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© 2008 American Society for Nutrition J. Nutr. 138:680-684, April 2008

Biochemical, Molecular, and Genetic Mechanisms

Biotin Deficiency Affects the Proliferation of Human Embryonic Palatal Mesenchymal Cells in Culture1,2

Ryusuke Takechi3,5, Ayumi Taniguchi3,6, Shuhei Ebara3, Toru Fukui4 and Toshiaki Watanabe3,*

3 Department of Dietary Environment Analysis, School of Human Science and Environment, Himeji Institute of Technology, University of Hyogo, Himeji, Japan and 4 Clinical Laboratory, Byotai Seiri Laboratory, Tokyo, Japan

* To whom correspondence should be addressed. E-mail: watanabe{at}shse.u-hyogo.ac.jp.


   ABSTRACT
TOP
 ABSTRACT
Introduction
 Materials and Methods
 Results
 Discussion
 LITERATURE CITED
 
It has recently been demonstrated that pregnancy in women may cause mild biotin deficiency without any clinical signs. However, the teratogenicity of biotin deficiency in humans has not been well investigated. On the other hand, our previous studies have shown that maternal biotin deficiency induces many kinds of malformations, such as cleft palate, micrognathia, and micromelia, in all animal fetuses. However the mechanism for cleft palate induction under biotin-deficient conditions is unknown. Therefore, to investigate the possible mechanisms for cleft palate induction in embryos, we investigated the effects of biotin deficiency on human embryonic palatal mesenchymal (HEPM) cells in culture in this study. HEPM cells were cultured in biotin-deficient and biotin-physiological (control) media for 5 wk. The proliferative availabilities of HEPM cells in the biotin-deficient state were significantly lower after wk 2 of culture (41.3% of the control). Biotin concentrations in biotin-deficient cells were drastically lower after wk 1 of culture, whereas those in the control cells remained at almost the same level. Biotinidase activities were also lower in biotin-deficient cells. Holocarboxylases in biotin-deficient cells were fewer after the first week of culture and were almost undetectable after wk 2. The amount of biotinylated histones in the nuclei of biotin-deficient cells was lower than in the control cells. This suppressed proliferation of mesenchymal cells may delay or inhibit the growth of palatal processes in embryos and thus it may partially contribute to the mechanisms for cleft palate induction.


   Introduction
TOP
 ABSTRACT
 Introduction
Materials and Methods
 Results
 Discussion
 LITERATURE CITED
 
Biotin is a kind of water-soluble vitamin that is classified as a B vitamin. Biotin was initially discovered as a factor that prevented dermatitis, hair loss, and neurological abnormalities, which are produced in rats by a diet containing egg white. Biotin acts as a prosthetic cofactor for enzymes that catalyze carboxylation. In mammals, biotin functions as a cofactor for biotin-dependent carboxylases: acetyl-CoA carboxylase {alpha} (ACC{alpha}),7 ACCβ, pyruvate carboxylase (PC), propionyl-CoA carboxylase (PCC), and methylcrotonyl-CoA carboxylase (MCC).

Two isoforms of ACC have been identified in mammals: cytosolic ACC{alpha} and mitochondrial ACCβ. ACC{alpha} participates in fatty acid synthesis in the cytoplasm by providing the substrate malonyl-CoA. In contrast, ACCβ participates in the regulation of fatty acid oxidation in the mitochondria. PC catalyzes the synthesis of oxaloacetate from pyruvate. This step in the citric cycle is also the rate-limiting step in gluconeogenesis and PC serves as the key regulatory enzyme for this process. PCC catalyzes the conversion of propionyl CoA into methylmalonyl CoA in the catabolic pathway for the branched-chain amino acids, odd-chain length fatty acids, and cholesterol. MCC is essential for the catabolism of leucine (1). 3-Methylcrotonyl CoA is converted into 3-methylglutaconyl CoA, which is hydrated to form 3-hydroxy-3-methylglutaryl CoA (2).

