Esterification of all-trans-retinol in normal human epithelial cell strains and carcinoma lines from oral cavity, skin and breast: reduced expression of lecithin:retinol acyltransferase in carcinoma lines

Xiaojia Guo1, Alberto Ruiz2,3, Robert R. Rando4, Dean Bok2,3 and Lorraine J. Gudas1

1 Department of Pharmacology, Weill Medical College of Cornell University, 1300 York Avenue, New York, NY 10021,
2 Department of Neurobiology, Brain Research Institute and
3 Jules Stein Eye Institute, School of Medicine, University of California, Los Angeles, CA 90095 and
4 Department of Biological Chemistry and Molecular Pharmacology, Harvard Medical School, Boston, MA 02115, USA


    Abstract
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 Abstract
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 Materials and methods
 Results
 Discussion
 References
 
When exogenous [3H]retinol (vitamin A) was added to culture medium, normal human epithelial cells from the oral cavity, skin, lung and breast took up and esterified essentially all of the [3H]retinol within a few hours. As shown by [3H]retinol pulse–chase experiments, normal epithelial cells then slowly hydrolyzed the [3H]retinyl esters to [3H]retinol, some of which was then oxidized to [3H]retinoic acid (RA) over a period of several days. In contrast, cultured normal human fibroblasts and human umbilical vein endothelial cells (HUVEC) did not esterify significant amounts of [3H]retinol; this lack of [3H]retinol esterification was correlated with a lack of expression of lecithin:retinol acyltransferase (LRAT) transcripts in normal fibroblast and HUVEC strains. These results indicate that normal, differentiated cell types differ in their ability to esterify retinol. Human carcinoma cells (neoplastically transformed epithelial cells) of the oral cavity, skin and breast did not esterify much [3H]retinol and showed greatly reduced LRAT expression. Transcripts of the neutral, bile salt-independent retinyl ester hydrolase and the bile salt-dependent retinyl ester hydrolase were undetectable in all of the normal cell types, including the epithelial cells. These experiments suggest that retinoid-deficiency in the tumor cells could develop because of the lack of retinyl esters, a storage form of retinol.

Abbreviations: AHD-2, aldehyde dehydrogenase-2; ARAT, acyl CoA:retinol acyltransferase; DMEM, Dulbecco's modified Eagle's medium; HMEC, normal human mammary epithelial cells; NHBE, normal human bronchial epithelial cells; HPLC, high performance liquid chromatography; HUVEC, human umbilical vein endothelial cells; LRAT, lecithin:retinol acyltransferase; NHEK, normal human epidermal keratinocytes; PBS, phosphate-buffered saline; RA, all-trans-retinoic acid; RALDH-2, retinaldehyde dehydrogenase-2; REH, retinyl ester hydrolase; ROH, all-trans-retinol; SCC, squamous cell carcinoma.


    Introduction
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Retinoids (vitamin A and its metabolites) can act as chemopreventive and/or chemotherapeutic agents for several types of cancer (14). They have major effects on the growth and differentiation of normal, premalignant and malignant epithelial cells both in vitro and in vivo (5). Retinol can be metabolized to retinyl esters and to various structurally related compounds, such as retinoic acid (RA), retinaldehyde, 4-oxoretinol, 14-hydroxy-4–14-retroretinol (14-HRR) and anhydroretinol in many cell types (611). While RA in particular has been demonstrated by many researchers to be useful in the prevention and treatment of cancer in humans (12,13), more recently other retinoids such as anhydroretinol have been shown to prevent cancer in animal models (14).

Retinyl esters are the major metabolites of retinol in some normal cells and tissues, but other cell types are thought not to be able to esterify retinol. For example, human keratinocytes (1520), human intestinal Caco-2 cells (21), cultured tracheal epithelial cells (22), retinal pigment epithelial cells (2325), liver (2630) and mammary epithelial cells (3133) exhibit a high level of retinol esterification activity. Two enzyme activities can catalyze retinyl ester synthesis: acyl CoA:retinol acyltransferase (ARAT) and lecithin:retinol acyltransferase (LRAT) both esterify retinol, but these two enzyme activities differ in their substrate preferences and in their sensitivities to various inhibitors (26,27,34). LRAT uses the acyl group at the sn1 position of membrane phospholipid (34) as an acyl donor, whereas ARAT uses acyl CoA (35). ARAT catalyzes esterification of free retinol (26,27,34,36), while LRAT can use both free retinol and retinol bound to the cellular retinol-binding protein I as a substrate (37). However, it has been shown that an increase in cellular retinol-binding protein (CRBP-I) does not enhance retinyl ester storage in transgenic animals (38). An LRAT partial cDNA was recently cloned from human retinal pigment epithelium cells. This cDNA hybridizes to a major RNA transcript of ~5.0 kb and minor transcripts of 2.2–2.5 kb in several tissues, including the testis and liver (39). The ARAT gene has not yet been cloned.

