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Originally published In Press as doi:10.1074/jbc.M908096199 on April 11, 2000

J. Biol. Chem., Vol. 275, Issue 24, 18489-18494, June 16, 2000
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Cellular Release of [18F]2-Fluoro-2-deoxyglucose as a Function of the Glucose-6-phosphatase Enzyme System*

Corradina CaracóDagger , Luigi Aloj§, Li-Yuan Chen||, Janice Y. Chou||, and William C. Eckelman**

From the Positron Emission Tomography and § Nuclear Medicine Departments, Clinical Center, || Heritable Disorders Branch, NICHD, National Institutes of Health, Bethesda, Maryland 20892

Received for publication, October 6, 1999, and in revised form, April 5, 2000

    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

[18F]-2-Fluoro-2-deoxyglucose (FDG) is a glucose analog currently utilized for positron emission tomography imaging studies in humans. FDG taken up by the liver is rapidly released. This property is attributed to elevated glucose-6-phosphatase (Glc-6-Pase) activity. To characterize this issue we studied the relationship between Glc-6-Pase activity and FDG release kinetics in a cell culture system. We overexpressed the Glc-6-Pase catalytic unit in a Glc-6-Pase-deficient mouse hepatocyte (Ho-15) and in A431 tumor cell lines. Glc-6-Pase enzyme activity and FDG release rates were determined in cells transfected with the Glc-6-Pase gene (Ho-15-D3 and A431-AC3), in mock-transfected cells of both cell lines, and in wild-type mouse hepatocytes (WT10) as control. Although the highest level of Glc-6-Pase activity was measured in A431-AC3, Ho-15-D3 cells showed much faster FDG release rates. The faster FDG release correlated with the level of glucose 6-phosphate transporter (Glc-6-PT) mRNA, which was found to be expressed at higher levels in Ho-15 compared with A431 cells. Overexpression of Glc-6-PT in A431-AC3 produced a dramatic increase in FDG release compared with control cells. This study gives the first direct evidence that activity of the Glc-6-Pase complex can be quantified in vivo by measuring FDG release. Adequate levels of Glc-6-Pase catalytic unit and Glc-6-PT are required for this function. FDG-positron emission tomography may be utilized to evaluate functional status of the Glc-6-Pase complex.

    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Glucose-6-phosphatase (Glc-6-Pase),1 most abundantly found in liver and kidney, catalyzes the last step of both gluconeogenesis and glycogenolysis. This enzyme is a key protein in the regulation of glucose homeostasis. Its function is to dephosphorylate glucose 6-phosphate (Glc-6-P), so that free glucose can be transported out of cells and released into the blood. Thus, the activity of this enzyme controls liver glucose production. Defects of the Glc-6-Pase enzyme system cause glycogen storage disease type 1 (GSD-1), which manifests with severe hypoglycemia and hepatomegaly caused by accumulation of glycogen. The Glc-6-Pase enzyme system is associated with the endoplasmic reticulum and has multiple components (1). Current understanding of this system postulates that Glc-6-P is transported into the microsome by a Glc-6-P translocase (Glc-6-PT) (2). Once inside the microsome, Glc-6-P is dephosphorylated by the Glc-6-Pase catalytic unit, and finally phosphate and glucose are transported out of the microsome via specific transport proteins (T2 and T3, respectively). The cDNA and gene for Glc-6-PT have been recently described (3, 4). Full characterization of this system is still pending.

