Expression and functional contribution of hTHTR-2 in thiamin absorption in human intestine

Hamid M. Said,1,2 Krishnaswamy Balamurugan,1,2 Veedamali S. Subramanian,1,2 and Jonathan S. Marchant3

1Veterans Affairs Medical Center, Long Beach 90822; 2University of California College of Medicine, Irvine, California 92697; and 3University of Minnesota Medical School, Minneapolis, Minnesota 55455

Submitted 21 August 2003 ; accepted in final form 22 October 2003


    ABSTRACT
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The aim of this study was to investigate expression and relative contribution of human thiamin transporter (hTHTR)-2 toward overall carrier-mediated thiamin uptake by human intestinal epithelial cells. Northern blot analysis showed that the message of the hTHTR-2 is expressed along the native human gastrointestinal tract with highest expression being in the proximal part of small intestine. hTHTR-2 protein was found, by Western blot analysis, to be expressed at the brush-border membrane (BBM), but not at the basolateral membrane, of native human enterocytes. This pattern of expression was confirmed in studies using a fusion protein of hTHTR-2 with the enhanced green fluorescent protein (hTHTR2-EGFP) expressed in living Caco-2 cells grown on filter. Pretreating Caco-2 cells (which also express the hTHTR-2 at RNA and protein levels) with hTHTR-2 gene-specific small interfering RNA (siRNA) led to a significant (P < 0.01) and specific inhibition (48%) in carrier-mediated thiamin uptake. Similarly, pretreating Caco-2 cells with siRNA that specifically target hTHTR-1 (which is expressed in Caco-2 cells) also significantly (P < 0.01) and specifically inhibited (by 56%) carrier-mediated thiamin uptake. When Caco-2 cells were pretreated with siRNAs against both hTHTR-2 and hTHTR-1 genes, an almost complete inhibition in carrier-mediated thiamin uptake was observed. These results show that the message of hTHTR-2 is expressed along the human gastrointestinal tract and that expression of its protein in intestinal epithelia is mainly localized to the apical BBM domain. In addition, results show that this transporter plays a significant role in carrier-mediated thiamin uptake in human intestine.

thiamin transport; polarized expression; small interfering RNA; Caco-2 cells


VITAMIN B1 (THIAMIN) is a member of the water-soluble vitamin family of micronutrients. It plays an essential role in normal cellular functions, growth, and development. In its coenzyme form, i.e., thiamin pyrophosphate, thiamin plays a vital role in metabolism and energy production reactions that include the decarboxylation of pyruvic acid and {alpha}-ketoglutamic acids and the utilization of pentose in the hexose monophosphate shunt (2). Thiamin deficiency in humans leads to a variety of clinical abnormalities including neurological and cardiovascular disorders (2, 27, 28). On the other hand, optimization of thiamin levels appear to have the potential of preventing diabetic retinopathy and blocking tissue damage caused by hyperglycemia of diabetes (10). Thiamin deficiency and suboptimal levels represent significant nutritional problems and occur under a variety of conditions including alcoholism (8, 14, 26) and diabetes mellitus (24). Deficiency in thiamin accumulation also occurs in the inherited condition of thiamin-responsive megaloblastic anemia (TRMA) (19, 21). A mutational defect in the human thiamin transporter (hTHTR)-1 is believed to be the cause of TRMA (3, 6, 9, 12).

Humans and other mammals cannot synthesize thiamin and thus must obtain the vitamin from exogenous sources via intestinal absorption. The intestine, therefore, plays a critical role in maintaining and regulating normal thiamin body homeostasis. Absorption of thiamin from the intestinal lumen involves a specialized, pH-dependent, Na+-independent, carrier-mediated mechanism (4, 5, 13, 18, 22, 23). Insight into the molecular identity of the transport mechanism of thiamin in the human intestine has begun to emerge recently after the cloning of hTHTR-1 and hTHTR-2 (the products of the SLC19A2 and SLC19A3 genes, respectively) from a number of human tissues (3, 6, 7, 9, 12, 16). cDNA of the hTHTR-1 codes for a protein of 497 amino acids that has 12 putative transmembrane domains and shares 40% identity (at the amino acid level) with the human reduced folate carrier (hRFC; the product of the human SLC19A1 gene) (3, 6, 9, 12). However, no functional overlap was found between these two transporters, i.e., the hTHTR-1 does not transport folate and the hRFC does not transport thiamin (6). hTHTR-1 is expressed in different tissues of the human gastrointestinal tract, including the small and large intestine, as well as in the human-derived intestinal epithelial models, Caco-2 cells (17). cDNA of the hTHTR-2 codes for a protein of 496 amino acids that again has 12 putative transmembrane domains (7, 16). At the amino acid level, hTHTR-2 shares 48 and 39% identity with the hTHTR-1 and the hRFC, respectively, but it does not transport folate either (7, 16). Little is currently known about the expression of the hTHTR-2 along the native human gastrointestinal tract and at the opposing membranes of the polarized human intestinal epithelial cells. Also not known is the relative contribution of this transporter toward overall carrier-mediated thiamin uptake by human intestinal epithelial cells. Our aim in this study was to address these issues by using preparations from native human gastrointestinal tract and the human-derived intestinal epithelial Caco-2 cells [a suitable model for studying thiamin uptake in the human intestine (22)].