While the functions of retinyl esters are not fully understood, it is believed that retinyl esters act as a storage form for retinol both in the liver and in many other tissues in the body. We have recently shown that carcinoma cells of the breast, oral cavity and skin are deficient in the esterification of retinol (18,33). These recent data, together with the aforementioned data, suggest that the lack of retinyl esters in carcinoma cells may be associated with or even contribute to their tumorigenic phenotype (18,33).

Hydrolysis of retinyl esters can also occur, both in hepatic cells and in other types of epithelial cells (4045). Recently, a neutral, bile salt-independent retinyl ester hydrolase (REH) was purified (46) and a hepatic, bile salt-dependent REH was cloned and shown to be identical to pancreatic carboxylester lipase (47). The REH(s) which are responsible for retinyl ester hydrolysis in many extra-hepatic tissues have not been well characterized (for review, see ref. 6), though REHs have been described in tissues and cell types in addition to liver. In retinal pigment epithelium, all-trans-retinyl esters are substrates for an isomerohydrolase which converts the esters into 11-cis-retinol; this is then oxidized and converted to 11-cis-retinaldehyde, the chromophore for rhodopsin and cone pigments (48,49). In adipocytes, there is evidence that retinyl esters can be hydrolyzed by a cyclic AMP-dependent enzyme-like hormone-sensitive lipase (50).

In contrast to normal epithelial cells, there are some reports that retinol, although readily taken up by normal human fibroblasts, is not metabolized to either RA or retinyl esters in these cells (51,52). In another study, it was reported that cultured human dermal fibroblasts, treated with retinol, metabolized retinol to RA and retinyl esters (53). Little is known about retinol metabolism in normal human endothelial cells. It was previously reported that isolated endothelial cells from the liver contained very low levels of retinoids (28). However, retinoids can influence endothelial cell growth, morphology and gene expression (5458).

We report here the analysis of retinol metabolism in several types of cultured normal human cells, including fibroblasts, endothelial cells and various types of epithelial cells. We demonstrate that normal epithelial cells, but not fibroblasts and endothelial cells, esterify large amounts of retinol. We also show that these retinyl esters can be hydrolyzed in normal epithelial cells to retinol, RA and other retinol metabolites over ~6–8 days. In contrast, cultured human carcinoma cells esterify very little or no retinol, and thus have essentially no or extremely limited retinyl ester stores. Finally, we demonstrate that the lack of ability to esterify retinol in the oral cavity, skin and breast carcinoma cell lines is correlated with a great reduction in LRAT protein levels.


    Materials and methods
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 Materials and methods
 Results
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Materials
Radiolabeled retinol (all-trans-[11,12-3H], sp. act. 27–47 Ci/mmol) was purchased from New England Nuclear/Dupont (Boston, MA). All other chemicals used, unless specified, were purchased from Sigma Chemical Co. (St Louis, MO).

Cells and culture conditions
The origins and properties of the cell strains used have been described previously (18,33,59,60; see also Table IGo). The fibroblast cell strains CCD42-SK and GM0970 were obtained from the American Type Culture Collection (Rockville, MD). The normal human umbilical vein endothelial cells (HUVEC), normal human bronchial epithelial cells (NHBE), normal human mammary epithelial cells (HMEC) and normal human epidermal keratinocytes (NHEK) were from Clonetics Corp. (Walkersville, MD). The ADO74 and 184B5 lines were from Dr M.Stampfer. For maintenance of the cell strains, the NT2 human teratocarcinoma line was cultured in Dulbecco's modified Eagle's medium (DMEM) plus 10% fetal calf serum (FCS); OKF4, OKP-7 and SCC-25 were cultured in keratinocyte serum-free medium (Life Technologies, Grand Island, NY) according to the manufacturer's instructions; SCC-40 cells were maintained in a mixture of DMEM and Ham's F12 medium (1:1) supplemented with 5% fetal calf serum (FCS), 0.4 µg/ml hydrocortisone, 10 µg/ml epidermal growth factor and 5 µg/ml insulin. MCF-7 and MDA-MB-231 were cultured in DMEM plus 10% FCS and 5 µg/ml insulin. ADO74 cells (61,62) were cultured in MEGM (Clonetics); CCD42-SK and GM0970 were cultured in DMEM plus 10% FCS. The HUVEC, NHBE, HMEC and NHEK were cultured in endothelial growth medium (EGM), bronchial epithelial growth medium (BEGM), mammary epithelial growth medium (MEGM) and keratinocyte growth medium (KGM), respectively (Clonetics). None of the normal cell strains used for experiments had been passaged more than eight times. For radiolabeling and for northern and western analyses of all cell strains and lines, DMEM plus 5% FCS was used. The cells were switched to this consensus medium when [3H]retinol was added.


