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Cell Growth & Differentiation Vol. 13, 227-236, May 2002
© 2002 American Association for Cancer Research

10-Formyltetrahydrofolate Dehydrogenase, One of the Major Folate Enzymes, Is Down-Regulated in Tumor Tissues and Possesses Suppressor Effects on Cancer Cells1

Sergey A. Krupenko2 and Natalia V. Oleinik

Department of Biochemistry and Molecular Biology, Medical University of South Carolina, Charleston, South Carolina 29425


    Abstract
 TOP
 Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
 References
 
Our studies showed that an abundant folate enzyme, 10-formyltetrahydrofolatedehydrogenase (FDH), is strongly down-regulated in several types of cancer on both the mRNA and the protein level. Transient expression of FDH in several human prostate cancer cell lines, a hepatocarcinoma cell line, HepG2, and a lung cancer cell line, A549, suppressed proliferation and resulted in cytotoxicity. In contrast, overexpression of a catalytically inactive FDH mutant did not inhibit proliferation, which suggests that the suppressor effect of FDH is a result of its enzymatic function. Because the FDH substrate, 10-formyltetrahydrofolate, is required for de novo purine biosynthesis, we hypothesized that the inhibitory effects of FDH occur through the depletion of intracellular 10-formyltetrahydrofolate followed by the loss of de novo purine biosynthesis. The ultimate impact is diminished DNA/RNA biosynthesis. Indeed, supplementation of FDH-overexpressing cells with 5-formyltetrahydrofolate or hypoxanthine reversed the FDH growth-inhibitory effects. Hence, down-regulation of FDH in tumors is proposed to be one of the cellular mechanisms that enhance proliferation.


    Introduction
 TOP
 Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
 References
 
Folate coenzymes are involved in several major cellular processes including biosynthesis of precursors for DNA and RNA (1, 2, 3) . Because of the crucial role of folates in cell function, disruption of folate metabolism results in the impairment of cell division (1) . Rapidly dividing cells such as cancer cells are critically dependent on an abundant supply of fully reduced folate, and, thus, folate metabolism is essential for cell proliferation (4) . Therefore, for many years, key enzymes of folate metabolism have been targeted as a means to prevent tumor development (4, 5, 6, 7) . Although many enzymes are involved in folate metabolism (1 , 8) , to date only relatively few of them have been identified as important elements of tumor proliferation and, accordingly, potential anticancer targets (4, 5, 6, 7 , 9) . The role of FDH,3 one of the most abundant folate enzymes, has yet to be explored in cell proliferation. FDH comprises about 1% of the total pool of soluble cell protein in liver cytosol (8) . It converts 10-formyl-THF to THF, thus regulating two major intracellular reduced-folate pools (10) . The physiological role of FDH is not completely clear. Because the FDH substrate, 10-formyl-THF, is involved in two reactions of de novo purine biosynthesis, it is believed that the enzyme serves to recycle 10-formyl-THF that is not required for purine biosynthesis back to the THF pool, where folate is available for other one-carbon reactions (1) . The dehydrogenase reaction carried out by FDH removes carbon units from the folate pool in the form of CO2. Therefore, it has also been postulated that the enzyme serves to remove excess one-carbon units from the folate pool of the cell (11 , 12) . However, the extent of the "excess one-carbon units" has never been defined nor experimentally tested. It has also been proposed that the enzyme plays a key role in the metabolism of formate by removing it as CO2, thus protecting the cell from formate intoxication (13) . On the basis of the fact that the FDH substrate, 10-formyl-THF, is necessary for two reactions of de novo purine biosynthesis, we have suggested that FDH might regulate purine biosynthesis through control of the level of 10-formyl-THF. De novo purine biosynthesis is especially active in the liver, whereas nonhepatic tissues, other than placenta, are capable of only limited de novo synthesis of purines (14) . It has been assumed that the needs of other tissues are supplied by de novo biosynthesis in the liver (15) . This explains the high levels of FDH in liver. Not much is known, however, about the role of FDH in tissues other than liver. In particular, the function of the enzyme in cancer cells has never been addressed. The present studies were undertaken to examine the influence of FDH on cancer cell growth.


    Results
 TOP
 Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
 References
 
Tissue Distribution of FDH mRNA.
To address the question of whether FDH expression is tissue specific, we estimated the levels of FDH message in several normal and tumor human tissues. To accomplish this task, we applied PCR techniques and used commercially available human cDNA panels (Clontech). The level of a particular cDNA reflects the level of corresponding mRNA and, hence, the tissue distribution of the protein. The commercial panels are normalized with regard to cDNA levels using four constitutively expressed genes. Therefore, these panels allow direct comparison of message levels between different tissues. We found that levels of FDH message vary significantly, which suggested that the enzyme expression is tissue specific (Fig. 1)Citation . Three tissues (liver, kidney, and pancreas) have the highest level of FDH message. The high level of FDH message in liver and kidney evidently reflects the fact that these are specialized organs of folate metabolism (16) . Some other folate enzymes are also highly expressed in these two organs (16, 17, 18, 19) . Lung, prostate, brain, skeletal muscle, heart, ovary, thymus, and testis displayed a moderate level of FDH message. Several tissues, including placenta, spleen, colon, small intestine, and leukocytes, revealed very low or nondetectable levels of FDH message. The tissue expression pattern of FDH is not unusual: numerous studies revealed that enzymes involved in folate metabolism display a tissuespecific expression (16, 17, 18, 19, 20, 21) . Interestingly, our experiments revealed uniformly low FDH mRNA levels in tumors, compared with normal tissues, which suggested strong down-regulation of the enzyme.