[18F]-2-Fluoro-2-deoxyglucose (FDG) is a glucose analog currently utilized for positron emission tomography (PET) imaging studies in humans. FDG uptake has been used for many years to measure in vivo regional glucose utilization (5). This tracer competes with glucose for phosphorylation by hexokinase or glucokinase. After it is phosphorylated, FDG-6-phosphate (FDGlc-6-P) does not undergo glycolysis. In tissues with elevated glucose metabolic rates, such as tumors, with low or absent dephosphorylating activity, FDGlc-6-P is trapped inside cells. This property allows for imaging of areas with increased FDG retention and has been extensively applied to visualize, stage, and monitor progression of tumors (6). On the other hand, in organs such as the liver, FDG is taken up and rapidly released, presumably for the presence and activity of the Glc-6-Pase enzyme (7). Preliminary data indicate that in patients with GSD-1, FDG is more avidly retained in the liver compared with normal subjects (8). The different steps in FDG metabolism, developed from the seminal work by Sokoloff and co-workers (9, 10) using [14C]deoxyglucose, can be described by the two tissue compartment model. Kinetic analysis of dynamic PET-FDG studies allows estimation of the rate constants for each step (Fig. 1). The current understanding of FDG metabolism is that the k4 parameter is tightly linked to the presence and activity of the Glc-6-Pase enzyme system. Therefore, in vivo measurement of k4 for FDG may provide a non-invasive quantitative method to evaluate liver Glc-6-Pase function and also be a tool to monitor the effectiveness of gene therapy in patients with GSD type 1. 


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  		Fig. 1.   Schematic representation of the two tissue compartment model for FDG. Transport of FDG into and out of the cell is mediated by glucose transport proteins and is represented, respectively, by the K1 and k2 rate constants. FDG inside the cell is transformed to FDG-6-P via the action of glucokinase; this step is represented by the k3 rate constant. Dephosphorylation of FDG is represented by the k4 rate constant; this step is presumed to be mediated by the action of Glc-6-Pase and/or other phosphatases present intracellularly. Changes of intracellular Glc-6-Pase activity should alter measurements of this parameter in the liver in vivo.

To characterize further this issue, we have analyzed the FDG turnover properties of cultured cells and verified if correlations exist between Glc-6-Pase activity and whole cell FDG release kinetics. We have studied three different cell lines as follows: A431, derived from a human epidermoid carcinoma; Ho-15 cells, derived from the liver of a Glc-6-Pase-deficient mouse (11); and a hepatocyte cell line derived from a normal mouse as control (WT10). In the first two cell lines we isolated clones that overexpress the Glc-6-Pase catalytic unit by transfection. Furthermore, A431 cells were also infected with a recombinant adenovirus containing the human Glc-6-PT cDNA (Ad-hGlc-6-PT-5'FLAG) to overexpress the Glc-6-PT.

    EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Cell Culture-- The Glc-6-Pase-deficient mouse hepatocyte cell line (Ho-15) and normal mouse hepatocyte cell line (WT10) were established by transformation of primary hepatocytes from a Glc-6-Pase-deficient mouse and a normal mouse, respectively, by the SV40 temperature-sensitive A255 virus (12). Both cell lines were cultured in alpha -minimum Eagle's medium supplemented with 4% fetal bovine serum and 100 nM dexamethasone at 34 °C (permissive temperature). In all experiments herein described, cells were placed in an incubator at 37 °C the night before use.

A431 cells were cultured in Dulbecco's modified Eagle's medium supplemented with 10% fetal bovine serum, at 37 °C (13).

Establishment of Glc-6-Pase Expression in Cell Lines-- A431 cells were permanently transfected with the pCR3.1 (Invitrogen, San Diego, CA) plasmid alone or containing the full coding sequence (nucleotides 65-1165, GenBankTM accession number U01120) for the human liver Glc-6-Pase catalytic unit. Transfected clones were established by selection in medium containing the neomycin analog G418 (Life Technologies, Inc., 75 µg/ml).

Ho-15 were permanently transfected with the pcDNA3.1/Zeo plasmid (Invitrogen) containing the full coding sequence (nucleotides 46-1425, Gen Bank GenBankTM accession number U00445) for the mouse liver Glc-6-Pase catalytic unit. The need for a different construct was dictated by the fact that cells were derived from a knockout mouse and thus were constitutively neomycin-resistant. A parallel transfection with pcDNA3.1/Zeo plasmid alone was also performed. Transfected clones were established by selection in medium containing zeocin (Invitrogen, 100 µg/ml).