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Table I. Summary of human cell strains and lines
 
[3H]Retinol radiolabeling
All retinoid solutions and samples were handled under red or dim light. Cells were plated at 1x106 cells per 60 mm dish 24 h before [3H]retinol addition. Cells were washed three times with the consensus medium before labeling and cultured for various periods in 2 ml of labeling medium containing 50 nM [3H]retinol in the consensus medium (~2 µCi/ml). A separate control consisting of labeling medium without cells was included during the incubation period. Cells and one quarter of the medium were collected. Cells were washed once with 0.5 ml phosphate-buffered saline (PBS) and removed from the monolayer in 0.5 ml PBS by scraping. Samples were stored at –70°C until needed for retinoid extraction. The cell numbers were counted from parallel dishes from each treatment at the time of cell harvest.

For the pulse–chase experiments, cells were cultured for 24 h in 100 nM [3H]retinol. Cells were then rinsed three times with warm PBS over a 30 min period and incubated in growth medium without [3H]retinol for an additional 1–8 days. At various times after the removal of [3H]retinol from the medium, dishes of cells were harvested and retinoids were extracted and subjected to high performance liquid chromatography (HPLC) analysis.

Extraction of retinoids and HPLC
The retinoids were extracted as described previously (63). Non-radiolabeled retinoid standards were added to the samples before extraction. Briefly, 350 µl acetonitrile/butanol (50:50, v/v), 0.1% butylated hydroxytoluene was added to 0.5 ml of cells or medium samples. The mixtures were vortexed thoroughly for 30 s. After addition of 300 µl of a saturated (1.3 kg/l) K2HPO4 solution and thorough mixing, the samples were centrifuged for 10 min at 3000xg. The upper organic layer was collected and transferred to an injector vial for automated HPLC analysis.

HPLC analysis was performed using a Waters Millenium system (Waters Corp., Milford, MA) to separate the various retinoids. Samples were applied to an analytical 5 µm reversed-phase C18 column (Vydac, Hesperia, CA) at a flow rate of 1.5 ml/min. The gradient consisted of a 35 min linear gradient from 15 mM ammonium acetate, pH 6.5, in water to 85% acetonitrile in a 10 min linear gradient from 85% acetonitrile to acetonitrile–dichloromethane (80:20) followed by a 15 min hold. Non-radiolabeled retinoid standards were run concurrently and monitored at a wavelength of 340 nm while an A-500 radiochromatography detector (Packard Instruments, Downers Grove, IL) was used to monitor the labeled retinoids.

Retinoids were identified by HPLC based on at least two criteria: an exact match of the retention times of unknown peaks with those of authentic retinoid standards and identical UV spectra (220–400 nm) of unknowns against spectra from authentic retinoid standards during HPLC by the use of the photodiode array detector. RA was also identified by the shift of the retention time of the methylated RA derivative to the same position as the corresponding methyl ester of the RA standard. The methyl ester of RA was synthesized by reaction with diazomethane (15).