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Chemicals and reagents. Custom-made [3H]thiamin (specific activity 555 GBq/mmol; radiochemical purity >98%) was purchased from American Radiolabeled Chemicals (St. Louis, MO). The enhanced green fluorescent protein vector (EGFP-N3) was from BD Biosciences (Palo Alto, CA). TRIzol reagent and Lipofectamine 2000 were purchased from Life Technologies (Rockville, MD). DNA oligonucleotide primers were ordered from Sigma Genosys (Woodlands, TX). Routine biochemicals, enzymes, and cell culture reagents were all of molecular biology quality and purchased from either Fisher Scientific (Tustin, CA) or Sigma (St. Louis, MO).

Cell culturing and uptake investigation. Caco-2 cells were purchased from American Type Culture Collection (Rockville, MD) and were used between passages 17 and 27. Cells were grown either on solid support as described by us previously (22) or on filter (see Generation of hTHTR2-EGFP fusion protein, cell transfection, and confocal imaging) and were maintained in DMEM supplemented with 10% fetal bovine serum, glutamine (0.29 g/l), sodium bicarbonate (2.2 g/l), penicillin (100,000 U/l), and streptomycin (10 mg/l). Uptake studies were performed by using confluent monolayers (3–5 days after confluence) of Caco-2 cells. Uptake was measured at 37°C in Krebs-Ringer buffer (in mM: 133 NaCl, 4.93 KCl, 1.23 MgSO4, 0.85 CaCl2, 5 glucose, 5 glutamine, 10 HEPES, and 10 MES, pH 7.4). Labeled and unlabeled thiamin were added to the incubation medium at onset of incubation, and uptake was examined during the initial linear period (22). Reaction was then terminated by the addition of 2 ml of ice-cold buffer followed by immediate aspiration. Cells were then rinsed twice with ice-cold buffer, digested with 1 ml of 1 N NaOH, neutralized with HCl, and then counted for radioactivity. Protein content of cell digests was measured in parallel wells by using a Bio-Rad kit (Bio-Rad, Hercules, CA).

In studies examining the effect of pretreatment of Caco-2 cells with siRNA, custom-made hTHTR-1 and hTHTR-2 gene-specific siRNAs (double-stranded RNAs of 21 nucleotides; for hTHTR-1 the siRNA was 5'-aagttactgtcgaagtgccacdTdT-3'; for hTHTR-2 the siRNA was 5'-aaggagtgaagaccatgcaggdTdT-3') chemically synthesized by a commercial vendor (Qiagen-Xeragon, Germantown, MD) were used. Both strands of the siRNAs were modified by the addition of dTdT overhang at their 3' ends to increase stability. Transient transfection of subconfluent (~70%) Caco-2 cells with the specific siRNA (0.2 µg/well) were performed, and control cells were transfected with scrambled siRNA (5'-aacgcgcccagagcgcagctcdTdT-3') as recently described by us (1). Cells were maintained until 3 to 5 days after confluence (i.e., 5 to 7 days after pretreatment with the siRNA) and then used in the specific experiments.

Northern blot analysis, semiquantitative PCR, and Western blot analysis. Northern blot analysis was performed as described by us previously (23). Briefly, the full-length open reading frame of the hTHTR-2 cDNA cloned in our laboratory from Caco-2 cells was labeled with [32P]dCTP and used as a probe on a commercially available blot that contains poly(A)+ RNA (2 µg) from different regions of the human digestive tract (Clonetech, Palo Alto, CA). The probe was simultaneously used on a blot containing an equivalent amount of poly(A)+ RNA (2 µg) isolated from Caco-2 cells. The blots were stripped and reprobed with a labeled human {beta}-actin. The message of the hTHTR-2 was normalized relative to the {beta}-actin signal by using the UnscaniT gel software (Silk Scientific, Salt Lake City, UT).

Semiquantitative PCR to quantitate levels of hTHTR-1 and hTHTR-2 RNA was performed by using total RNA isolated from control and siRNA-pretreated Caco-2 cells. For hTHTR-1, the primers were forward, 5'-AGCCAGACCGTCTCCTTGTA-3' and reverse, 5'-TAGAGAGGGCCCACCACAC-3'. For hTHTR-2, the primers were: forward, 5'-TTCCTGGATTTACCCCACTG-3' and reverse, 5'-GTATGTCCAAACGGGGAAGA-3'. For {beta}-actin, the primers were forward, 5'-CATCCTGCGTCTGGACCT-3' and reverse, 5'-TAATGTCACGCACGATTTCC-3'. For hTHTR-1, the PCR products were analyzed between cycles 22 and 28 (linear range). For hTHTR-2, the PCR products were analyzed between cycles 26 and 32 (linear range). The level of expression was normalized relative to human {beta}-actin.