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Fig. 1. Levels of FDH message in normal and tumor human tissues estimated by PCR techniques. Top row, levels of FDH message; bottom row, the level of message for constitutively expressed glyceraldehyde-3-phosphate dehydrogenase. Lung (1), muscle (2), liver (3), kidney (4), heart (5), placenta (6), spleen (7), thymus (8), small intestine (9), leukocytes (10), testis (11), ovary (12), lung (13), lung carcinomas LX-1 (14), lung carcinoma GI-117 (15), pancreas (16), pancreatic adenocarcinoma GI-103 (17), prostate (18), and prostatic adenocarcinoma (19) were evaluated. Carcinomas are marked according to Clontech.

 
FDH Levels in Normal versus Tumor Tissues.
To further explore the question of whether normal and tumor tissues differ in the levels of FDH, we performed immunohistochemical staining of tissue preparations with FDH-specific antiserum. In these experiments, we used commercially available hybrid-ready slides (Novagen), which were suitable for immunohistochemical localization of proteins with antibody probes, and antiserum raised against rat FDH. Our preliminary experiments showed that this antiserum interacts with human FDH as well as with rat FDH, obviously because of high sequence similarity (92%) between the two enzymes. Normal human liver, lung, prostate, pancreas, and ovary tissues were compared in these experiments with preparations obtained from tumors that arose from the tissues of the same origins. In these experiments, we evaluated a total of 15 preparations from tumors of different origin. Intense staining with FDH-specific antiserum was observed in all of the normal tissues (Fig. 2Citation ; pancreatic preparations are not shown), which indicated substantial levels of FDH. In contrast, all of the preparations from tumor tissues showed significantly decreased staining, almost at background levels, which indicated a strong down-regulation of FDH. A selection of 8 from a total of 15 tissues examined is shown in Fig. 2Citation (the other 7 tissues displaying the same effects are not shown). Using in situ hybridization on the same tissue preparations that were used for immunostaining, we compared FDH mRNA levels in normal versus tumor tissues. These experiments indicated significantly decreased FDH mRNA levels in all of the tumors compared with the normal tissues. Eight of a total of 15 tissues examined are shown in Fig. 3Citation ; the remaining tissues exhibited the same effects (not shown). Overall, our experiments showed an ubiquitous lack of both FDH and its message in all of the tumors examined.



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Fig. 2. Immunohistochemical staining of human tissues with FDH-specific antiserum. For each tissue: top row, staining with FDHspecific antiserum (AS); bottom row, control staining with preimmune serum (PI). Brown color, the presence of FDH. For each sample, a representative area from the middle of tissue section is shown. x200.

 


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Fig. 3. In situ hybridization of human tissues with FDH mRNA antisense probe. For each tissue: top row, staining with the antisense probe (A-Pr); bottom row, control hybridization with sense oligonucleotide (S-Pr) complimentary to the antisense probe. White spots correspond to FDH mRNA. For each sample, a representative area from the middle of the tissue section is shown. x200.

 
Comparison of FDH Message Levels in Matched Pairs.
We also estimated FDH mRNA levels in matched samples from normal and tumor tissues. These are the samples of cDNAs obtained from mRNAs isolated from normal and tumor counterpart tissues of the same individual. Analysis of such pairs allows one to directly compare expression of genes in tumor versus normal tissues. Eight preparations from lung were examined in these experiments (Fig. 4)Citation . We have found that in five cases (pairs 2, 3, 4, 5, and 8) an FDH message was essentially absent in tumor but not in normal counterpart tissue. In three pairs the presence of FDH cDNA was not detected in normal tissue or cancer tissue (pairs 1, 6, and 7). Analysis of matched cDNA pairs from individuals with breast and ovarian cancers also revealed either down-regulation of FDH expression in tumor tissues compared with corresponding normal tissues or very low FDH message in both normal and tumor tissues (Fig. 4)Citation . These experiments showed that, whereas there are variations between individuals in FDH expression in normal tissues, tumor tissues have uniformly low or undetectable levels of FDH mRNA, which indicates strong down-regulation of FDH expression.



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Fig. 4. Comparison of levels of FDH mRNA in normal and tumor tissues. Each pair of tissues was obtained from the same individual. For each pair: upper row, levels of FDH message; bottom row, message levels for constitutively expressed glyceraldehyde-3-phosphate dehydrogenase. Lung: pairs 1–3 and 5–7, squamous cell carcinoma; pairs 4 and 8, adenocarcinoma. Breast: pairs 1, 3, and 4, lobular carcinoma; pair 2, tubular adenocarcinoma. Ovary: papillary serous carcinoma (1), serous cystadenocarcinoma (2), adenocarcinoma (3). N, normal tissue; T, tumor tissue.