Histones are basic proteins associated with DNA in nucleosomes, which consist of core histones (H2A, H2B, H3, and H4) and linker histones (H1). Histones are subjected to post-translational modifications, such as methylation, acetylation, phosphorylation, ubiquitination, and biotinylation. Hymes et al. (3) have proposed a reaction mechanism by which the enzyme biotinidase mediates the covalent binding of biotin to histones. Recent studies have shown new roles for biotin in cell proliferation. Stanley et al. (4) provided evidence that the biotinylation of histones increases in response to cell proliferation in human lymphocytes. Histone biotinylation increases early in the cell cycle (G1 phase) and remains elevated during later phases (S, G2, and M phases). In addition, enriched biotinylated histones were observed in the transcriptionally silent chromatin of chicken erythrocytes (5).

We first found increased fetal malformations and mortality in biotin-deficient pregnant rodents (6–9). In these studies, biotin deficiency was induced by feeding raw egg white to the animals. The incidence of fetal malformations was extremely high in mice. The defects consisted mainly of cleft palate, micrognathia, and micromelia. We have also shown differences in teratogenic susceptibility among rodent species. No malformations were observed in biotin-deficient rats. This may be caused by differences in biotin transport from the mother to the fetus (8). The pregnant dams showed no signs of biotin deficiency and gained normal amounts of weight, the same as the normal pregnant dams from our previous studies. It has been demonstrated that the fetal to maternal biotin ratio increases during the 2nd trimester of normal human pregnancies (10). These findings suggest substantial utilization of biotin by the fetus and high fetal sensitivity to biotin deficiency.

Although a high incidence of cleft palate induction under biotin-deficient conditions has been reported in many studies, its mechanisms are unknown. Formation of the secondary palate occurs early in embryonic development and involves a complex series of steps, including both the growth and differentiation of epithelial and mesenchymal cells (11,12). Human embryonic palatal mesenchymal (HEPM) cells were originally derived from the mesenchyme of the secondary palatal shelves from a human embryonic abortus aborted after 56 d of gestation. HEPM cells have previously been used in studies to determine the mechanisms for cleft palate induction by various teratogen, to determine the role that various growth factors play in the regulation of embryonic development, and to assess the teratogenic potential of environmental agents (13). Therefore, using HEPM cells, we undertook this study to determine how biotin deficiency affects the formation of the palate in embryos and the specific mechanisms of cleft palate induction in culture.


   Materials and Methods
TOP
 ABSTRACT
 Introduction
 Materials and Methods
Results
 Discussion
 LITERATURE CITED
 
    Cell culture. We purchased HEPM cells from the American Type Culture Collection (ATCC). The cells were cultured at 37°C in an incubator with 5% CO2. The culture medium consisted of DMEM (Gibco Japan) supplemented with 10% (v:v) fetal bovine serum (FBS) and 2% (v:v) penicillin streptomycin glutamine. The medium was changed every 3 d, or when its color turned from pink-red to yellow-orange, and the cells were split every week.

To obtain the control and biotin-deficient media, 1 Affinity Pak Immobilized Avidin Column (Pierce Biotechnology) containing 1 mL of immobilized avidin gel was used to strip the biotin from 50 mL of the FBS. The biotin-stripped FBS was stored at –20°C until the next use. For biotin-deficient and biotin-physiological (control) media, d-biotin (dissolved in ethanol and stored at –20°C) was added to make the final concentrations of biotin in the media 1.5 nmol/L and 10 nmol/L, respectively. The concentration in the control media was the same as the serum biotin concentration in healthy adults measured by microbiological assay. The concentration in the deficient media was >2 SD below normal biotin concentration in serum. Both biotin-deficient and control media were stored at 4°C and freshly made at least once per month.

    Proliferation of HEPM cells. The cells were subcultured in a control medium for 24 h in a 75-cm2 culture flask. Subsequently, the cells were split with 3 mL of trypsin and seeded into 30-mm diameter multi-cell culture dishes at a density of 4.0 x 104 cells per dish. The cells were cultured in triplicate in both biotin-deficient and control media and each medium was changed every 3 d. Each cell under both conditions was split on d 7, washed twice with 1 mL of ice-cold PBS, and split with 200 µL of trypsin. At this time, the proliferated cells were counted using a hemocytometer and reseeded at a density of 4.0 x 104 cells per dish. The number of proliferated cells was measured for 5 wk and the data thus collected was utilized as an indicator of cell proliferating availability. Also, the remaining cells were stored to measure their biotin concentration and biotinydase activity. Each week in the time axes represents a point when the cells had proliferated for 1 wk (Figs. 1 and 2).