Western blot analysis was performed by using apical and basolateral membrane proteins isolated from the jejunum of organ donors by an established procedure (4, 5). These samples were provided to us by our collaborator Dr. Pradeep K. Dudeja of the University of Illinois (Chicago, IL). In addition, the Western blot analysis was performed on the membranous proteins isolated from control and siRNA-pretreated Caco-2 cells. All membrane protein suspensions contained 1 mM PMSF, 1 µg/ml aprotinin, and 0.5 µg/ml leupeptin. Protein (150 µg) samples were treated with Laemmli sample buffer and resolved on a 10% SDS-polyacrylamide gel. After electrophoresis, the proteins were electroblotted onto an immunoblot polyvinylidene difluoride membrane (Bio-Rad) overnight, washed twice with PBS-Tween 20 for 10 min, and blocked with 5% dried milk in PBS-Tween 20. The membrane samples were then probed with either anti-human hTHTR-1-specific polyclonal antibodies (1:10,000 diluted in PBS-Tween 20) (23) or anti-human hTHTR-2-specific polyclonal antibodies (1:5,000 diluted in PBS-Tween 20) raised for us by Sigma Genosys. The polyclonal antibodies were raised against a specific sequence in the COOH terminus of the hTHTR-2 polypeptide (CENPDVSHPEEESNI) that corresponded with amino acids 477–490. Blots were then washed twice with PBS-Tween 20 buffer (Sigma) and reacted with goat anti-rabbit IgG conjugated to horseradish peroxidase (1:5,000 diluted in PBS-Tween 20) for 1 h at room temperature. The blots were finally washed twice with PBS-Tween 20 for 10 min each time, and the color was developed by using an enhanced chemiluminescence kit (Amersham).

Real-time PCR analysis. The real-time PCR was used to determine the level (amount) of endogenous hTHTR-1 and hTHTR-2 in Caco-2 cells. Total RNA was isolated from Caco-2 cells by using TRIzol reagent following the protocol described by the manufacturer (Life Technologies). Five micrograms of total RNA was reverse transcribed with oligo(dT) primers by using Superscript II (Life Technologies) following the manufacturer's procedures. After the reverse transcription, all samples were diluted with sterile water and three different dilutions were used for each real-time PCR assay (QuantiTect SYBRgreen PCR Kit, Qiagen, Valencia, CA). Real-time PCR was carried out based on Light Cycler technology to accurately analyze the differences in hTHTR-1 and hTHTR-2 gene expression in the Caco-2 cells. Gene-specific primers corresponding to the PCR targets were designed by using the specifications given by the vendors (Bio-Rad). Primers used for hTHTR-1, hTHTR-2, and {beta}-actin were as described earlier. Each SYBRgreen reaction (20 µl total volume) contained 9 µl of diluted cDNA as template. The amplification program consisted of 1 cycle of 95°C with a 30-s hold ("hot start") followed by 40 cycles of 95°C for 1 min, specified annealing temperature with 15-s hold, 72°C with 30-s hold for extension, and data acquisition. Amplification was followed by a melting curve analysis program run for one cycle at 95°C with 0-s hold, 65°C with 10-s hold, and 95°C with 0-s hold at the step acquisition mode. A negative control without cDNA template was run with every assay to assess specificity. In all cases, the final PCR products were analyzed on 4% agarose gels and data were expressed relative to human {beta}-actin.

Generation of hTHTR2-EGFP fusion protein, cell transfection, and confocal imaging. Coding region of the full-length hTHTR-2 was generated by PCR by using gene-specific primers (forward primer, 5'-CCGCTCGAGATGGATTGTTACAGAACTTCACTAAG-3' and reverse primer, 5'-CGGGATCCTTAGAGTTTTGTTGACATGATGATATTAC-3') and subcloned into pGEM-T vector. The clone was then amplified by using primers to fuse it with EGFP (5'-CCGCTCGAGATGGATTGTTACAGAACTTCACTAAG-3' and 5'-CGGGATCCGAGTTTTGTTGACATGATGATATTAC-3') under conditions specified previously (25). The PCR product as well as the EGFP-N3 vector were then digested with the restriction enzymes BamHI and XhoI, and the products were gel isolated and then ligated together to generate an in-frame fusion protein (hTHTR2-EGFP) with EGFP fused to the COOH terminus of hTHTR-2. The nucleotide sequence of each of these constructs was confirmed by sequencing (Laragen, CA). Generation of the hTHTR1-EGFP fusion protein has been described recently by us (25). Caco-2 cells were seeded onto collagen-coated filters (Corning Costar, Cambridge, MA) and grown until confluency. Individual filter dishes were transiently transfected ~5 days postconfluency in DMEM containing ~2 µg plasmid DNA. Cells were imaged 24–48 h later by using a Bio-Rad MRC 1024 confocal scanner attached to an Olympus Provis AX70 upright microscope equipped with a x40 water-immersion objective for imaging Caco-2 cells grown on filters. Fluorophores were excited by using the 488-nm line from an argon ion laser, and emitted fluorescence was monitored with a 530 ± 20-nm band-pass filter (EGFP).

Statistical analysis. All measurements were made on at least three different samples. Uptake data presented in this paper are means ± SE of multiple separate uptake determinations and were expressed by femtomoles or nanomoles per milligram protein per unit time. Statistical differences were evaluated by one-way ANOVA followed by Tukey's honestly significant difference test; comparison was made relative to simultaneously run controls. Kinetic parameters of the saturable thiamine uptake process determined by subtracting the diffusing component (determined from the slope of the uptake line at a high pharmacological concentration of thiamin of 1 mM and the point of origin from total uptake at each concentration examined) were calculated by using a computerized model of the Michaelis-Menten equation as described previously by Wilkinson (29).


    RESULTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Expression of the hTHTR-2 mRNA along the human gastrointestinal tract. In this study, we examined the expression of the hTHTR-2 mRNA along the length of the human gastrointestinal tract. The study was performed by means of Northern blot analysis by using a commercially available blot containing mRNA isolated from different native human gastrointestinal tissues (see MATERIALS AND METHODS). Results (Fig. 1) showed that the hTHTR-2 message was expressed, although at different levels, in all of these tissues. Two main bands were observed: a major band at ~4.0 kb and a minor band at ~3.5 kb. Relative expression of the major band in the different gastrointestinal tissues (normalized to {beta}-actin) was as follows: duodenum > jejunum > ileocecum > stomach > ascending colon > descending colon = transverse colon > ileum > rectum > cecum > esophagus.



View larger version (48K):
[in this window]
[in a new window]
 
Fig. 1. Expression of human thiamin transporter (hTHTR)-2 mRNA along the native human gastrointestinal tract and in Caco-2 cells. Northern blot analysis was performed by using a commercially available blot (Clontech) that contained 2 µg of poly(A)+ RNA from the indicated gastrointestinal tissue and from Caco-2 cells. The blot was probed with a randomly labeled hTHTR-2 cDNA and then with labeled human {beta}-actin cDNA.

 

Membrane expression of the hTHTR-2 protein in polarized human intestinal epithelial cells. In this study, we examined by means of Western blot analysis the expression of the hTHTR-2 protein at the brush-border membrane and the basolateral membrane domains of the native human enterocytes by using brush-border and basolateral membrane preparations isolated from the jejunum of organ donors by an established procedure (4, 5). We also examined expression of the hTHTR-2 protein in the membrane fraction of Caco-2 cells, our in vitro model system in these investigations (22). Specific polyclonal antibodies raised for us by a commercial vendor were used in this study (see MATERIALS AND METHODS). Results showed that the hTHTR-2 protein is expressed in the brush-border membrane but not in the basolateral membrane of the human jejunum intestinal epithelial cells (Fig. 2). Expression of the hTHTR-2 was also found in the membranous fraction of Caco-2 cells (Fig. 2). Specificity of the hTHTR-2 polyclonal antibodies was confirmed by preincubating the antibodies with the free synthetic peptide (against which the antibodies were raised), which competed out the specific band (Fig. 2).



View larger version (18K):
[in this window]
[in a new window]
 
Fig. 2. Expression of the hTHTR-2 protein in native human jejunal brush-border membrane (BBM), basolateral membrane (BLM), and in membranous fraction of Caco-2 cells. Western blot analysis was run by using 150 µg protein from native human jejunal brush-border membrane (lane 1) and BLM (lane 2) as well as from membranous fraction of Caco-2 cells (lanes 3 and 4). The blot was probed with specific polyclonal antibodies directed against a specific peptide of the hTHTR-2 protein and detected by using the enhanced chemiluminescence system. In lane 4, the polyclonal antibodies were first treated with antigenic peptide and then used to probe membranous fraction of Caco-2 cells.

 

To confirm the finding on the pattern of expression of the hTHTR-2 protein in the polarized human enterocytes, we used polarized Caco-2 cells grown on filter and transfected them with cDNA, encoding the fusion protein hTHTR2-EGFP. We then imaged the living Caco-2 cells by using a confocal microscope. The results (Fig. 3, A and C) showed the hTHTR2-EGFP fusion protein to be mainly expressed at the apical membrane domain of Caco-2 cells with little expression at the basolateral membrane domain of the cells. For comparison, we also resolved the polarity of expression of hTHTR-1, again fused with EGFP (hTHTR1-EGFP) under similar conditions. Expression of hTHTR-1 protein was found at both the apical and the basolateral membrane domains, with slightly higher expression at the basolateral compared with the apical membrane domain (Fig. 3, B and D).



View larger version (22K):
[in this window]
[in a new window]
 
Fig. 3. Distribution of hTHTR2-enhanced green fluorescent protein (EGFP) and hTHTR1-EGFP in Caco-2 cells grown on filters. Confocal images lateral (x-y) and axial (z) sections (white lines) showing expression of hTHTR2-EGFP (A) and hTHTR1-EGFP (B) in confluent Caco-2 monolayers grown on filters. Cells were imaged 24–48 h after transient transfection (see MATERIALS AND METHODS). C and D: relative fluorescence intensity of hTHTR2-EGFP and hTHTR1-EGFP, respectively, at the apical (AP) and basolateral (BL) membrane domains of Caco-2 cells.

 

Kinetic parameters of cloned hTHTR-2 in Caco-2 cells. Although the cDNA of the hTHTR-2 was cloned a few years ago (7, 16), no study is available describing the kinetic parameters (i.e., apparent Km and Vmax) of the thiamin uptake process by this system. To address this issue, we transiently expressed the hTHTR-2 cDNA in Caco-2 cells and examined thiamin uptake as a function of concentration (10–75 nM). Uptake of thiamin by the induced carrier-mediated system (calculated by subtracting uptake by control cells from that of cDNA-transfected cells) was determined at each thiamin concentration, and the kinetic parameters of the induced system were determined as described in MATERIALS AND METHODS. Results (Fig. 4) showed that the induced thiamin uptake process is saturable as a function of concentration with an apparent Km of 27.1 ± 7.9 nM, and a Vmax of 144.0 ± 17.1 fmol·mg protein-1·7 min-1.