Verification of Glc-6-Pase mRNA Expression-- Expression of the transfected gene in both cell lines was verified by Northern blot analysis. Fifteen µg of total RNA were denatured by heating at 65 °C for 15 min and loaded onto a 1% agarose formaldehyde gel containing ethidium bromide (14). The samples were electrophoresed for 2 h at 100 V. The ethidium signal was captured using a Molecular Dynamics 595 Fluorimager (Sunnyvale, CA), and the gel contents were subsequently transferred to nylon membranes using a Turboblotter apparatus (Schleicher & Schuell). After UV cross-linking, the membranes were hybridized with the following 32P-labeled probes. For human beta -actin an oligonucleotide complementary to bases 1101-1148 of the cDNA (GenBankTM accession number X00351) was used. For human Glc-6-Pase an oligonucleotide complementary to bases 1121-1165 of the cDNA (GenBankTM accession number U01120) was used. For mouse Glc-6-Pase an oligonucleotide complementary to bases 1373-1417 of the cDNA (GenBankTM accession number U00445) was used. All oligonucleotide probes were labeled at the 5'-end. For the detection of mouse beta -actin, the human cDNA plasmid purchased from the American Type Culture Collection (Manassas, VA) was used after labeling by random priming. After hybridization the membranes were washed twice in 2× SSC with 0.1% SDS at 70 °C, exposed overnight on imaging plates, and analyzed using a FUJI BAS-1500 PhosphorImager (Stamford, CT). All probes were tested on commercially available Northern blots (CLONTECH, Palo Alto, CA) prior to use in the experiments reported.

Alternatively, the presence of mouse Glc-6-Pase mRNA was detected by reverse transcriptase PCR, using standard procedures. Reverse transcription was performed with random hexamers as primers, and the PCR step was performed using 22-base oligonucleotides designed to amplify a 541-bp portion of the cDNA corresponding to bases 66-606 of the mouse Glc-6-Pase gene (GenBankTM accession number U00445).

The presence of mRNA for the Glc-6-PT protein in the A431 and Ho-15 cells was assessed by Northern blot. Total RNA was isolated by the guanidinium thiocyanate/CsCl method (15), and poly(A)+ RNA was obtained by oligo(dT)-cellulose chromatography. RNA was fractionated by electrophoresis through 1.2% agarose gels containing 2.2 M formaldehyde and transferred to a Nytran membrane by electroblotting. The filters were hybridized at 62 °C in a buffer containing 5× SSC, 50% formamide, 50 mM sodium phosphate buffer, pH 6.5, 8× Denhardt, 1% SDS, 200 µg/ml sonicated salmon sperm DNA, and a uniformly labeled mouse Glc-6-Pase, Glc-6-PT, or beta -actin riboprobe.

Construction of Ad-hGlc-6-PT-5'FLAG-- The hGlc-6-PT-5'FLAG construct containing the eight-amino acid FLAG marker peptide, DYKDDDDK (Scientific Imaging Systems, Eastman Kodak Co.), at the N terminus of human Glc-6-PT has been described (4). A recombinant adenovirus containing the hGlc-6-PT-5'FLAG (Ad-hGlc-6-PT-5'FLAG) carrying the viral E1-deleted mutation was generated through homologous recombination in 293 cells between a co-transfected linearized transfer vector (pAv6-hGlc-6-PT-5'FLAG) and the large ClaI fragment of wild-type adenoviral Ad5 as described previously (16, 17). pAv6-hGlc-6-PT-5'FLAG transfer vector contained the entire coding region (nucleotides 166-1486) of the human Glc-6-PT cDNA under the control of the RSV promoter.

Overexpression of Glc-6-PT-- Overexpression of Glc-6-PT was achieved by infecting A431 cells or A431-AC3 cells for 24 h at 37 °C with lysates containing Ad-hGlc-6-PT-5'FLAG obtained from Ad-hGlc-6-PT-5'FLAG-infected 293 cells. The amounts of virus used were determined in preliminary experiments by testing Glc-6-PT protein production by Western blot analysis using a monoclonal antibody against the FLAG epitope (Scientific Imaging System) as described previously (4). The immunocomplex was detected with the horseradish peroxidase-linked chemiluminescent system using the SuperSignal West Pico Chemiluminescent Substrate obtained from Pierce. Northern blot analysis on total RNA was also performed to compare hGlc-6-PT mRNA expression before and after infection.