 
FDH Level in Normal and Regenerating Rat Liver.
To explore whether the FDH inhibitory effect is tumor specific or is associated with proliferation in general, we measured FDH levels in normal and regenerating rat liver. This was done on whole liver samples by immunoblot techniques with FDH-specific antiserum. In normal adult rat liver, 60% of the cells are hepatocytes, and the remainder are various types of nonparenchymal cells (22) . The hepatocytes, however, are much larger and constitute about 90% of the liver volume (22) . Regeneration of cells within the remnant liver after a partial hepatectomy is initiated by hepatocytes, with peak DNA synthesis between 24 and 48 h (23 , 24) . Other cells of the liver enter into DNA synthesis 24 h after hepatectomy, with a peak of DNA synthesis at 48 h or later (23 , 24) . Therefore, liver samples 24 h, 48 h, 96 h, and 7 days posthepatectomy, and a control sample from normal nonregenerating liver, were analyzed in our experiments. These experiments revealed that levels of FDH were similar in all of the preparations (Fig. 5)Citation .



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Fig. 5. Levels of FDH in normal and regenerating rat liver. FDH was detected by immunoblot techniques using FDH-specific polyclonal antiserum in liver samples obtained 24 h, 48 h, 96 h, and 7 days after partial (70%) hepatectomy. The control sample represents the level of FDH in resting (nonmanipulated) rat liver. About 15 µg of the total cytosolic liver protein was loaded per each lane.

 
Elevation of FDH in Cultured Cells.
To test the influence of FDH on cancer cells, we elevated FDH in different cell lines using transient expression. The prostate cancer PPC-1, lung cancer A549, hepatocarcinoma HepG2, and transformed kidney embryonic 293A cell lines were used in these experiments. All of these cell lines are of human origin. Immunoblot assays with FDH specific antiserum showed that all of these cell lines except for 293A did not express FDH (Fig. 6A)Citation . Consistent with this, only 293A cells displayed FDH activity (data not shown). Cells were transfected with FDH cDNA cloned into pcDNA 3.1+ vector using LipofectAMINE. Transfected cells were selected for resistance to neomycin, which enabled the elimination of nontransfected cells. In control experiments, cells were transfected with the vector that did not contain the FDH cDNA insert (that is, with the "empty" vector). Immunoblot assay with FDH specific antiserum showed significant levels of expressed FDH in all of the cell lines beginning 48 h posttransfection (Fig. 6A)Citation . We observed that expression of FDH inhibited the proliferation of all of the cell lines except for 293A, whereas proliferation of control cells transfected with the empty vector was not inhibited (Fig. 6B)Citation . Overexpression in the same cell lines of catalytically inactive FDH mutant, C707A (25) , did not inhibit cell proliferation (Fig. 6B)Citation . This suggests that the observed effects of FDH are the result of its enzymatic activity. Similar results were obtained with three other prostate cell lines, Tsu-Pr1, Du-145, and PC-3. Expression of the wild-type FDH inhibited cell proliferation, whereas expression of the C707A mutant did not affect the cells (data not shown).



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Fig. 6. Elevation of FDH in cultured cells by transient expression. A, levels of FDH in nontransfected cells (Lanes 1) and transfected cells expressing wild type FDH (Lanes 2) or C707A FDH mutant (Lanes 3), assayed by immunoblot techniques (see "Materials and Methods" for details). B, influence of transient FDH expression on cell proliferation. Lines 1 ({image}), cells transformed with empty vector pcDNA3.1+ (control); lines 2 ( ), cells transformed with vector expressing wild-type FDH; and lines 3 (····), cells transformed with vector expressing C707A FDH mutant. The type of cell line is marked on the blot and graphs.

 
Reversal of FDH Inhibitory Effects.
To examine whether suppressor effects of FDH can be reversed by media supplementation with high concentrations of folate and/or purine, we grew transiently transfected cell in the presence of 10 µM 5-formyl-THF or 100 µM hypoxanthine, or their combination. Three cell lines, A549, PPC-3, and PC-1, were used in these experiments. We found that, in the presence of either 5-formyl-THF or hypoxanthine, all of the studied cells were able to form clones capable of FDH expression (Fig. 7, A and C)Citation . In contrast, there was no cell survival without supplementation (Fig. 7A)Citation . Supplementation with 5-formyl-THF resulted in slightly better rescue than did supplementation with hypoxanthine, whereas the strongest effect was observed with the combination of folate and purine (Fig. 7B)Citation . Although all of the cell lines were able to form clones when supplemented with folate/purine, A549 cells had the highest survival rate (Fig. 7B)Citation .



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Fig. 7. Reversal of FDH inhibitory effects by 5-formyl-THF and hypoxanthine. A, clonal growth assay of PPC-1 cell line expressing FDH. B, colony formation by cell lines in rescue experiments; average of a duplicate is shown for each value. C, levels of FDH in cells rescued with hypoxanthine (HX), 5-formyl-THF (F), or their combination (HX+F).

 

    Discussion
 TOP
 Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
 References
 
The present studies revealed that FDH expression is highly tissue specific. Moreover, we observed a lack of FDH and FDH mRNA in several types of cancer, although they were present in corresponding normal tissues. We have further demonstrated that transient FDH expression suppresses the proliferation of some types of cancer cells. This suggests that a lack of FDH in tumor tissues is not a coincidental deletion of a nonessential enzyme but rather is associated with neoplastic transformation. Whereas some metabolic enzymes possess regulatory functions different from their catalytic functions (26 , 27) , such a noncatalytic regulatory function has not been demonstrated for FDH. Hence, the fact that expression of the catalytically inactive FDH mutant, C707A, does not produce growth-inhibitory effects strongly supports the concept that the growth inhibition is a result of FDH enzymatic activity.