Figure 1
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  		FIGURE 1  Proliferation of HEPM cells cultured in control or biotin-deficient media for 5 wk. The number of cells proliferating from 4 x 104 was counted. Values are means ± SD, n = 3. Means without a common letter differ, P < 0.05.

 

Figure 2
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  		FIGURE 2  Biotin concentrations (A) and biotinidase activities (B) in HEPM cells cultured in control or biotin-deficient media for 5 wk. Values are means of triplicate samples, n = 1. Intracellular biotin was extracted in PBS and measured by bioassay using Lactobacillus plantarum ATCC 8014. Biotinidase activities were measured by colorimetric assay. Biotinidase activities are expressed as µmol p-aminobenzoate liberated·min–1·L–1 sample.

 
    Measurement of biotin concentration and biotinidase activity. The HEPM cells collected and stored were thawed on ice. The cells were pelleted at 2500 x g for 5 min at 4°C and resuspended in 500 µL of PBS. Intracellular biotin was extracted using a sonicator. The sonicated sample was centrifuged at 2500 x g; 10 min at 4°C and supernatant was collected. The pellet was resuspended in 200 µL of PBS and the sonication and centrifugation were repeated.

We measured biotin concentration in the samples (cells and media) by bioassay using Lactobacillus plantarum ATCC 8014 (14) and determined biotinidase activity using the colorimetric method by measuring the liberation of p-aminobenzoate from N-biotinyl-p-aminobenzoate (15).

    Detection of biotin-dependent carboxylases. The stored HEPM cells were thawed and sonicated in 300 µL of solubilization buffer [PBS supplemented with 1% (v:v) Triton-X 100 and 0.02% (v:v) protease inhibitor cocktail] on ice. The solubilized cells were centrifuged at 800 x g; 5 min at 4°C and the supernatant was stored at –40°C until next use. Each supernatant was adjusted to the same concentration of protein and subjected to SDS-PAGE with NuPAGE 3–8% precast gradient gels (Invitrogen Japan). After being transferred to a polyvinylidene difluoride membrane, biotin-dependent carboxylases were probed with streptavidin peroxidase (Sigma-Aldrich Japan) and detected using the enhanced chemiluminescence (ECL, Amersham Biosciences) technique.

As for the measurement of the total amount of carboxylases (total number of apo- and holo-carboxylases), the apocarboxylases in the samples were biotinylated in vitro by a technique modified from Rodriguez-Melendez et al. (16) and Desjardins and Dakshinamurti (17). Biotinylated carboxylases were subjected to SDS-PAGE as described above.

Protein concentration was measured using a modified Lowry's method using a BCA protein assay kit (Pierce Biotechnology).

    Detection of biotinylated histones in cell nuclei. The stored HEPM cells were thawed and sonicated in 300 µL of solubilization buffer [PBS supplemented with 1% (v:v) Triton-X 100 and 0.02% (v:v) protease inhibitor cocktail] on ice. The solubilized cells were centrifuged at 100 x g; 10 min at 4°C. The supernatant was collected and centrifugation was repeated. Cell nuclei pellets were combined and resuspended in 100 µL of 0.5 mol/L HCl containing 2% (v:v) 2-mercaptoethanol and 2 mmol/L phenylmethanesulfonyl fluoride and then incubated at 4°C overnight. The solution was centrifuged at 10,000 x g; 4 min at 4°C and the supernatant was collected. The pellet was resuspended in 50 µL of the HCl mixture for another 6 h at 4°C and the solution was centrifuged at 10,000 x g; 4 min at 4°C. The supernatant was combined with the first supernatant. The combined supernatant was adjusted to approximately pH 7 using 4 mol/L KOH and stored at –40°C until next use.

The extracted histones were subjected to SDS-PAGE. Biotinylated histones were detected by use of a streptavidin peroxidase probe as described in the section entitled "Detection of Biotin-Dependent Carboxylases."

The amounts of all types of histones were detected as follows: the electrophoresed gel was transferred into a plastic container and gently washed with distilled water several times. Then, the gel was heated in water using a microwave for ~1 min to dissolve out the SDS in the gel. This process was repeated 3 times. The histones were detected using a Coomassie stain procedure.