RNA isolation and northern blot analysis
Total cellular RNA was isolated from cultured cells using RNA Stat-60 (Tel-Test, Friendswood, TX) according to the manufacturer's instruction. RNA was size-fractionated by electrophoresis on 1% agarose/2.2 M formaldehyde gels, transferred to nylon filters by blotting and attached to the filters using a UV Stratalinker 1800. The cDNA probes used in this analysis were radiolabeled with [32P]dCTP using a random primer labeling kit (Boehringer Mannheim, Indianapolis, IN) according to the manufacturer's directions. Glyceraldehyde phosphate dehydrogenase cDNA was used as a probe for northern blots as described previously (18,64). Aldehyde dehydrogenase-2 (AHD-2) cDNA, isolated from a murine liver cDNA library by this laboratory (65), was used as a probe; this cDNA encodes an aldehyde dehydrogenase class I enzyme, also called ALDH-1 in human. The human LRAT cDNA clone was an EcoRI fragment (840 bp) as described (39). The human neutral, bile salt-independent REH and the hepatic, bile sale-dependent REH were EST clones #g4069398 and #g2237729, respectively, from Genome Systems (St Louis, MO).

Blots were prehybridized and hybridized at 42°C in 50% (w/v) formamide/5x SSC, 50 mM NaH2PO4, pH 7.4, 5 mM EDTA, 0.08% polyvinylpyrrolidone, 10% (w/v) bovine serum albumin and 10% (w/v) salmon sperm DNA. After 10–16 h of hybridization, blots were washed twice in 2x SSC, 0.1% SDS for 20 min at room temperature, and twice in 0.2x SSC, 0.1% SDS at 50°C.

Western analysis
This procedure was carried out as described previously (39). Briefly, polyclonal antisera were generated in rabbits to a mixture of two different LRAT peptides. Total cell protein was used, and blot analysis on nitrocellulose filters was performed using antiserum diluted to 1:1000 for detection of LRAT. Protein bands were detected by the ECL system (Pierce, Rockford, IL).


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 Materials and methods
 Results
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 References
 
Analysis of metabolism in normal cultured human fibroblasts and in endothelial and epithelial cells
We examined two different normal human fibroblast cell strains, GM0970 and CCD42-SK, for their ability to metabolize [3H]retinol (see Table IGo for a list of cell strains and lines analyzed). Essentially no retinol metabolism occurred in GM0970 (Figure 1AGo) and CCD42-SK (data not shown) even after 15–24 h of culture in the presence of [3H]retinol. We also examined HUVEC for their ability to metabolize [3H]retinol. Again, only trace amounts of retinyl esters were found in these cells even after 15 h of culture in the presence of 50 nM [3H]retinol (Figure 1BGo).



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Fig. 1. Metabolism of [3H]retinol in various types of normal cells. HPLC tracings of (A) GMO970, a human normal fibroblast cell strain, (B) a primary culture of HUVEC, (C) HNBE cells and (D) NHEK. Cells were cultured in the presence of [3H]retinol for varying times; 15 h is shown. Cells and one quarter of the medium were harvested, and retinoids were extracted and separated by reversed-phase HPLC analysis as described in Materials and methods. Only the intracellular retinoids are shown. Non-radiolabeled retinoids were included with each sample as standards to determine the elution times of the various retinoids. Arrows indicate the elution positions of [3H]RA (RA, 18 min) and [3H]retinol (ROH, 29.8 min), and the bracket indicates the [3H]retinyl esters at 48–57 min. Note that the y axes are different in the four panels; there were 1.3x106 GM0970 cells, 1.2x106 HUVEC, 1.0 x106 HNBE cells and 9.7x105 NHEK. This experiment was performed three times with very similar results. Data from one experiment are shown here.

 
In contrast, both NHBE (Figure 1CGo) and NHEK (Figure 1DGo) exhibited extensive esterification of [3H]retinol to retinyl esters, with the predominant retinyl ester being retinyl oleate. After 15 h of culture in the presence of [3H]retinol in the medium, the concentration of [3H]retinol was much higher in NHBE than in NHEK (Figure 1Go, compare panels C and D). We observed similar levels of [3H]retinol esterification in exponentially growing cultures of epithelial cells and in confluent cultures (data not shown).

Thus, we conclude that normal human epithelial cells take up and esterify a large proportion of the [3H]retinol to which they are exposed in the medium, consistent with our previous data using normal human mammary epithelial cells (33) and normal cell strains from the oral cavity and skin (18). In contrast, human fibroblast strains and HUVEC do not esterify retinol under these conditions. Our data are consistent with two previous reports relating to cultured fibroblasts in which it was shown that there was no significant retinol metabolism in murine 10t1/2 cells (51) and human dermal fibroblasts (52). This lack of retinol esterification in normal human dermal fibroblasts and endothelial cells may reflect a less important role for endogenous retinoids in these cell types.