There are clearly many changes in protein expression during neoplastic transformation. Microarray analysis has shown drastically different patterns of gene expression between normal and cancer tissues at the transcript level (28 , 29) . However, there are not as many examples of the lack of a critical metabolic enzyme such as FDH. The lack of FDH is remarkably widespread in tumors of different origin, which suggests that down-regulation of the enzyme is an important factor of tumor proliferation. Tissue distribution of folate enzymes, as well as changes in enzyme expression during neoplastic transformation, suggests that they are tightly regulated in the cell (16, 17, 18, 19, 20, 21) . Changes in expression of folate-related proteins are best studied in connection with cellular response to antifolate treatment. Thus, alterations in expression of certain genes of folate-related proteins targeted by antifolates are one of the mechanisms of tumor cell resistance (4 , 5 , 30) . In most cases, however, in contrast to FDH, an increase in the production of folate enzymes is observed (5 , 16 , 19 , 21) .

Although FDH is one of the most abundant folate enzymes, it is not among the proteins the role of which has been extensively evaluated in cellular metabolism. The only known function of FDH is to catalyze the conversion of 10-formyl-THF to THF. As such, this reaction can regulate these two major reduced folate pools. Mechanistically, increased FDH levels will result in decreased intracellular 10-formyl-THF concentrations, whereas depletion of the enzyme, conversely, should result in an increase of the 10-formyl-THF pool. Therefore, down-regulation of FDH in tumors is in agreement with the observation that tumor tissues have increased concentrations of 10-formyl-THF compared with normal tissues (31) . Deletion of FDH in mice leads to low THF and increased 10-formyl-THF levels (32) , which supports a role for the enzyme in the regulation of the two reduced folate pools. The FDH substrate, 10-formyl-THF, is required for de novo purine biosynthesis (3) . Rapidly proliferating tissues have increased demand for nucleic acid biosynthesis and, hence, require high levels of nucleic acid precursors including purines (33) . Therefore, in general, the key enzymes of the de novo and salvage pathways of purine biosynthesis are expected to be increased, and the opposing catabolic enzymes decreased, during malignant transformation and tumor progression (33) .

FDH was first described as an abundant protein of liver cytosol (10 , 11 , 34) . According to some reports, it comprises ~1% of the total cytosolic protein in liver (8) . It is not known whether intracellular FDH is lower in proliferating hepatocytes and at what stage of liver development FDH reaches these high levels. It would be expected, however, that if observed FDH down-regulation in tumors is associated with proliferation in general, in regenerating liver, the enzyme is also down-regulated. Subsequently, high levels of FDH in nonproliferating hepatocytes should be reached by upregulation during differentiation. Yet, our study revealed consistently high levels of FDH in regenerating liver, which indicated that, in this system, proliferation is not accompanied by enzyme down-regulation. These results further suggest that FDH down-regulation in tumors might not be associated simply with proliferation.

In de novo purine biosynthesis, 10-formyl-THF donates two carbon atoms to the purine ring in two reactions catalyzed by GART and AICART (3) . Inhibition of these enzymes by antifolates has shown antitumor activity in cell culture and animal studies (35 , 36) . Antifolates that target GART/AICART block de novo purine biosynthesis, which results in a rapid drop of intracellular ATP and GTP pools (35 , 37) . Likewise, depletion of the intracellular 10-formyl-THF pool would be expected to generate an effect similar to that of the inhibition of GART/AICART with regard to purine biosynthesis and, correspondingly, with regard to cancer cell proliferation. The enzyme reaction carried out by FDH is a direct way to deplete intracellular 10-formyl-THF. Elevated FDH in vivo should reduce 10-formyl-THF in the cell, making it unavailable for biosynthetic reactions and, thus, inhibiting de novo purine biosynthesis and cell proliferation. As a secondary effect, elevated FDH may also inhibit DNA repair, resulting in accumulation of DNA damage. It is also likely that it will impact the energetic potential and rates of phosphorylation in the cell by diminishing the levels of ATP and GTP. Therefore, multiple inhibitory effects can result from elevated FDH. The proposed mechanism rationalizes down-regulation of the enzyme in tumor tissues: FDH levels in normal tissues are too high to allow de novo purine biosynthesis adequate to the increased demand of cancer cells for DNA/RNA synthesis. Therefore, depletion of the enzyme in tumors could be one of the mechanisms to support high intracellular 10-formyl-THF levels.

We propose that suppressor effects of FDH is similar to the effects of antifolates targeting purine metabolism and occur through depletion of the intracellular 10-formyl-THF pool followed by arrest of the de novo purine biosynthesis. The antiproliferative effects of antifolates targeting GART, with resultant inhibition of the de novo purine biosynthesis, can be ablated by supplying enough exogenous purine to maintain intracellular pools via salvage pathways (38) . It has been shown that preformed purine, hypoxanthine, or the purine precursor, aminoimidazole carboxamide, can completely reverse the effects of the GART inhibitor DDATHF on purine pools and can protect cells against DDATHF growth suppression (37) . It has also been shown that 5-formyl-THF protects cells from the suppressor effects of GART inhibitors (35) . In the cell, 5-formyl-THF is readily converted to 10-formyl-THF, and, therefore, supplementation of cultured cells with 5-formyl-THF would be expected to increase intracellular 10-formyl-THF levels (39 , 40) . Similar to experiments using antifolates, FDH suppressor effects were at least partially overcome by the addition of excess 5-formyl-THF or hypoxanthine, which suggests that FDH cytotoxicity occurs through the disturbance of intracellular folate/purine pools.