    Statistical analysis. All values are means ± SD. Cell proliferation was compared between the control and biotin-deficient cells for 5 wk by 2-way ANOVA for repeated measures and post hoc analyses were conducted using the Tukey-Kramer test. All statistical tests were 2-sided at the 5 and 1% levels and performed using SPSS software (version 12.0; SAS Institute). Graphs were drawn using a GraphPad Prizm (version 3.6; Synergy Software).


   Results
TOP
 ABSTRACT
 Introduction
 Materials and Methods
 Results
Discussion
 LITERATURE CITED
 
    Cell proliferative availability. The proliferation rates of HEPM cells cultured in biotin-control media did not change during the culture period. However, the proliferation rates of HEPM cells cultured in biotin-deficient media were significantly suppressed from wk 2 of culture (Fig. 1). At wk 5 of culture, a reduction in the number of proliferated cells was identified clearly using a microscope.

    Intracellular biotin concentration and biotinidase activity. In control cells cultured in the media containing a physiological concentration of biotin, the intracellular biotin concentrations, ranging from 457 to 917 fmol/cell, did not change much throughout the 5 wk of the culture (Fig. 2A). However, the biotin concentrations in the biotin-deficient cells were drastically lower than in the control cells at any given point in time, which did not change throughout the culture period, remaining in the range from 8.1 to 28.3 fmol/cell.

Biotinidase activity in the control cells remained at almost the same level throughout the experiment. However, the biotinidase activity in biotin-deficient cells was lower after wk 2 of culture and reached an almost undetectable level after wk 4 of culture (Fig. 2B).

    Biotin-dependent carboxylases. The levels of electrophoresis band areas and densities of ACC, PC, MCC, and PCC were suppressed in biotin-deficient cells from wk 1 of culture and reached an undetectable level from wk 2 of culture, whereas the levels in the control cells did not change much throughout the culture period (Fig. 3). In the control cells, the total mass of PC and MCC did not change much throughout the culture period (Fig. 4). However, the total mass of PC and MCC in biotin-deficient cells was lower from wk 3 of culture and reached an undetectable level from wk 4 of culture. Additionally, the expression of PC and MCC in biotin-deficient cells at wk 1 and 2 of culture was the same as that in the control cells.


Figure 3
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  		FIGURE 3  Biotinylation of biotin-dependent carboxylases. Biotin-dependent holocarboxylases of ACC, PC, MCC, and PCC in the control or deficient cells from wk 1 to 5 of culture were separated on SDS-PAGE and probed by western blotting with streptavidin peroxidase.

 

Figure 4
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  		FIGURE 4  Total mass of biotin-dependent carboxylases in control or biotin-deficient cells for 5 wk separated and probed by western blotting after the in vitro biotinylation of carboxylases. The protein bands of ACC and PCC were not detectable by western blotting using streptavidin peroxidase in our method.

 
    Biotinylation of histones. The total expression of histones was detected using a Coomassie stain (Fig. 5A). The levels of each type of histone, H1, H2A, H2B, H3, and H4, in biotin-deficient cells were almost the same as those in the control cells. Biotinylated histones were detected by using streptavidin peroxidase after separation by SDS-PAGE (Fig. 5B). The protein bands for H1, H3, and H4 were all low in the control cells. However, no protein bands for biotinylated histones were detectable in biotin-deficient cells.


Figure 5
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  		FIGURE 5  Electrophoresis images of nuclear histones. (A) Histones H1, H2A, H2B, H3, and H4 were extracted in HCl and separated on SDS-PAGE. The total expression of histones was detected using a Coomassie Blue gel stain. (B) The biotinylation of histones was measured by western blotting using streptavidin peroxidase and labeled anti-biotin. The protein bands of H1 were almost undetectable by western blotting in our method. Lane 1 shows the control cells and lane 2 shows the biotin-deficient cells at wk 5 of culture.