Retinol esterification in cultured normal epithelial cell strains and carcinoma cell lines
Normal mammary epithelial cell strains (HMEC and ADO74), NHEK and an oral cavity epithelial cell strain (OKP-7) were cultured for various times in the presence of [3H]retinol. Approximately 6–8 h after 50 nM [3H]retinol had been added to the medium, it had all been esterified. Figure 2Go shows retinoids extracted from cells 22 h after addition of [3H]retinol to the culture medium [except for ADO74 (33)]. It can be seen that there is almost no [3H]retinol in the normal epithelial cells, as it has all been converted to various types of [3H]retinyl esters, with [3H]retinyl oleate and [3H]retinyl palmitate the most prevalent (Figure 2A–CGo). The MCF-7 breast carcinoma cell line, in contrast, even after 22 h of culture in the presence of [3H]retinol in the medium, did not exhibit [3H]retinol esterification (Figure 2EGo). While some [3H]retinol esterification occurred in the SCC-12 skin squamous carcinoma line (Figure 2FGo), the SCC-40 carcinoma line, from the soft palate, did not show significant [3H]retinol esterification even after 22 h of exposure to [3H]retinol in the medium (Figure 2GGo). As a result, the intracellular levels of [3H]retinyl esters in the tumor cell lines cultured in the presence of [3H]retinol are much lower than those achieved in the normal cell strains, and much less of the total [3H]retinol added to the medium is metabolized by the tumor cells (compare Figure 2Go panels E–G with panels A–C; and Figure 2Go, panels I and J for quantification).



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Fig. 2. Metabolism of [3H]retinol in normal human epithelial cell strains [(A) HMEC, (B) NHEK and (C) OKP-7] and human carcinoma lines [(E) MCF-7, (F) SCC-12 and (G) SCC-40]. Non-radiolabeled standards are shown in panels D and H; 1, all-trans-4-oxoretinol; 2, all-trans RA; 3, all-trans-retinol; 4, all-trans-retinaldehyde; 5, all-trans-retinyl acetate; 6, retinyl palmitate. Cells were labeled for 22 h with 50 nM [3H]retinol. Cells and one quarter of the medium were harvested, and retinoids were extracted and separated by reversed-phase HPLC analysis. Only the intracellular retinoids are shown. Non-radiolabeled retinoids were included with each sample as standards to determine the elution times of the various retinoids. The data for each sample are plotted as [3H] counts per minute against time. The peaks that correspond to [3H]retinol and [3H]retinyl esters are at 30.5 min and 47–56 min, respectively. The data from panels A, B, C, E, F and G are shown in a quantitative format in panels I and J. Panel I shows the intracellular [3H]retinyl ester levels normalized to 1x106 cells. Panel J shows the total [3H]retinol (intracellular plus in the medium) remaining at 22 h, calculated as described previously (18,64). This experiment was performed three times with very similar results. One representative HPLC tracing for each cell line is shown here.

 
[3H]Retinyl ester content in normal epithelial cell strains and carcinoma cell lines after [3H]retinol removal from the medium
As the carcinoma cells appeared to be unable to store retinol in the ester form, we next investigated how the concentrations of various retinoids in these cells changed after removal of [3H]retinol from the medium. Therefore, in this series of experiments the cells were cultured in the presence of 100 nM [3H]retinol and 900 nM retinol in the medium for 24 h, after which [3H]retinol was removed from the medium. The cells were washed several times. At various times after the removal of [3H]retinol from the medium, dishes of cells were harvested, retinoids were extracted and [3H]retinoids were analyzed by HPLC. In the normal cell strains from the oral cavity (OKP-7) and the breast (ADO74), substantial amounts of [3H]retinyl esters were present at the time of [3H]retinol removal from the medium (day 0, Figure 3Go). The intracellular concentrations of [3H]retinyl esters remained high the next 4–8 days (Figure 3Go, and data not shown). Intracellular concentrations of [3H]RA and [3H]retinol were 1–2 µM and 3–10 µM, respectively, in the OKP-7 and ADO74 normal cell strains over this time period (Figure 3A and BGo; see also panel E, an enlarged scale of internal [3H]RA from panels A and B). Even 4 days after the removal of [3H]retinol from the medium, the intracellular concentration of [3H]retinyl esters was ~26 µM in the OKP-7 cells and 30 µM in ADO74 cells.