It is interesting that FDH overexpression did not inhibit the proliferation of the human embryonic kidney cell line 293A. This cell line, in contrast to the other cell lines examined, expresses significant levels of endogenous FDH. Because this cell line is of a noncancerous origin, this supports the concept that the FDH-inhibitory effects are tumor cellspecific. Another explanation of this phenomenon could be that these cells are less dependent on de novo purine biosynthesis but, rather, use an alternative salvage pathway to supply purines for DNA/RNA synthesis (33) . The extent to which purine salvage has an impact on cellular metabolism is not clear (38) . Experiments using radioactively labeled nucleic acids as metabolic tracers have demonstrated that little of the nucleosides ingested in the diet are incorporated into cellular nucleic acids (41 , 42) . These findings suggest that the de novo pathway of nucleotide biosynthesis is the primary source of nucleic acid precursors. However, cells of various types are expected to be different in their relative dependence on de novo versus salvage purine pathway.

Thus, FDH appears to be ubiquitously depleted in tumors and possesses the ability to inhibit cancer cell proliferation when overexpressed, which, in turn, implies that the enzyme could be a critical growth regulator. It is most likely that the inhibitory effect of FDH on proliferation is a result of direct influence on cellular metabolism of rapidly proliferating cells. At present, it is not clear whether the FDH loss in cells that normally express the enzyme will promote cell transformation or whether FDH down-regulation is a secondary effect in initiated cells. It is also not clear whether the inhibitory effect of FDH is entirely tumor cell-specific or whether it is more generally associated with proliferating cells. The high levels of the enzyme observed in both resting and regenerating liver, however, as well as the absence of inhibitory effects on the human embryonic kidney cell line, implies that the FDH suppressor effects may be tumor specific.


    Materials and Methods
 TOP
 Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
 References
 
Materials.
The pcDNA 3.1+ expression vector was obtained from Invitrogen (Carlsbad, CA). Tissue slides were purchased from Novagene (Madison, WI). Panels of cDNAs and matched cDNA pairs were obtained from Clontech (Palo Alto, CA). Pfu Turbo DNA polymerase was purchased from Stratagene (La Jolla, CA). Restriction enzymes were from New England Biolabs (Beverly, MA). Biochemical reagents were purchased from Sigma (St. Louis, MO). Cell media and reagents were purchased from Life Technologies, Inc. (Rockville, MD). Five matched cDNA pairs were a kind gift from Dr. Besim Ogretmen, Medical University of South Carolina (Charleston, SC).

PCR.
Amplifications were performed in a Stratagene RoboCycler thermal cycler in 250-µl PCR tubes (Perkin-Elmer, Norwalk, CT) using Pfu Turbo DNA polymerase. The total volume of the reaction mixture was 50 µl. To ensure exponential amplification, thus enabling the accurate detection of the FDH message, conditions for PCR experiments were empirically determined. The optimal amplification was achieved in 33 cycles with the following cycle parameters: denaturation at 94°C for 1 min, annealing at 65°C for 1 min, and extension at 68°C for 3 min. The runs also included an initial 1-min hold at 94°C and a final 10-min hold at 68°C. Samples (10 µl from each PCR mixture) were analyzed by standard horizontal gel electrophoresis in 1.0% agarose containing ethidium bromide and were visualized using a Bio-Rad imaging system (Hercules, CA).

Immunohistochemical Staining.
Immunohistochemical staining was performed using polyclonal antibodies raised against rat FDH. Before staining, tissue slides were deparaffinized in xylene and rehydrated in descending concentrations of ethanol (100–35%) in water. Endogenous peroxidase activity was blocked by treating the slides with 0.3% solution of hydrogen peroxide in 100% methanol for 30 min at room temperature. The slides were treated with unmasking solution (Vector laboratories, Burlingame, CA), washed with PBS (Life Technologies, Inc.), and incubated at room temperature in PBS containing 1.5% normal goat serum for 1 h and then for 1.5 h in PBS containing 1.5% normal goat serum and FDH-specific antiserum (1:600 dilution). The staining procedure was further carried out using Vectastain Elite ABC kit, a peroxidase-based detection system (Vector Laboratories), according to the manufacturer’s directions. The final peroxidase-labeled complex was visualized using a mixture of 0.01% 3,3'-diaminobenzidine tetrahydrochloride and 0.01% hydrogen peroxide. The tissue sections were counterstained by hematoxylin, dehydrated, mounted with coverslips, and examined using an Axioskop 20 upright microscope (Carl Zeiss, Oberkochen, Germany). In the control experiments, tissue slides were subjected to the same procedure using preimmune serum instead of FDH-specific antiserum.

In Situ Hybridization.
To generate a probe for in situ hybridization, the translated region of human FDH cDNA was amplified from cDNA library using PCR techniques and cloned into pRSET vector (Invitrogen). A 250-bp fragment was excised and recloned in the opposite orientation immediately downstream of T7 promoter to allow the synthesis of the antisense strand. The vector was linearized by cutting at the end of the 250-bp sequence to stop the synthesis at this point. The 250-bp antisense mRNA probe for hybridization was synthesized with T7 reverse transcriptase and digoxigenin-labeled UTP using a DIG RNA labeling kit (Roche Molecular Biochemicals, Indianapolis, IN). Hybridization was performed as described elsewhere (43) using ELF 97 mRNA in situ hybridization kit (Molecular Probes, Eugene, OR) and Sure site hybridization kit (Novagen). A digoxigenin-labeled probe was stained with digoxigenin-specific antibodies conjugated with biotin (step I), followed by staining with streptavidin conjugated with alkaline phosphatase (step II). The final complex was visualized by incubation with an alkaline phosphatase substrate that yields an insoluble colored product. The control (sense) probe was generated by the same procedure. The staining preparations were examined using an Olympus IX70 fluorescence microscope.