Enzyme Activity Assay-- Glc-6-Pase enzyme assays were performed by measuring the ability of fully disrupted microsomes to hydrolyze [3H]2-deoxyglucose 6-phosphate ([3H]DGlc-6-P). Mouse liver microsomes were prepared as described previously from fresh tissue, disrupted, and stored frozen at -70 °C at a concentration of 10 mg of protein/ml (18). Microsomes from cultured cells were isolated with the same centrifugation procedure, disrupted, and resuspended in buffer containing 100 mM KCl, 5 mM MgCl2, 20 mM NaCl, 20 mM MOPS, at pH 7.2. [3H]DGlc-6-P was synthesized incubating [3H]DG (1 mCi, 60 Ci/mmol, American Radiolabeled Chemicals, St. Louis, MO) with 10 units/ml agarose-immobilized hexokinase in 40 mM potassium phosphate buffer at pH 7.6 containing 40 mM KCl, 5 mM ATP, and 5 mM MgCl2 for 1 h at 33 °C in a shaking water bath. The hexokinase beads were separated by centrifugation, and the supernatant was collected and applied to AG 1X-8 resin (Bio-Rad). The columns were washed with water to remove unbound [3H]DG, and then [3H]DGlc-6-P was eluted with 0.5 M NaCl. The Glc-6-Pase assays were carried out at 33 °C in a total volume of 250 µl. Each sample contained 0.1 µCi of [3H]DGlc-6-P and microsomal protein at a concentration of 0.5 mg/ml. In the control samples no microsomal extract was added to determine background dephosphorylation. Between 5 and 60 min after addition of the radioactivity, aliquots of the mixture were collected and the reaction terminated by adding to 0.5 ml of 0.3 M ZnSO4. After mixing, 0.5 ml of a saturated solution of Ba(OH)2 were added, and after vortexing the tubes were centrifuged for 2 min at 14,000 rpm in an Eppendorf centrifuge. Aliquots of the supernatant were counted for each sample, and relative amounts of [3H]DG formed were determined (19). The rate of [3H]DGlc-6-P dephosphorylation is expressed as the background subtracted percentage of the initial [3H]DGlc-6-P present transformed per min.

Measurement of [18F]FDG Release Rate-- [18F]FDG was prepared in the NIH Cyclotron facility by standard means (20). Cells were plated in 12-well multiwell plates at a density of 2.5 × 105/well and allowed to grow for 3 days. On the day of the experiment, cells were incubated in Dulbecco's modified Eagle's medium without glucose supplemented with 0.5% fetal bovine serum, containing [18F]FDG (~1 × 106 cpm/well) for 60 min. After 60 min one plate for each clone was washed twice with fresh medium and incubated for different times in cold medium containing 1 g/liter glucose (5.5 mM). At each time point triplicate wells were rinsed twice with cold phosphate-buffered saline, and cell-associated counts were recovered by lysing the cells in 1 M NaOH. Radioactivity in cells and medium was then determined for each sample. Protein concentrations in the cell extracts were monitored throughout the experiment to verify that cells were not detaching from the plate during the washes. Cell-associated radioactivity was plotted as the percent of the activity present at time 0 (end of uptake period). In separate experiments, cells were solubilized after extensive washing in 0.1 M NaOH at different times following the 60-min incubation in uptake medium. The fraction of intracellular FDGlc-6-P in these samples was determined with the ZnSO4 and Ba(OH)2 method described above.

    RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Transfection of A431 cells with the plasmid containing the human Glc-6-Pase catalytic unit gene yielded numerous G418-resistant clones. In 13 clones Glc-6-Pase mRNA expression was determined. The positive clone with the highest level of Glc-6-Pase mRNA expression (A431-AC3) and one mock-transfected clone were selected for further characterization (Fig. 2). RT-PCR analysis of A431 wild-type (A431-WT) and A431-mock cells for the presence of human Glc-6-Pase mRNA was negative. The presence of mRNA for mouse Glc-6-Pase in WT10, used as control, was confirmed by RT-PCR (not shown).


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  		Fig. 2.   Northern blot analysis of A431 cells and transfected clones (A431-mock and A431-AC3) for human Glc-6-Pase mRNA expression. A431-WT and mock-transfected cells show absent expression of Glc-6-Pase mRNA. Results were confirmed by RT-PCR.