View larger version (14K):
[in this window]
[in a new window]
 
Fig. 4. Initial rate of thiamin uptake by the induced carrier in Caco-2 cells transfected with hTHTR-2 cDNA. Cells were transiently transfected with 2.0 µg of hTHTR-2 cDNA. Initial rate [i.e., 7 min (22)] of thiamin uptake by confluent monolayers of Caco-2 cells as a function of concentration over the range of 10 to 75 nM was then examined in Krebs-Ringer buffer (pH 7.4). Kinetic parameters of the induced carrier system were determined as described in Kinetic parameters of cloned hTHTR-2 in Caco-2 cells. Data are means ± SE of at least 3 separate uptake determinations.

 

Evidence for existence of a saturable thiamin uptake system in human intestinal epithelial cells that operates in nanomolar concentration range. Little is known about thiamin uptake by human intestinal epithelial cells as a function of concentration over the nanomolar range. This is because most of the previous investigations on the subject have focused mainly on the micromolar concentration range to establish saturability (13, 22, 23). This includes the study from our laboratory with Caco-2 cells that reported an apparent Km of 3.18 ± 0.56 µM (22). The latter apparent Km is similar to the apparent Km of hTHTR-1 (2.5 ± 0.6 µM) following expression of cDNA of that system in HeLa cells (6). hTHTR-1 is also expressed in native human intestine and in Caco-2 cells (17). Because human intestinal epithelial cells (including Caco-2 cells) express hTHTR-2 and this transporter has an apparent Km in the nanomolar range (see above), we sought in this investigation to determine whether Caco-2 cells indeed posses a saturable system for thiamin uptake in the nanomolar concentration range. We therefore examined thiamin uptake by Caco-2 cells in the nanomolar range (10–75 nM) and observed clear saturation in the uptake process (Fig. 5). Kinetic parameters of the saturable system were calculated as described in MATERIALS AND METHODS and found to be 26.9 ± 4.2 nM and 177.7 ± 11.6 fmol·mg protein-1·7 min-1 for the apparent Km and Vmax, respectively.



View larger version (14K):
[in this window]
[in a new window]
 
Fig. 5. Initial rate of thiamin uptake by Caco-2 cells as a function of concentration in the nanomolar range. Confluent monolayers of Caco-2 cells were incubated in Krebs-Ringer buffer, pH 7.4, at 37°C for 7 min in the presence of 10–75 nM thiamin. Kinetic parameters of the carrier-mediated component were calculated as described in MATERIALS AND METHODS. Data are means ± SE of at least 3 separate uptake determinations.

 

Relative contribution of hTHTR-2 toward overall carrier-mediated thiamin uptake by human intestinal epithelial cells. Our aim in this study was to determine the relative contribution of the hTHTR-2 toward overall carrier-mediated thiamin uptake in human intestinal epithelial cells by using Caco-2 cells as the model. Because levels of hTHTR-2 and hTHTR-1 were determined individually in different studies by using different intestinal samples, we sought to first determine the relative expression of the two thiamin transporters in the same RNA samples of Caco-2 cells. This was performed by means of real-time PCR as described in MATERIALS AND METHODS. The results showed the level of expression of hTHTR-2 to be ~3.3-fold lower than that of hTHTR-1 in these cells. To determine the relative contribution of the hTHTR-2 toward overall carrier-mediated thiamin uptake by confluent monolayers of Caco-2 cells, we used a well-established approach to selectively and specifically silence the hTHTR-2 gene by means of small interfering RNA (siRNA) (1, 11, 15) then after 5 to 7 days examined the effect of such pretreatment on carrier-mediated thiamin uptake by confluent Caco-2 monolayers (see MATERIALS AND METHODS). First, we verified that the siRNAs used were able to silence the hTHTR-2 gene. This was performed by determining the level of the hTHTR-2 message by semiquantitative PCR in siRNA-pretreated and control cells. Results showed that pretreating Caco-2 cells with siRNAs led to a severe reduction in the endogenous hTHTR-2 mRNA level compared with control cells (Fig. 6A). The level of the human {beta}-actin mRNA, however, was not affected by the siRNA pretreatment, demonstrating the specificity of the effect (Fig. 6A). Furthermore, we also determined (by Western blot analysis) the level of hTHTR-2 protein in siRNA-pretreated and control Caco-2 cells and found the level to be substantially reduced in the siRNA-pretreated cells compared with controls (Fig. 8A). These findings confirm the effectiveness of the siRNA approach in silencing the individual thiamin transporter gene in these cells. We then examined the effect of pretreating the Caco-2 cells with hTHTR-2-specific siRNA on the initial rate [3 min (22)] of carrier-mediated uptake of a physiological concentration (15 nM) of thiamin across the apical membrane of confluent monolayers of these cells. Results showed that such pretreatments lead to a significant (P < 0.01; 48%) inhibition in carrier-mediated thiamin uptake compared with control (Fig. 7). When a similar approach of siRNA pretreatment was used to silence the gene of the other thiamin transporter, i.e., hTHTR-1, such pretreatment again led to a severe reduction in the endogenous hTHTR-1 RNA and protein levels (Figs. 6B and 8B, respectively) and to a significant (P < 0.01; 56%) inhibition in carrier-mediated uptake of thiamin (15 nM) (Fig. 7). When Caco-2 cells were pretreated with genespecific siRNAs against the hTHTR-1 and the hTHTR-2 simultaneously, a severe reduction of the individual endogenous mRNA and protein levels were observed (Figs. 6C and 8, A and B), and there was an almost complete inhibition of carrier-mediated thiamin (15 nM) uptake (Fig. 7). Uptake of the unrelated biotin (15 nM), however, was not affected by such siRNA pretreatment (112 ± 1.8 and 115 ± 3.1 fmol·mg protein-1·3 min-1 in cells pretreated simultaneously with a combination of hTHTR-2- and hTHTR-1-specific siRNAs, and control cells, respectively), indicating specificity of the observed effects.



