymocytes, but that there is also a change in the relative proportions of cells in the different maturation states, suggesting that a specific stage in the T-cell maturation process is especially sensitive to biotin deficiency.

From birth through adulthood, the lymphoid tissue of the thymus is progressively replaced by adipose tissue. This process is known as age-related thymus involution and is characterized by the following: 1) a progressive loss of cellularity; 2) a disruption of the normal architecture of the organ; 3) a decrease in the relative number of double-positive lymphocytes with a corresponding increase in the percentage of double-negative cells; 4) a progressive loss of cortical epithelial cells; and 5) a low efflux of mature T cells to the periphery (35–37).

Protein-energy malnutrition accelerates thymus involution (38,39). The accelerated thymus involution in biotin-deficient mice is not due to a deficiency of biotin inducing a state equivalent to protein-energy malnutrition because in the latter, the proportion of both CD4+ and CD8+ mature T lymphocytes decreases (40), whereas in biotin-deficient mice, both subpopulations in spleen (11) and thymus (present work) increase.

It was suggested that the adverse effects of biotin deficiency on the immune system are not likely to be specific to such deficiency (27). The best known function of biotin is its role as a cofactor of cellular carboxylases. Due to the central role of these enzymes in the metabolism of carbohydrates and amino acids, biotin deficiency might have an effect on physiologic phenomena secondary to its effect on cellular metabolism. However, the different times at which the decrease in enzymatic activity of PC and PCC vs. the alterations in the immune system become evident suggest that these effects are not a direct consequence of the low level of the carboxylases activity. Regulation of biotin metabolism (both free and protein-bound) is complex and is affected by several factors including the dietary biotin intake (31,41), differential effects at the transcription and expression levels of carboxylases (12,41–43), and conditions such as pregnancy, lactation, anticonvulsant therapy, and old age (27,29). More biochemical, metabolic, and molecular studies are required to discern whether the observed effects of biotin deficiency on the spleen (11) and thymus (present work) are an indirect consequence of the metabolic alterations derived from a low level of activity of carboxylases or are caused directly by the lack of another function of biotin.

In addition to its role as cofactor of carboxylases, biotin modulates gene transcription (44); it is involved in the regulation of the expression of hepatic glucokinase (45), pancreatic glucokinase (46), and PC, PCC, and holocarboxylase synthetase in rats (47). Moreover, biotin was shown to induce expression of holocarboxylase synthetase, ACC, and PCC in humans (48). In addition, there are other observations that cannot be related directly to the role of biotin as a cofactor of carboxylases, such as the following: 1) the presence of substantial amounts of biotin in the nucleus of certain tumor cells (49); 2) biotin-induced histone modifications (50); 3) enzymatic biotinylation of histones (51); 4) the effect of biotin on the expression of interleukin-2 and of the {gamma} chain of its receptor (52); and 5) the modulation of transcription factors Sp1 and Sp3 (53). All of the above indicate that in addition to its role as cofactor of enzymes, biotin can have effects on gene expression. The number and nature of genes whose expression might be modulated by biotin are only beginning to be explored. Analyses of the effect of biotin on genes involved in control of T-cell maturation by hormones, chemokines, and cytokines, might help in understanding the deleterious effect of biotin deficiency on the immune phenomena.


   ACKNOWLEDGMENTS
 
The authors are indebted to Rafael Saavedra for his critical reading of the manuscript. We also thank Claudia Garay for her expert technical assistance, Georgina Díaz for her expert advice and help in the care of the animals, and René Rosiles and Rosa Elena Méndez for their help in the X-ray exam of mice. Blanca A. Rosales-Báez corrected the English version of the manuscript.


   FOOTNOTES
 
1 Supported by a research grant (IN208399) from the Dirección General de Asuntos del Personal Académico, UNAM, México. Back

3 Abbreviations used: ACC, acetyl-CoA carboxylase; CD3{epsilon}, {epsilon} chain of the T-cell receptor-associated CD3 complex; CD4, L3T4 antigen of differentiation expressed on T lymphocytes; CD8a, Ly-2 antigen of differentiation expressed on the surface of T lymphocytes; CD25, {alpha} chain of the interleukin 2 receptor; CD44, glycoprotein expressed on hematopoietic and non-hematopoietic cells; CyChrome, peridinin chlorophyll protein-cyanin 5.5; DN, double negative (CD4–CD8–) thymocytes; DP, double-positive (CD4+CD8+) thymocytes; FITC, fluorescein isothiocyanate; mAbs, monoclonal antibodies; MCC, methylcrotonyl CoA carboxylase; PC, pyruvate carboxylase; PCC, propionyl CoA carboxylase; PE, phycoerytrin; PerCP, peridinin chlorophyll protein; SP, single-positive T lymphocyte, either CD4+CD8– or CD4–CD8+. Back

Manuscript received 21 January 2004. Initial review completed 4 February 2004. Revision accepted 23 May 2004.


   LITERATURE CITED
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 LITERATURE CITED
 

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