Glc-6-Pase enzyme activity was determined on fully disrupted microsomes prepared from A431-WT, A431-AC3, and A431-mock. Mouse liver tissue and WT10 hepatocytes were also measured as control (Fig. 3). Values for A431-AC3 were 2.5-fold higher compared with mouse liver (1.08 ± 0.23%/min, mean ± S.D. versus 0.44 ± 0.05, respectively). Dephosphorylating activities in A431-WT and A431-mock were comparable to those found in background (control) tubes. Glc-6-Pase activity in WT10 was 5 times lower than in mouse liver.


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  		Fig. 3.   [3H]DGlc-6-P dephosphorylating activity. Comparison between fully disrupted microsomes obtained from mouse liver, A431 wild-type and transfected clones (A431-mock and A431-AC3), and normal mouse hepatocytes (WT10) is shown. A431-AC3 microsomes show the highest level of activity. In A431-WT and A431-mock only background levels were detected. Error bars = S.D.; n = 3.

The FDG release kinetics for the A431 cells are shown in Fig. 4 in comparison with WT10. Both A431-AC3 and A431-mock show slower release rates than the hepatocytes. The A431-AC3 showed only a slight increase in release rate, compared with the A431-mock clone (half-times were 230 and 300 min, respectively). Release rates in WT10 were much more rapid than in both these cells (half-time = 119 min).


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  		Fig. 4.   FDG release. Comparison of A431-mock () and A431-AC3 (open circle ) with normal mouse hepatocytes (×) is shown. Normal mouse hepatocytes show much more rapid release than both A431 clones. A431-AC3 shows only slightly (<30%) faster release rates than A431-mock. Error bars = S.D.; n = 3 per time point.

Transfection efficiency of the Ho-15 cell line with the mouse Glc-6-Pase-containing plasmid was very low. We obtained 8 zeocin-resistant clones; however, only 3 of these clones could be further characterized due to the poor growth of the remaining clones. Out of these 3, a single clone (Ho-15-D3) was found to express the mouse Glc-6-Pase mRNA. Transfection of Ho-15 with the mock plasmid yielded similar efficiency. A mock-transfected clone and the Ho-15-D3 underwent further analysis. Mouse Glc-6-Pase mRNA expression of these cell lines and of RNA extracted from mouse liver is shown in Fig. 5. The level of mRNA expression in Ho-15-D3 cells is similar to that of mouse liver. Mouse Glc-6-Pase mRNA was not detectable in Ho-15-WT and mock-transfected cells by RT-PCR as well.


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  		Fig. 5.   Northern blot analysis of mouse liver, Ho-15 wild-type (Ho-15-WT) cells, and transfected clones (Ho-15-mock and Ho-15-D3) for mouse Glc-6-Pase mRNA expression. Ho-15-WT and mock-transfected cells show no detectable expression of Glc-6-Pase mRNA. Results were confirmed by RT-PCR.

Microsomal Glc-6-Pase enzyme activity was approximately 4 times lower in Ho-15-D3 cells compared with mouse liver (Fig. 6). Compared with the Ho-15-WT and mock-transfected cells, however, values obtained in Ho-15-D3 cells were 3-4 times higher. Activity in Ho-15-D3 cells was very similar to that found for WT10 (see Fig. 3).


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  		Fig. 6.   [3H]DGlc-6-P dephosphorylating activity. Comparison between fully disrupted microsomes obtained from mouse liver, Ho-15 wild-type (Ho-15-WT) cells, and transfected clones (Ho-15-mock and Ho-15-D3) is shown. Ho-15-D3 microsomes show higher activity than Ho-15-WT and mock clones. Error bars = S.D.; n = 3.

FDG release in Ho-15-D3 is much more rapid compared with the mock-transfected clone (Fig. 7). Half-times were 120 and 361 min, respectively. In a separate experiment, a zeocin-resistant clone obtained by transfection with the mouse Glc-6-Pase containing plasmid (Ho-15-D1), in which Glc-6-Pase mRNA expression, however, was negative by Northern blot, showed release rates similar to those of the Ho-15-mock cells (not shown).