Cell Lines.
The lung carcinoma cell line A549, hepatocarcinoma cell line HepG2, kidney transformed embryonic cell line 293A, prostate carcinoma cell line DU-145, and prostate adenocarcinoma cell line PC-3 were obtained from American Type Culture Collection. The prostate carcinoma cell lines Tsu-Pr1 and PPC-1 were a kind gift from Dr. James S. Norris, Medical University of South Carolina. The A549, 293A, Tsu-Pr1, and PPC-1 cell lines were maintained in RPMI 1640 supplemented with 10% heat-inactivated fetal bovine serum, 2 mM glutamine and 1 mM sodium pyruvate (complete medium). The PC-3 and DU-145 cell lines were maintained in the complete medium supplemented with 1.5 g/liter sodium bicarbonate and 0.1 mM nonessential amino acids. The HepG2 cell line was propagated in MEM Eagle with 2 mM L-glutamine supplemented with 10% fetal bovine serum and Earle’s PBS adjusted containing 1.5 g/liter sodium bicarbonate, 0.1 mM nonessential amino acids, and 1.0 mM sodium pyruvate. All of the cells were grown at 37°C under humidified air containing 5% CO2.

Detection of FDH in Cultured Cells.
Attached cells (~1 x 106) were washed from culture medium by rinsing in PBS and were lysed by adding 50 mM Tris-HCl buffer (pH 8.0) containing 0.15 M NaCl, 2 mM EDTA, 1% Triton X-100, and protease inhibitors (Sigma). FDH was determined in the cell lysate by immunoblotting with FDH-specific polyclonal antiserum (44) using an ECL kit (Amersham Pharmacia Biotech, Piscataway, NJ).

Detection of FDH in Liver Samples.
Samples of regenerating and normal rat liver were kindly provided by Dr. Donna Beer Stolz (University of Pittsburgh School of Medicine, Pittsburgh, PA). The samples were obtained from Fisher 344 male rats weighing ~200–220 g at 24 h, 48 h, 96 h, and 7 days after partial (70%) hepatectomy. Frozen liver samples were homogenized with Polytron in 4 volumes of 0.25 M sucrose containing 10 mM ß-mercaptoethanol and protease inhibitor mixture (Sigma). The homogenate was centrifuged at 17,000 x g for 15 min. The supernatant was subjected to SDS-PAGE followed by immunoblot with FDH-specific polyclonal antiserum (44) using an ECL kit.

Expression Vector Construction.
Rat liver FDH cDNA was excised from pVL1393/FDH vector (25) and recloned into pcDNA3.1+. pVL1393/FDH vector was cut with XbaI restriction endonuclease, treated with Pfu Turbo DNA polymerase to create blunt ends, and then cut with EcoRI restriction endonuclease to yield linear FDH cDNA having blunt 5'-end and EcoRI-cohesive 3'-end. Correspondingly, pcDNA3.1+ vector was treated with HindIII restriction endonuclease, Pfu Turbo DNA polymerase, and EcoRI restriction endonuclease to yield linear vector having blunt 3'-end and EcoRI-cohesive 5'-end. The two fragments were ligated using a Rapid DNA Ligation kit (Roche Molecular Biochemicals).

Transient Expression of FDH in Mammalian Cells.
Cells (~2 x 105) were transfected with the pcDNA3.1/FDH construct using LipofectAMINE Plus reagent (Life Technologies, Inc.) or FuGENE 6 (Roche Diagnostic, IN) according to the manufacturer’s directions. Antibiotic G418 (Sigma) was added to the culture medium at a concentration of 500 µg/ml 48 h later to allow the selection of transfected cells. Every 24 h, cells were detached by treatment with trypsin and counted with a hemocytometer. Experiments were performed in triplicate, and data were expressed as the mean of the triplicates. FDH expression was detected by immunoblotting analysis as described above.

Clonal Growth Assay.
Cells transfected with FDH as described above were grown in medium supplemented with 10 µM 5-formyl-THF, or 100 µM hypoxanthine, or their combination. In control plates, cells were grown on regular medium without supplementation. Formation of clones was detected 12 days posttransfection by staining with crystal violet (45) . The medium was removed from the plates, the cells were rinsed with PBS, fixed with 4% formaldehyde for 1 h, stained with 0.2% crystal violet for 20 min, and the clones were counted. In a parallel experiment, clones were examined for FDH expression. FDH was measured by SDS-PAGE, followed by immunoblot with FDH-specific antiserum and an ECL kit as described above. All of the experiments were performed in duplicate.


    Acknowledgments
 
We thank Dr. David G. Priest for helpful discussion and critical reading of the manuscript, Dr. Besim Ogretmen for the gift of lung tumor cDNA samples, and Dr. Donna Beer Stolz for the kindly provided samples of regenerating rat liver.