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Fig. 3. Kinetics of [3H]retinol metabolism in (A) OKP-7, (B) ADO74, (C) SCC-40 and (D) MDA-MB-231 cells. Cells were radiolabeled with 100 nM [3H]retinol plus 900 nM retinol for 24 h. Cells were then washed three times with PBS and the medium was replaced with fresh growth medium (day 0). At various times thereafter, cell samples and samples of medium were harvested, and retinoids were extracted and separated by reversed-phase HPLC analysis. Cell numbers for each cell line were determined by counting a separate dish, seeded with the same number of cells, at the time of harvest. The intracellular concentration of each of the [3H]retinyl esters, [3H]retinol and [3H]RA produced during the chase period after removal of the [3H]retinol label on day 0 was calculated as described (18,33) and plotted (on the y axis) against days after [3H]retinol removal (on the x axis). This experiment was performed twice, with very similar results. Data from one experiment are shown here. Note the differences in the y axes between the normal cell strains and the carcinoma lines. (E) intracellular levels of [3H]RA from panels A–D, on a more sensitive scale. ROH, retinol; RE, retinyl esters.

 
In contrast, the SCC-40 squamous cell carcinoma line from the oral cavity and the MDA-MB-231 breast carcinoma line had no detectable retinyl esters at the time of [3H]retinol removal (day 0, Figure 3C and DGo). The [3H]retinol concentration in these cells decreased very rapidly after removal of [3H]retinol from the medium, and within 48 h no [3H]retinol was detectable in SCC-40 and MDA-MB-231 cells (Figure 3Go, note the difference in y axes between the graphs of retinoid concentrations in the normal epithelial cell strains and the carcinoma lines). Similar results were obtained for the breast carcinoma line MCF-7 (data not shown). We conclude from this experiment that, in the normal cell strains, retinyl esters are slowly hydrolyzed over a period of several days to generate retinol and more bioactive retinoids such as RA. In contrast, the carcinoma lines become profoundly retinoid deficient when cultured in the absence of retinol in the medium.

Expression of genes encoding enzymes involved in retinol metabolism in cultured normal human epithelial, endothelial and fibroblast cells
We next examined normal human cell strains for expression of genes encoding various enzymes involved in the metabolism of retinol, including genes encoding enzymes involved in the conversion of retinol to RA and the gene encoding LRAT. For these experiments cells were cultured in the presence or absence of 1 µM unlabeled RA for 48 h, followed by cell harvesting and RNA isolation. HUVEC and the normal fibroblast strains CCD42-SK and GM0970 did not express detectable levels of LRAT transcripts (Figure 4AGo). In HMEC and HNBE cells, expression of both the 5 kb and 2.5 kb LRAT transcripts was detected (Figure 4BGo). The human NT2 teratocarcinoma cell line, which esterifies large amounts of retinol (X.Guo and L.J.Gudas, unpublished), also expresses high levels of the 5 kb and 2.5 kb LRAT transcripts (Figure 4A and BGo). Thus, the levels of LRAT mRNA expression correlate with the abilities of these various normal cell types to esterify retinol, as shown in Figures 1 and 2GoGo. Culture in the presence of RA for 48 h did not greatly affect the levels of LRAT mRNA in these cells; a slight increase in LRAT mRNA was noted after RA addition.



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Fig. 4. Northern blot analysis of LRAT transcripts in cultured normal cell strains, cultured normal epithelial cell strains and cultured carcinoma cell lines. Cells were cultured as described in Materials and methods, in the presence (+) or absence (–) of 1 µM RA for 48 h. RNA was isolated from the cells and northern blot analysis was performed. Total RNA (10 µg) was loaded in each lane. Autoradiograms of blots hybridized to 32P-radiolabeled cDNA probes are shown. These experiments were performed three times with very similar results; one experiment is shown. (A) Cell lines CCD-42SK, GM0970, HUVEC and NT2; (B) cell strains NT-2, HMEC and HNBE; (C) cell lines ADO74, MCF-7 and MDA-MB-231. See Table IGo for details of cell strains and lines. Top, human LRAT cDNA probe; bottom, GAPDH cDNA probe for a loading control. Transcripts of 5 and 2.5 kb are shown by larger bold lines, and transcripts of 3.0 and 1.5 kb by smaller lines. The exposure times for LRAT (2 days) and GAPDH (12 h) were the same for all of the lanes.