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  		Fig. 7.   FDG release. Comparison of Ho-15-mock (black-square) and Ho-15-D3 cells () is shown. Addition of Glc-6-Pase catalytic unit in Ho-15 cells causes a 3-fold increase in FDG release rate. Values for Ho-15-D3 are similar to those obtained in WT10 (see Fig. 4). Error bars = S.D.; n = 3 per time point.

The chemical form of intracellular FDG was determined in all cell lines tested. At the end of the uptake period, 95 ± 0.1% (mean ± S.D.) of intracellular radioactivity is in the form of FDGlc-6-P in both A431-mock and A431-AC3 cells. In Ho-15-mock and Ho-15-D3, these values were 97.4 ± 0.2 and 97.1 ± 0.2%, respectively. These ratios do not change with time, and 120 min after the uptake period, virtually all intracellular radioactivity is in the form of FDGlc-6-P in Ho-15-mock and Ho-15-D3 cells (99.5 ± 0.1% in both cell lines).

A431 and Ho-15 cells, along with the respective transfected clones, were also tested for the expression of Glc-6-PT mRNA by Northern blot (Fig. 8). Expression in Ho-15 cells and Ho-15-D3 was found to be much higher than in A431 and A431-AC3 cells.


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  		Fig. 8.   Northern blot analysis of A431 wild-type (A431-WT) and Ho-15 wild-type (Ho-15-WT) cells and respective transfected clones (A431-AC3 and Ho-15-D3) for Glc-6-Pase catalytic unit and Glc-6-PT mRNA expression. Liver-derived cells show higher level of expression of Glc-6-PT compared with A431.

To obtain higher levels of Glc-6-PT expression, A431-WT and A431-AC3 cells were infected with Ad-hGlc-6-PT-5'FLAG. Protein expression was confirmed by Western blot analysis for the presence of the FLAG epitope (Fig. 9A). The levels of expression of Glc-6-PT mRNA (Fig. 9B) were considerably higher in infected compared with untreated cells. In A431-AC3 cells overexpressing Glc-6-PT, a dramatic increase in FDG release was observed compared with untreated A431-AC3 cells (Fig. 10A). The increase in FDG release rate was directly proportional to the amount of Ad-hGlc-6-PT-5'FLAG used for infection. There was no effect of Ad-hGlc-6-PT-5'FLAG infection on the FDG release of A431-WT cells (Fig. 10B).


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  		Fig. 9.   A, Western blot analysis of A431 wild-type (A431-WT) and A431-AC3 cells 24 h after Ad-hGlc-6-PT-5'FLAG infection. Untreated cells were used as control. Decreasing titers of Ad-hGlc-6-PT-5'FLAG (from 1 to 0.01×) were used to infect A431-AC3 cells. The membrane was probed with a monoclonal antibody against the FLAG epitope. Bottom panel, Amido Black staining of the membrane for total protein. B, Northern blot analysis of A431-WT and A431-AC3 before and 24 h after treatment with Ad-hGlc-6-PT-5'FLAG (1×). Markedly higher levels of Glc-6-PT mRNA are seen in both cell types after infection. Bottom panel, ethidium bromide stain of the gel showing 28 S ribosomal RNA band.


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  		Fig. 10.   A, FDG release in A431-AC3 cells 24 h after addition of Ad-hGlc-6-PT-5'FLAG at 1 (), 0.1 (×), and 0.01× (triangle ) dilution, compared with untreated cells (open circle ) (same titers as in Fig. 9). B, FDG release in untreated A431-WT cells () compared with Ad-hGlc-6-PT-5'FLAG-treated A431-WT cells at 1× dilution (black-square). Notice the shorter time scale compared with previous figures. Error bars = S.D.; n = 3 per time point.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

The recent cloning of the genes that encode for the human (21) and mouse (22) Glc-6-Pase catalytic unit and Glc-6-PT (3, 23) gives us the opportunity to better understand the effect of this enzyme system on FDG efflux kinetics. In the present study we have investigated the relationship between the presence of these factors and FDG release in whole cells.