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

1 Supported by NIH Grant DK54388 and a grant from the South Carolina Commission on Higher Education Research Initiative. Back

2 To whom requests for reprints should be addressed, at Department of Biochemistry and Molecular Biology, Medical University of South Carolina, 173 Ashley Avenue, Room 512-B BSB, Charleston, SC 29425. Phone: (843) 792-4321, extension 17; Fax: (843) 792-4322; E-mail: krupenko{at}musc.edu Back

3 The abbreviations used are: FDH, 10-formyl-THF dehydrogenase; THF, tetrahydrofolate; GART, glycinamide ribonucleotide formyltransferase; DDATHF, 5,10-dideaza-5,6,7,8-THF; AICART, 5-aminoimidazol-4-carbox-amide ribonucleotide formyltransferase. Back

Received for publication 12/20/01. Revision received 3/13/02. Accepted for publication 3/20/02.


    References
 TOP
 Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
 References
 

  1. Wagner C. Biochemical role of folate in cellular metabolism Bailey L. B. eds. . Folate in Health and Disease, 23-42, Marcel Dekker, Inc. New York 1995.
  2. Bailey L. B., Gregory J. F. Folate metabolism and requirements. J. Nutr., 129: 779-782, 1999.[Abstract/Free Full Text]
  3. Benkovic S. J. The transformylase enzymes in de novo purine biosynthesis. Trends Biochem. Sci., 9: 320-322, 1984.
  4. Allegra C. J. Antifolates: the new millennium. Semin. Oncol., 26: 1-2, 1999.[Medline]
  5. Chu E., Allegra C. J. Antifolates Chabner B. A. Longo D. L. eds. . Cancer Chemotherapy and Biotherapy: Principles and Practice, 109-148, Lippincott-Raven Publishers Philadelphia 1996.
  6. Kaye S. B. New antimetabolites in cancer chemotherapy and their clinical impact. Br. J. Cancer, 78: 1-7, 1998.
  7. Priest D. G., Bunni M. A. Folates and folate antagonists in cancer chemotherapy Bailey L. B. eds. . Folate in Health and Disease, 379-403, Marcel Dekker, Inc. New York 1995.
  8. Kisliuk R. Folate biochemistry in relation to antifolate selectivity Jackman A. L. eds. . Antifolate Drugs in Cancer Therapy, 13-36, Humana Press Inc. Totowa, NJ 1999.
  9. Calvert H. An overview of folate metabolism: features relevant to the action and toxicities of antifolate anticancer agents. Semin. Oncol., 26: 3-10, 1999.[Medline]
  10. Kutzbach C., Stokstad E. L. R. 10-Formyl tetrahydrofolate: NADP oxidoreductase. Methods Enzymol., 18B: 793-798, 1971.
  11. Scrutton M. C., Beis I. Inhibitory effects of histidine and their reversal. The roles of pyruvate carboxylase and N-10-formyltetrahydrofolate dehydrogenase. Biochem. J., 177: 833-846, 1979.[Abstract/Free Full Text]
  12. Krebs H. A., Hems R., Tyler B. The regulation of folate and methionine metabolism. Biochem. J., 158: 341-353, 1976.[Abstract/Free Full Text]
  13. Tephly T. R. The toxicity of methanol. Life Sci., 48: 1031-1041, 1991.[Medline]
  14. Wyngaarden J. B., Kelley W. N. Gout Stanbury J. B. Wyngaarden J. B. Fredrickson D. S. Goldstein J. L. Brown M. S. eds. . The Metabolic Basis of Inherited Disease, 1044-1114, New York, McGraw-Hill, Inc. 1983.
  15. Rudolph F. B., Kulkarni A. D., Fanslow W. C., Pizzini R. P., Kumar S., Van Buren C. T. Role of RNA as a dietary source of pyrimidines and purines in immune function. Nutrition, 6: 45-52, 1990.[Medline]
  16. Freemantle S. J., Moran R. G. Transcription of the human folylpoly-{gamma}-glutamate synthetase gene. J. Biol. Chem., 272: 25373-25379, 1997.[Abstract/Free Full Text]
  17. Peri K. G., MacKenzie R. E. Transcriptional regulation of murine NADP(+)-dependent methylenetetrahydrofolate dehydrogenase-cyclohydrolase-synthetase. FEBS Lett., 294: 113-115, 1991.[Medline]
  18. Girgis S., Nasrallah I. M., Suh J. R., Oppenheim E., Zanetti K. A., Mastri M. G., Stover P. J. Molecular cloning, characterization and alternative splicing of the human cytoplasmic serine hydroxymethyltransferase gene. Gene, 210: 315-324, 1998.[Medline]
  19. Galivan J., Ryan T., Rhee M., Yao R., Chave K. Glutamyl hydrolase: properties and pharmacologic impact. Semin. Oncol., 26: 33-37, 1999.
  20. Gaughan D. J., Barbaux S., Kluijtmans A. J., Whitehead A. S. The human and mouse methylenetetrahydrofolate reductase (MTHFR) genes: genomic organization, mRNA structure and linkage to the CLCN6 gene. Gene, 257: 279-289, 2000.[Medline]
  21. Ross J. F., Chaudhuri P. K., Ratnam M. Differential regulation of folate receptor isoforms in normal and malignant tissues in vivo and in established cell lines. Physiologic and clinical implications. Cancer (Phila.), 73: 2432-2443, 1994.[Medline]