 

   Discussion
TOP
 ABSTRACT
 Introduction
 Materials and Methods
 Results
 Discussion
LITERATURE CITED
 
In this study, HEPM cells were cultured in control and biotin-deficient media for 5 wk and the cell proliferation rates were measured every week. The proliferation of biotin-deficient cells was significantly lower after wk 2 of culture than that of the control cells. Crisp et al. (18) have shown decreased cell proliferation rates of human choriocarcinoma JAr cells under biotin-deficient cell culture conditions and this result is consistent with our results. In contrast, the cell proliferation rates of small cell lung cancer NCI-H69 cells (19) and Jurkat cells (20) did not decrease under biotin-deficient culture conditions. This suggests that the susceptibilities differ among the cell lines and that cell proliferative availability may be specifically affected by biotin deficiency.

On the other hand, some studies have revealed that biotin is associated positively with cell proliferation. Zempleni and Mock (21,22) have shown that mitogen-stimulated proliferating peripheral blood mononuclear cells (PBMC) and human lymphocytes accumulated biotin significantly faster than unstimulated controls. For example, when proliferation was stimulated by incubation with pokeweed mitogen for 3 d, biotin uptake increased in a dose-dependent fashion from 481 to 722% of the control value. Other evidence has been provided by a basic study reporting the significant stimulation of BHK cell growth in culture by free biotin, which is not protein bound and is thus more bioavailable (23). This study has also shown that free biotin was found in relatively large amounts in fetal sera and only a very small percentage of free biotin was contained in adult sera and was characterized by lower growth-stimulating activity. In addition, Mantagos et al. (10) reported that human fetal plasma contained more biotin [784 ± 327 ng/L (3.21 ± 1.34 nmol/L)] than maternal plasma [131 ± 102 ng/L (0.54 ± 0.42 nmol/L)]. These studies suggest that biotin utilization is significantly greater in the fetus, which may be consistent with our previous studies. We showed a high incidence of fetal malformations caused by biotin deficiency, whereas the pregnant dams showed no specific signs of biotin deficiency (6,8,24–27).

The absence of certain growth factors causes cells to enter a specialized resting state termed G0 (28). Nutrient deficiencies often arrest cells at the G0 phase and the cells neither enter the cell cycle nor divide. Dakshinamurti et al. (29,30) have shown that human cervical carcinoma cells halt in the G0 phase when the cells are incubated in biotin-free media. Studies on PBMC provided evidence that the biotin uptake rates of the different cell cycle phases were significantly different and the peak biotin uptake was observed in the G1/S phase (21). However, the mechanisms for biotin usage in cell proliferation and/or differentiation are uncertain.

Possible causes of the decreased proliferation of HEPM cells were also investigated in this study. The biotinylation of carboxylases is the best known physiological function of biotin. These biotin-dependent carboxylases play essential roles in cellular biosynthetic pathways. The activities of biotin-dependent carboxylases increase in mitogen-stimulated PBMC cells (22); hence, the blockage of these metabolic pathways by inactivated carboxylases may indirectly affect cellular proliferation. In HEPM cells, the amounts of biotinylated biotin-dependent carboxylases were drastically lower in biotin-deficient cells after wk 1 of culture than in control cells. Evidence exists that the biotinylation of biotin-dependent carboxylases in rats is associated with dietary biotin intake (31), suggesting that a reduction in the biotinylation of carboxylases in HEPM cells might be regulated by limited intracellular access to biotin. Additionally, in Jurkat cells and JAr cells, the biotinylation of carboxylases decreased when the cells were cultured in biotin-deficient media (18,20). However, it was 3–4 wk before carboxylase biotinylation was suppressed, whereas the process required only 1 wk in HEPM cells. This suggests that the biotinylation of carboxylases in HEPM cells is more easily affected by biotin deficiency.

The study on CD-1 mice showed that the carboxylases in fetuses were much lower than in dams, suggesting that the fetuses are more sensitive to biotin deficiency. Biotin-dependent carboxylase activity in the central nervous system cells, JAr cells, NCI-H69 cells, and Jurkat cells is significantly lower under biotin-deficient cell culture conditions (18–20,32). In these studies, carboxylase activity decreased within a relatively shorter period, a few days, than did the amount of biotinylated carboxylases, which required more than a few weeks to decline. Previous studies have shown that cell proliferative availabilities were kept normal for several weeks or more under biotin-deficient conditions, even after carboxylase activity decreased at an early stage (18,20).