In our first approach to understanding the role of these proteins, we have analyzed the effect of Glc-6-Pase catalytic unit expression on FDG release from A431 cells. These cells are derived from a human epidermoid carcinoma. Cancer cells, in general, are believed to have low or absent Glc-6-Pase activity, which was confirmed in A431-WT cells by mRNA analysis (Fig. 2). Enzymatic activity in these cells, as assessed by the dephosphorylation assay, was similarly found to be very low compared with that measured in the mouse liver microsome preparation where Glc-6-Pase activity is known to be present. Overexpression by transfection of the human Glc-6-Pase mRNA in these cells was successful, and very high levels of expression were obtained in the A431-AC3 clone as proven both by the Northern blot experiment (Fig. 2) and the enzymatic activity assay (Fig. 3). However, release kinetics of FDG in these cells were not influenced by the presence of the Glc-6-Pase catalytic unit (Fig. 4). It is surprising to note that FDG release in A431-AC3 cells is much slower than in the WT10, even though A431-AC3 extracts show much higher dephosphorylating activity than the ones obtained from the hepatocytes. Furthermore, FDG release in A431-AC3 cells is only 30% faster than that found in the mock-transfected A431, indicating very little effect, if any, of the transfected protein on the FDG release process in these cells. These data are in agreement with the notion that for the Glc-6-Pase system to function it requires more than the catalytic unit alone (24).

It is reasonable to believe that the other components of the Glc-6-Pase enzyme system are involved in Glc-6-Pase-dependent FDG release. These components must be present in liver-derived cells. Therefore, our next approach was to study how this protein affects FDG release in hepatocytes derived from a normal mouse and in hepatocytes derived from a Glc-6-Pase-deficient mouse (Ho-15) (11). Comparison of the behavior of these two cell lines constitutes an optimal model for studying the effect of the Glc-6-Pase protein on FDG release. Glc-6-Pase activity is present in WT10, and it is present at higher levels compared with Ho-15 cells. In WT10, however, Glc-6-Pase activity is lower than in microsomes extracted directly from mouse liver. Glc-6-Pase activity can be reestablished in the Ho-15 cells by transfection, and the levels of Glc-6-Pase activity achieved in the Ho-15-D3 clone are similar to those found in WT10 (Figs. 3 and 6). FDG release rates of the mock-transfected Ho-15 cells are ~3 times slower than in the Ho-15-D3 cells (Fig. 7). Furthermore, the release rates of the Ho-15-D3 cells are nearly identical to those measured for the WT10, indicating unequivocally the role of the Glc-6-Pase catalytic unit in promoting FDG release of these two liver-derived cell lines.

Intracellular FDG was found to be virtually all in the form of FDGlc-6-P, in each of the cell lines tested. This suggests that the residence time of FDG intracellularly is very short and that the molecule is rapidly transported out of the cell if it is not in phosphorylated form, regardless of which cell line is being analyzed. These data support the relationship between the ability to dephosphorylate in Ho-15-D3 cells and the increased release rates of FDG, indicating that dephosphorylation is indeed controlling the release of radioactivity. Similar evidence is also provided by PET studies in humans (25). The k2 rate constant for FDG in the liver has been found to be in the order of 1 min-1 (half-time of less than 1 min). Therefore, in vivo, the intracellular residence time of free FDG is very short as well. The overall release process is under the control of the k4 parameter which is slower (0.014 min-1, half-time of approximately 50 min) indicating that the rate at which FDGlc-6-P is being transformed into FDG is controlling the release of radioactivity.

The high level of expression of Glc-6-Pase catalytic unit in A431-AC3 cells and relatively unchanged FDG release rates compared with A431-WT cells may be explained by the fact that FDGlc-6-P does not come in contact with the catalytic unit at all. Other evidence is provided by the fact that intracellular radioactivity remains in phosphorylated form even though Glc-6-Pase activity is very high in the cellular extracts. In experimental studies in COS-1 cells, Glc-6-P transport into microsomes was found to be increased by simultaneous transfection of Glc-6-Pase catalytic unit and Glc-6-PT, rather than the overexpression of each protein individually, suggesting that adequate levels of both components of the system are required (26). The low levels of Glc-6-PT RNA expression in A431 cells, compared with Ho-15 hepatocytes (Fig. 8), suggest that inadequate amounts of Glc-6-PT protein produced in A431-AC3 cells may be the rate-limiting factor in FDG release.