 
With the exception of the human endothelial cells, none of these various human cell strains, when cultured in the presence or absence of RA, expressed detectable levels of AHD-2 mRNA; however, the mouse liver cells, used as a positive control expressed a high level of AHD-2 mRNA (data not shown). AHD-2, which is called ALDH-1 in humans, can use retinaldehyde as a substrate, converting it to RA. The RA hydroxylase (CYP26) gene was not expressed by any of these normal cell strains at detectable levels (data not shown).

A comparison between normal and tumor cell lines of the expression of genes encoding enzymes for retinol metabolism
In the next series of experiments, the expression of LRAT mRNA was examined in a normal cell strain and in two carcinoma lines (Figure 4CGo). The normal mammary epithelial cell strain ADO74 expressed two major LRAT transcripts of 5 and 2.5 kb (see bold lines, Figure 4CGo). MDA-MB-231 and MCF-7, estrogen receptor-negative and -positive breast cancer lines, respectively, did not express detectable levels of LRAT transcripts of 5 and 2.5 kb. However, these tumor lines had transcripts of ~3 and 1.5 kb (narrower lines, Figure 4CGo). These aberrantly sized transcripts may reflect the use of alternative polyadenylation sequences in the tumor cells, alternative transcription start sites or alternative splicing. In summary, the expression of the 5.0 and 2.5 kb LRAT transcripts by the normal epithelial cells correlates well with the ability of the cells to esterify retinol.

The AHD-2 gene was not expressed at detectable levels in the cultured normal human cell strain or tumor cell lines shown in Figure 4CGo (data not shown). We did not detect transcripts of the bile salt-dependent REH gene or the bile salt independent REH gene in the normal cell strain ADO74 or in the tumor lines (data not shown). The RA hydroxylase (CYP26) gene was strongly expressed by the MCF-7 cell line after RA addition, but was not expressed at detectable levels in other tumor lines or normal cell strains cultured with or without RA (data not shown). Thus, there was no correlation of the expression of these enzymes with the tumor cell phenotype.

Analysis of LRAT protein levels in normal cell strains and carcinoma cell lines
In all of the normal human epithelial cell strains from breast, skin and oral cavity, cultured in consensus medium, an intense protein doublet of 62–65 kDa was detected which was reactive with the anti-LRAT antibody (Figure 5Go); the levels of these proteins did not change when the cells were first cultured for 48 h in the presence of 1 µM exogenous RA and then harvested for western analysis (data not shown). In contrast, in oral cavity, skin and mammary carcinoma lines, no protein reacted with the anti-LRAT antibody (Figure 5Go). Since small amounts of [3H]retinyl esters can be seen in SCC-12 cells (Figure 2Go), either ARAT or another enzyme must carry out esterification in these cells or a very small amount of LRAT protein, undetectable in this western assay, must be present in the SCC-12 cells. However, the major conclusion from these data is that the normal epithelial cell strains contained much higher levels of LRAT protein than the carcinoma lines (Figure 5Go). In human retinal pigment epithelium (Figure 5Go), a protein band of 25–26 kDa was observed by western analysis, using polyclonal antisera raised against a mixture of two LRAT peptides (39).



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Fig. 5. Western blot analysis of human LRAT protein. Ten micrograms of microsomal protein from tissues or 10 µg of whole cell lysate from cultured cells was loaded in each lane. Lane 1, retinal pigment epithelial cells; lane 2, ADO74; lane 3, MCF-7; lane 4, MDA-MB-231; lane 5, NHEK; lane 6, SCC-12; lane 7, SCC-13; lane 8, OKF4; lane 9, SCC-25; lane 10, OKP7; lane 11, SCC-40 (see Table IGo for description of lines). A rabbit polyclonal antiserum against a mixture of two LRAT peptides was used for the detection; the antiserum was used at a 1:1000 dilution. Molecular weight markers in kilodaltons are indicated on the left. This experiment was performed three times with similar results. One experiment is shown here.

 

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We have previously demonstrated that, relative to normal human cell strains, human carcinoma cell lines have a greatly reduced ability to metabolize [3H]retinol to [3H]retinyl esters (18,33). In this report, we extend these experiments to the analysis of the expression levels of one of the enzymes involved in retinol esterification, LRAT. We previously demonstrated that microsomal ARAT enzyme activity was lower in carcinoma cells than in normal epithelial cells (18). Under the conditions of our prior assays (18), we were primarily detecting ARAT enzyme activity and not LRAT enzyme activity in the microsomal protein extracts in the normal cell strains and the tumor cell lines. In this report, we demonstrate that LRAT protein levels are lower in the carcinoma cell lines than in the normal epithelial cell strains (Figure 5Go). Since the ARAT gene has not been cloned, we do not know if ARAT mRNA and protein levels are lower in the tumor cells, though we would predict that this is the case.