  22. Bucher N. L. R. Liver regeneration then and now Jirtle R. L. eds. . Liver Regeneration and Carcinogenesis, 1-25, Academic Press San Diego, CA 1995.
  23. Michalopoulos G. K., DeFrances M. C. Liver regeneration. Science (Wash. DC), 276: 60-66, 1997.[Abstract/Free Full Text]
  24. Ross M. A., Sander C. M., Kleeb T. B., Watkins C., Stolz D. B. Spatiotemporal expression of angiogenesis growth factor receptors during the revascularization of regenerating rat liver. Hepatology, 34: 1135-1148, 2001.[Medline]
  25. Krupenko S. A., Wagner C., Cook R. J. Cysteine 707 is involved in the dehydrogenase active site of rat 10-formyltetrahydrofolate dehydrogenase. J. Biol. Chem., 270: 519-522, 1995.[Abstract/Free Full Text]
  26. Hentze M. W. Enzymes as RNA-binding proteins: a role for (di)nucleotide binding domains?. Trends Biochem. Sci., 19: 101-103, 1994.[Medline]
  27. Jeffery C. J. Moonlighting proteins. Trends Biochem. Sci., 24: 8-11, 1999.[Medline]
  28. Cole K. A., Krizman D. B., Emmeret-Buck M. R. The genetics of cancer—a 3D model. Nat. Genet., 21: 38-41, 1999.[Medline]
  29. Kallioniemi O.-P., Wagner U., Kononen J., Sauter G. Tissue microarray technology for high-throughput molecular profiling of cancer. Hum. Mol. Genet., 10: 657-662, 2001.[Abstract/Free Full Text]
  30. Bertino J. R. Karnofsky memorial lecture. Ode to methotrexate. J. Clin. Oncol., 11: 5-14, 1993.[Medline]
  31. Barford P. A., Blair J. A. Effect of an implanted walker tumor on metabolism of folic acid in the rat. Br. J. Cancer, 38: 122-129, 1978.[Medline]
  32. Champion K. M., Cook R. J., Tollaksen S. L., Giometti C. S. Identification of a heritable deficiency of the folate-dependent enzyme 10-formyltetrahydrofolate dehydrogenase in mice. Proc. Natl. Acad. Sci. USA, 91: 11338-11342, 1994.[Abstract/Free Full Text]
  33. Weber G. Biochemical strategy of cancer cells and the design of chemotherapy: G. H. A. Clowes Memorial Lecture. Cancer Res., 43: 3466-3492, 1983.[Free Full Text]
  34. Rios-Orlandi E. M., Zarkadas C. G., MacKenzie R. E. Formyltetrahydrofolate dehydrogenase-hydrolase from pig liver: simultaneous assay of the activities. Biochim. Biophys. Acta, 87: 24-35, 1986.
  35. Beardsley G. P., Moroson B. A., Taylor E. C., Moran R. G. A new folate antimetabolite, 5,10-dideaza-5,6,7,8-tetrahydrofolate is a potent inhibitor of the de novo purine synthesis. J. Biol. Chem., 264: 328-333, 1989.[Abstract/Free Full Text]
  36. Baldwin S. W., Tse A., Taylor E. C., Rosowsky A., Shin C., Moran R. G. Structural features of 5,10-dideaza-5,6,7,8-tetrahydrofolate that determine inhibition of mammalian glycinamide ribonucleotide formyltransferase. Biochemistry, 30: 1997-2006, 1991.[Medline]
  37. Pizzorno G., Moroson B. A., Cashmore A. R., Beardsley G. P. (6R)-5,10-Dideaza-5,6,7,8-tetrahydrofolic acid effects on nucleotide metabolism in CCRF-CEM human T-lymphoblast leukemia cells. Cancer Res., 51: 2291-2295, 1991.[Abstract/Free Full Text]
  38. Kinsella A. R., Smith D., Pickard M. Resistance to chemotherapeutic antimetabolites: a function of salvage pathway involvement and cellular response to DNA damage. Br. J. Cancer, 75: 935-945, 1997.[Medline]
  39. Rhee M. S., Coward J. K., Galivan J. Depletion of 5, 10-methylenetetrahydro-folate and 10-formyltetrahydrofolate by methotrexate in cultured hepatoma cells. Mol. Pharmacol., 42: 909-916, 1992.[Abstract]
  40. Priest D. G., Schmitz J. C., Bunni M. A. Folate metabolites as modulators of anti-tumor drug activity. Adv. Exp. Biol. Med., 338: 693-698, 1993.[Medline]
  41. Savaiano D. A., Ho C. Y., Chu V., Clifford A. J. Metabolism of orally and intravenously administered purines in rats. J. Nutr., 110: 1793-1804, 1980.
  42. Savaiano D. A., Clifford A. J. Adenine, the precursor of nucleic acids in intestinal cells unable to synthesize purines de novo. J. Nutr., 111: 1816-1822, 1981.
  43. Wilkinson D. G. eds. . In Situ Hybridization: A Practical Approach, 1-158, Oxford University Press Oxford, England 1998.
  44. Krupenko S. A., Wagner C., Cook R. J. Expression, purification, and properties of the aldehyde dehydrogenase homologous carboxyl-terminal domain of rat 10-formyltetrahydrofolate dehydrogenase. J. Biol. Chem., 272: 10266-10272, 1997.[Abstract/Free Full Text]
  45. Lin H., Roberts E. S., Hollenberg P. F. Heterologous expression of rat P450 2E1 in a mammalian cell line: in situ metabolism and cytotoxicity of N-nitrosodimethyl-amine. Carcinogenesis (Lond.), 19: 321-329, 1998.[Abstract/Free Full Text]



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