To test this hypothesis, the protein was overexpressed in A431-WT and A431-AC3 cells (Fig. 9). Our results show that FDG release in A431-AC3 cells, overexpressing Glc-6-Pase catalytic unit and Glc-6-PT, occurs at higher rates than in A431-WT cells, A431-WT cells overexpressing the Glc-6-PT alone, or A431-AC3 cells overexpressing the catalytic unit alone. These results show that simultaneous overexpression of Glc-6-Pase catalytic unit and Glc-6-PT in A431 cells is necessary to increase FDG release to levels similar to those measured in hepatocytes.

Finally, our data show that FDG is released from all cells we tested at measurable rates even in the total absence of the Glc-6-Pase catalytic unit. In A431 cells, FDG is released with a half-time of 300 to 230 min (k = 0.002-0.003 min-1). In PET studies in humans, k4 values for tissues with low Glc-6-Pase activity such as gray matter of the brain have been reported to be 0.0055 min-1 (half-time of approximately 126 min) (27). Dephosphorylation of FDG both in vitro and in vivo, therefore, appears to be mediated at least in part by the action of Glc-6-Pase-independent mechanisms of FDGlc-6-P dephosphorylation.

We have shown, for the first time, a direct dependence between FDGlc-6-P dephosphorylation and FDG release in cultured cells. Increasing Glc-6-Pase catalytic unit activity in A431 cells to higher levels than in mouse liver does not influence FDG release kinetics, whereas increasing activity in Ho-15 cells results in a dramatic increase in FDG release. When Glc-6-PT is expressed at higher levels in A431-AC3 cells, release of FDG is also increased. These data are in agreement with evidence that adequate levels of Glc-6-PT are necessary to preserve Glc-6-Pase-dependent hydrolysis and subsequent release of glucose-, DG-, or FDG-6P. These results support the use of dynamic PET-FDG liver imaging to study in vivo functional status of the entire Glc-6-Pase system. This non-invasive method may be of clinical value to monitor the effectiveness of gene therapy in patients with GSD-1.

    ACKNOWLEDGEMENTS

We thank Dr. Domenico Accili and Dr. Kristina Rother (NICHD, National Institutes of Health) for their help in setting up the experiments with the normal hepatocytes.

    FOOTNOTES

* 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.

Dagger Current address: Istituto di Medicina Sperimentale e Biotecnologie, Consiglio Nazionale delle Ricerche, Localitá Burga, Piano Lago Mangone, 87050, Cosenza, Italy.

Current address: Centro di Studio per la Medicina Nucleare, Consiglio Nazionale delle Ricerche, c/o Dept. di Scienze Biomorfologiche e Funzionali, Universitá degli Studi di Napoli "Federico II", Via S. Pansini, 5, 80131, Napoli, Italy.

** To whom correspondence and reprint requests should be addressed: Positron Emission Tomography Dept., Warren G. Magnuson Clinical Center, Bldg. 10, Rm. 1C495, 10 Center Dr., MSC 1180, National Institutes of Health, Bethesda, MD 20892-1180. Tel.: 301-496-6455; Fax: 301-402-3521; E-mail: eckelman@nih.gov.

Published, JBC Papers in Press, April 11, 2000, DOI 10.1074/jbc.M908096199

    ABBREVIATIONS

The abbreviations used are: Glc-6-Pase, glucose-6-phosphatase; Glc-6-P, glucose 6-phosphate; GSD-1, glycogen storage disease type 1; Glc-6-PT, glucose 6-phosphate transporter; PET, positron emission tomography; FDG, [18F]-2-fluoro-2-deoxyglucose; WT, wild-type, Ad-hGlc-6-PT-5'FLAG, human Glc-6-PT gene-bearing recombinant adenovirus; DGlc-6-P, 2-deoxyglucose 6-phosphate; DG, 2-deoxyglucose; FDGlc-6-P, FDG-6-phosphate; RT-PCR, reverse transcriptase-polymerase chain reaction; MOPS, 4-morpholinepropanesulfonic acid.

    REFERENCES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

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