View larger version (28K):
[in this window]
[in a new window]
 
Fig. 6. Semiquantitative PCR analysis of hTHTR-1 and hTHTR-2 mRNA in Caco-2 cells. Semiquantitative PCR was performed on total RNA isolated from control Caco-2 cells and those pretreated 5–7 days earlier with gene-specific small interfering RNA (siRNA). A: hTHTR-2-specific siRNA-pretreated cells (left, control). PCR products were analyzed between cycles 26 and 32 (linear range). Representative gel at 29 cycles is shown. B: hTHTR-1-specific siRNA-pretreated cells (left, control). PCR products were analyzed between cycles 22 and 28 (linear range). Representative gel at 26 cycles is shown. C: hTHTR-1- and hTHTR-2-specific siRNA-pretreated cells (left, control). PCR products were analyzed at 29 cycles.

 


View larger version (18K):
[in this window]
[in a new window]
 
Fig. 8. Western blot analysis of hTHTR-2 and hTHTR-1 proteins in Caco-2 cells. Western blot analysis was performed as described in MATERIALS AND METHODS. Level of expression of hTHTR-2 protein (A) and hTHTR-1 protein (B) in control cells and those pretreated with gene-specific siRNAs are shown. Data are representative of 3 separate sets of experiments.

 


View larger version (17K):
[in this window]
[in a new window]
 
Fig. 7. Initial rate of carrier-mediated thiamine uptake by control and siRNA-pretreated Caco-2 cells. Control cells and those pretreated with siRNA 5–7 days earlier were incubated at 37°C in Krebs-Ringer buffer, pH 7.4. [3H]thiamin (15 nM) was added to the incubation medium at the onset of incubation. Uptake was measured after 3-min incubation [i.e., initial rate (22)]. Data are means ± SE of at least 3 separate uptake determinations and were evaluated by one-way ANOVA followed by Tukey's honestly significant difference test (a and e vs. b, c, and d, P < 0.01; b vs. c, P < 0.05; b and c vs. d, P < 0.01).

 


    DISCUSSION
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The aim of this study was to examine the expression of the hTHTR-2 message along the length of the native human gastrointestinal tract and to determine the membrane domain of the polarized enterocyte at which the thiamin carrier protein is expressed. Our aim was also to determine the relative contribution of hTHTR-2 toward overall carrier-mediated thiamin uptake by the human intestinal epithelial cells. Results of these studies have shown that the message of hTHTR-2 is expressed along the length of the gastrointestinal tract with the highest level being in the proximal half of the human small intestine (duodenal/jejunal areas). Such a finding coincides with the higher ability of the proximal part of the small intestine to absorb dietary thiamin (20). It is also worth noting that the human large intestine represented by the colon also expresses this thiamin transporter. This again agrees with our previous findings on the existence of a carrier-mediated thiamin uptake system in human colonocytes (23). The latter uptake mechanism is believed to be responsible for absorption of the thiamin generated by the normal microflora of the large intestine (23).

Our studies have also examined the membrane domain of the polarized human enterocytes at which the hTHTR-2 protein is expressed. Two approaches were employed in these studies: an immunoblotting approach using native human jejunal brush-border membrane and basolateral membrane preparations and a confocal imaging approach using a fusion protein of hTHTR2-EGFP expressed in living human intestinal epithelial Caco-2 cells. Our Western blot analysis revealed that the hTHTR-2 protein is expressed only at the brush-border membrane domain of the human intestinal absorptive epithelial cells and not at the basolateral membrane domain of these cells. These findings were confirmed by the confocal imaging studies that showed expression of the hTHTR-2 protein to be mainly confined to the apical membrane domain of Caco-2 cells grown on filters. The latter findings with regards to expression of the hTHTR2-EGFP fusion protein at the apical membrane of Caco-2 cells was in contrast to the finding in the same cells with regards to expression of the other hTHTR, i.e., hTHTR-1. Expression of the hTHTR1-EGFP fusion protein was found to be at both the apical and the basolateral membrane domains of the filter-grown Caco-2 cells, with the expression being slightly higher at the latter compared with the former membrane domain. These results require further confirmation in intact native human intestinal tissues and indicate that although messages of both of the hTHTRs are expressed in human intestinal epithelial cells, the membrane domain at which their proteins are expressed is different. The hTHTR-2 protein appears to be expressed mainly at the apical brush-border membrane domain in these cells, whereas the hTHTR-1 protein is expressed at both the apical and the basolateral membrane domains. These findings may have implication as to the role of each of these transporters in the regulation of intestinal thiamin-absorption process and thiamin body homeostasis in health and disease. Indeed, recent studies in our laboratory (unpublished observations) have shown that these two thiamin transporters respond differently to regulatory conditions, with hTHTR-2 being more sensitive to regulation. Further studies are required to enhance our understanding of the physiology and regulation of these two systems in intestinal epithelial cells.