The mechanism for the maintenance of cell proliferative availability with decreased carboxylase activity is unknown. Although the activities of carboxylases were not measured in this study, the amount of biotinylated carboxylases and the proliferation of HEPM cells showed a similar decrease and consistent results were observed with JAr cells. In contrast, cell proliferation in PBMC and Jurkat cells was not affected by biotin deficiency, whereas carboxylase biotinylation was significantly downregulated. Thus, the associations among cell proliferative availabilities, carboxylase biotinylation, and carboxylase activity are unclear. Studies in mice (33) and Jurkat cells (20) have provided evidence that the gene encoding the mRNA of the biotin-dependent carboxylases was not affected by biotin deficiency, although the activity and biotinylation of the carboxylases were decreased. These findings suggest that the biotin-dependent carboxylases may exist in tissues under biotin-deficient conditions even though the carboxylase biotinylation decreases.

To investigate the total mass of biotin-dependent carboxylases in HEPM cells, we measured the expression of carboxylases after the in vitro biotinylation of carboxylases in this study. As a result, a decrease in the total mass of the carboxylases was observed, suggesting that the expression of the mRNA encoding biotin-dependent carboxylases in HEPM might be regulated by biotin deficiency. This is obviously a different observation from those of previous studies and may suggest the specific effects of biotin on HEPM cells.

Histones are involved in the replication and repair of DNA and hence are directly related to cell proliferation and differentiation. The biotinylation of histones was significantly increased in proliferating PBMC and the incorporation of biotin into histones was greater at all phases of the cell cycle (G1, S, G2, and M) than in quiescent cells (4,28). In this study, the biotinylation of histones decreased in biotin-deficient HEPM cells and the results were consistent with the study on JAr cells (18). In both studies, decreased histone biotinylation was observed along with the decreased cell proliferation. In contrast, the biotinylation of histones is not decreased by biotin deficiency in Jurkat cells, the proliferation of which does not decrease under the biotin-deficient conditions (20). Similarly, the cell proliferation of NCI-H69 is not affected by biotin deficiency; only the biotinylation of histones H2 and H4 decreases, whereas that of histones H1 and H3 remains at the same levels (19).

Previous studies suggested that the biotinylation of histones might play a role in cell proliferation, gene silencing, and the repair of damaged DNA or apoptosis (4). Changes in the biotinylation of histones might affect processes such as gene expression within confined regions of chromatin without having a substantial effect on the overall biotinylation of histones in the chromatin as a whole. Because cell proliferation requires a substantial increase in both the replication and transcription of DNA, an increased requirement of biotin uptake by proliferating cells to biotinylate histones is a plausible mechanism by which this nutrient might exert at least some of its effects on cell proliferation.

In conclusion, the proliferative availability of HEPM cells in this study was lower in biotin-deficient cells than in control cells. The biotinylation of biotin-dependent carboxylases and histones was also drastically lower in biotin-deficient cells than in control cells, which resulted from limited access to intracellular biotin. Biotin deficiency may cause cleft palate via delayed or arrested growth of the embryonic palate by the suppressed cell proliferation of HEPM cells. Therefore, this study has revealed a possible mechanism for cleft palate induction by biotin deficiency.


   FOOTNOTES
 
1 Supported by a grant from the University of Hyogo. Back

2 Author disclosures: R. Takechi, A. Taniguchi, S. Ebara, T. Fukui, and T. Watanabe, no conflicts of interest. Back

5 Present address: School of Public Health, Curtin University of Technology, Perth, Australia. Back

6 Present address: Institute for Environmental and Gender Specific Medicine, Graduate School of Medicine, Juntendo University School of Medicine, Urayasu, Japan. Back

7 Abbreviations used: ACC, acetyl-CoA carboxylase; ATCC, American Type Culture Collection; FBS, fetal bovine serum; HEPM, human embryonic palatal mesenchymal; MCC, methylcrotonyl-CoA carboxylase; PBMC, peripheral blood mononuclear cells; PC, pyruvate carboxylase; PCC, propionyl-CoA carboxylase. Back

Manuscript received 4 November 2007. Initial review completed 28 November 2007. Revision accepted 9 January 2008.


   LITERATURE CITED
TOP
 ABSTRACT
 Introduction
 Materials and Methods
 Results
 Discussion
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