To determine the relative contribution of the hTHTR-2 toward overall carrier-mediated thiamin uptake by human intestinal epithelial cells, we used Caco-2 cells as a cellular model system and employed the recently developed siRNA as an approach to selectively silence this thiamin transporter. We then examined the effect of such a silencing of the hTHTR-2 on carrier-mediated thiamin uptake. Caco-2 cells were chosen because they express hTHTR-1 and hTHTR-2 at the RNA and protein levels. Pretreatment of Caco-2 cells with siRNA specific for the hTHTR-2 gene led to a substantial and specific silencing of the hTHTR-2 RNA and protein as indicated by PCR and Western blot analysis results shown in Figs. 6 and 8, respectively. Such a pretreatment also led to significant (P < 0.01; 48%) and specific inhibition in initial rate of uptake of a physiological concentration of thiamin. When a similar approach was used to silence hTHTR-1 in Caco-2 cells, again significant (P < 0.01; 56%) inhibition in carrier-mediated thiamin uptake was observed. When Caco-2 cells were simultaneously pretreated with siRNAs specific for hTHTR-1 and hTHTR-2, an almost complete inhibition in carrier-mediated thiamin uptake was observed. These findings suggest that both hTHTR-1 and hTHTR-2 play a role in carrier-mediated thiamin uptake by human intestinal epithelial cells and that together they account for total carrier-mediated thiamin uptake by these cells. How these two carrier systems are regulated and how they respond to physiological and pathological conditions requires further investigations.

In the present investigations, we also determined for the first time the kinetic parameters of thiamin uptake by the hTHTR-2 in human intestinal epithelial cells. The results showed uptake by this system to be saturable in the nanomolar range (apparent Km of 27.1 ± 7.6 nM). This is in contrast to the kinetic parameters of thiamin uptake mediated by hTHTR-1, a carrier that saturates in the micromolar range (apparent Km of 2.5 ± 0.6 µM) (6). Indeed, human intestinal epithelial Caco-2 cells appear to have both of these thiamin uptake systems functional as described above; this is further demonstrated by the findings of saturation in thiamine uptake in the nanomolar (Fig. 5) and micromolar (22) ranges.

In summary, results of these studies delineate the expression of the hTHTR-2 message along the gastrointestinal tract and show that the expression of the hTHTR-2 protein is mainly confined to the apical membrane domain of the polarized human enterocytes. In addition, hTHTR-1 and hTHTR-2 appear to play a role in the normal thiamin uptake process in human intestinal epithelial cells.


    ACKNOWLEDGMENTS
 
GRANTS

This study was supported by grants from the Department of Veterans Affairs and National Institute of Diabetes and Digestive and Kidney Diseases Grants DK-56061 and DK-58057 to (H. M. Said) and DK-63750 (to V. S. Subramanian).


    FOOTNOTES
 

Address for reprint requests and other correspondence: H. M. Said, Veterans Affairs Medical Center-151, Long Beach, CA 90822 (E-mail: hmsaid{at}uci.edu).

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


    REFERENCES
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Balamurugan K, Ortiz A, and Said HM. Biotin uptake by human intestinal and liver epithelial cells: role of the sodium-dependent multivitamin transport system, SMVT. Am J Physiol Gastrointest Liver Physiol 285: G73-G77, 2003.[Abstract/Free Full Text]
  2. Berdanier CD. Advanced Nutrition-Micronutrients. New York: CRC, 1998.
  3. Diaz GA, Banikazemi M, Oishi K, Desnick RJ, and Gelb BD. Mutations in a new gene encoding a thiamine transporter cause thiamine-responsive megaloblastic anaemia syndrome. Nat Genet 22: 309-312, 1999.[CrossRef][ISI][Medline]
  4. Dudeja PK, Tyagi S, Gill R, and Said HM. Evidence for carrier-mediated mechanism for thiamine transport to human jejunal basolateral membrane vesicles. Dig Dis Sci 48: 109-115, 2003.[CrossRef][ISI][Medline]
  5. Dudeja PK, Tyagi S, Kavilaveettil RJ, Gill R, and Said HM. Mechanism of thiamine uptake by human jejunal brush-border membrane vesicles. Am J Physiol Cell Physiol 281: C786-C792, 2001.[Abstract/Free Full Text]
  6. Dutta B, Huang W, Molero M, Kekuda R, Leibach FH, Devee LD, Ganapathy V, and Prasad PD. Cloning of the human thiamine transporter, a member of the folate transporter family. J Biol Chem 45: 31925-31929, 1999.[CrossRef]
  7. Eudy JD, Spiegelstein O, Barber RC, Wlodarczyk BJ, Talbot J, and Finnell RH. Identification and characterization of human and mouse SLC19A3 gene: a novel member of the reduced folate family of micronutrient transporter genes. Mol Genet Metab 71: 581-590, 2000.[CrossRef][ISI][Medline]
  8. Fennelly J, Frank O, Baker H, and Leevy CM. Peripheral neuropathy of the alcoholic. I. Aetiological role of aneurin and other B-complex vitamins. Brit Med J 5420: 1290-1292, 1964.[Medline]
  9. Fleming JC, Tartaglini E, Steinkamp MP, Schorderet DF, Cohen N, and Neufeld EJ. The gene mutated in thiamine-responsive anaemia with diabetes and deafness (TRMA) encodes a functional thiamine transporter. Nat Genet 22: 305-308, 1999.[CrossRef][ISI][Medline]
  10. Hammes HP, Du X, Edelstein D, Taguchi T, Matsumura T, Ju Q, Lin J, Bierhaus A, Naworth P, Hannak D, Neumaier M, Bergfeld R, Giardino I, and Brownlee M. Benfotiamine blocks three major pathways of hyperglycemia damage and prevents experimental diabetic retinopathy. Nat Med 9: 294-299, 2003.[CrossRef][ISI][Medline]
  11. Hannon GJ. RNA interference. Nature 418: 244-251, 2002.[CrossRef][ISI][Medline]
  12. Labay V, Raz T, Baron D, Mandel H, Williams H, Barret T, Szargel R, McDonald L, Shalata A, Nosaka K, Gregory S, and Cohen N. Mutations in SLC19A2 cause thiamine-response megaloblastic anaemia associated with diabetes mellitus and deafness. Nat Genet 22: 300-304, 1999.[CrossRef][ISI][Medline]
  13. Laforenza U, Patrini C, Alvisi C, Faelli A, Licandro A, and Rindi G. Thiamine uptake in human intestinal biopsy specimens, including observations from a patient with acute thiamin deficiency. Am J Clin Nutr 66: 320-326, 1997.[Abstract]
  14. Leevy CM and Baker H. Vitamins and alcoholism. Am J Clin Nutr 21: 325-328, 1968.
  15. Leirdal M and Sioud M. Gene silencing in mammalian cells by preformed small RNA duplexes. Biochem Biophys Res Commun 295: 744-748, 2002.[CrossRef][ISI][Medline]
  16. Rajgopal A, Edmondson A, Goldman D, and Zhao R. SLC19A3 encodes a second thiamine transporter ThTr2. Biochim Biophys Acta 1537: 175-178, 2001.[ISI][Medline]
  17. Reidling JC, Subramanian VS, Dudeja PK, and Said HM. Expression and promoter analysis of SLC19A2 in the human intestine. Biochim Biophys Acta 1561: 180-187, 2002.[ISI][Medline]
  18. Rindi G and Ferrari G. Thiamine transport by human intestine in vitro. Experientia 33: 211-213, 1977.[ISI][Medline]
  19. Rindi G, Patrini C, Laforenza U, Mandel H, Berant M, Viana MB, Poggi V, and Zarra AN. Further studies on erythrocytes thiamin transport and phosphorylation in seven patients with thiamin-responsive megaloblastic anaemia. J Inherit Metab Dis 17: 667-677, 1994.[ISI][Medline]
  20. Rindi G. Thiamine absorption in the small intestine. Acta Vitaminol Enzymol 6: 47-55, 1984.[Medline]
  21. Rogers LE, Porter FS, and Sidbury JB Jr. Thiamine-responsive megaloblastic anemia. J Pediatr 74: 494-504, 1969.[ISI][Medline]
  22. Said HM, Ortiz A, Kumar CK, Chatterjee N, Dudeja PK, and Rubin SA. Transport of thiamine in the human intestine: mechanism and regulation in intestinal epithelial cell model Caco-2. Am J Physiol Cell Physiol 277: C645-C651, 1999.[Abstract/Free Full Text]
  23. Said HM, Ortiz A, Subramanian VS, Neufeld EJ, Moyer MP, and Dudeja PK. Mechanism of thiamine uptake by human colonocytes: studies with cultured colonic epithelial cell line NCM460. Am J Physiol Gastrointest Liver Physiol 281: G144-G150, 2001.[Abstract/Free Full Text]
  24. Saito N, Kimura M, Kuchiba A, and Itokawa Y. Blood thiamin levels in outpatients with diabetes mellitus. J Nutr Sci Vitaminol (Tokyo) 33: 421-430, 1987.[Medline]
  25. Subramanian VS, Marchant JS, Parker I, and Said HM. Cell biology of the human thiamine transporter-1 (hTHTR1): intracellular trafficking and membrane targeting mechanisms. J Biol Chem 278: 3976-3984, 2003.[Abstract/Free Full Text]
  26. Tallaksen CME, Bohmer T, and Bell H. Blood and serum thiamin and thiamin phosphate esters concentrations in patients with alcohol dependence syndrome before and after thiamin treatment. Alcohol Clin Exp Res 16: 320-325, 1992.[ISI][Medline]
  27. Tanphaichirt V. Thiamin. In: Modern Nutrition in Health and Disease, edited by Shils ME, Olsen JA, and Shike M. New York: Lea and Febiger, 1994, p. 359-375.
  28. Victor M, Adams RD, and Collins GH. The Wernicke-Korsakoff Syndrome and Related Neurological Disorders Due to Alcoholism and Malnutrition. Philadelphia, PA: Davis, 1989.
  29. Wilkinson GN. Statistical estimation in enzyme kinetics. Biochem J 80: 324-332, 1961.[ISI][Medline]