We show that, even though the carcinoma cells contained some [3H]retinol when cultured in the presence of [3H]retinol, as soon as the medium was changed and the [3H]retinol was removed, the carcinoma cells had essentially no detectable internal [3H]retinyl ester stores or internal [3H]retinol (Figure 3Go). The lack of retinyl esters in the tumor cells most probably results from the low level of LRAT protein in the carcinoma cells relative to normal cell strains (Figure 5Go).

We found that the transcripts for the neutral bile salt-independent REH and the hepatic bile salt-dependent REH genes are not expressed at detectable levels in the normal and tumor cell lines (data not shown). These may not be the only genes encoding enzymes involved in retinyl ester hydrolysis; alternatively, a very low expression level may be sufficient to carry out the [3H]retinyl ester hydrolysis observed in the normal cell strains (Figure 3Go).

The mechanism by which the expression of the LRAT transcripts and protein (Figures 4 and 5GoGo) is altered in the carcinoma cells as compared with the normal epithelial cells is unclear at this time. As LRAT genomic clones containing the promoter region and other potential regulatory regions are not yet available, we have not determined why the LRAT transcripts in the tumor lines are of aberrant sizes (Figure 4BGo). However, we think that aberrant splicing, leading to the abnormally sized transcripts of 3 and 1.5 kb in the tumor cells, is the most likely explanation. These abnormally sized LRAT transcripts in the tumor cells apparently cannot be translated into LRAT protein, since no LRAT protein could be detected in these cells by western analysis whereas in normal cells there was a high level of an LRAT protein doublet at a molecular mass of 62–65 kDa (Figure 5Go).

It was previously shown that a major LRAT transcript of 5 kb is present in several tissues (39), consistent with our data for the sizes of the LRAT transcripts in normal epithelial cells (Figure 4Go). However, it was surprising that the mass of the LRAT protein, as revealed by western blot analysis, varies among normal tissues (Figure 5Go). We have performed RT-PCR analysis of the mRNA from ADO74, a normal human mammary epithelial cell strain, and have determined that the putative coding sequence (data not shown) is identical to the entire open reading frame from retinal pigment epithelium and so probably contains the peptide sequences against which the antibodies used were directed (39). However, we have preliminary data that the open reading frame continues in the 5' direction in LRAT from the ADO74 cells (X.Guo, unpublished). Therefore, although we do not yet know the mechanism(s) whereby the larger protein species arise in some tissues (e.g. multiple transcription start sites, alternative splicing), we are confident that the antibodies are LRAT-specific.

The functions of many retinol metabolites, including the various esters of retinol, are not fully understood. While data in the literature suggest that retinyl esters play an important role in retinol storage in various cell types in the body (for review, 66), further analysis of the actions of retinyl esters will require methods for altering retinyl ester formation from retinol in cells to assess some of the consequences in terms of cell growth and differentiation. Our data indicating that metabolism of [3H]retinol to [3H]retinyl esters is much lower in carcinoma lines than in normal cell strains may have important clinical implications. In cultured tumor cell lines and biopsies taken directly from patients (59,6775), there are low or undetectable levels of the mRNA for RARß, a RA receptor gene that is RA-inducible in many cell types; RARß has been implicated as a biomarker, reflecting the content of active retinoids in the cells. We suggest that RARß mRNA levels are low in the carcinoma cells in part because of their deficiency in retinyl ester stores relative to normal epithelial cells. The impairment in the ability to convert retinol to retinyl esters in the tumor cells could lead to their inappropriate growth and to the loss of normal differentiation responses because of the lack of a sufficient amount of internal retinol, stored as retinyl esters. With respect to cancer therapy, if the decrease in LRAT mRNA and protein levels results from an oncogene-associated inhibition of gene transcription, it is possible that drugs can be developed which will prevent this inhibition of LRAT transcription. If the low level of LRAT protein in carcinoma lines results from the aberrant splicing of the LRAT gene in the tumor cells, this may be more difficult to correct. An alternative therapy may involve the delivery of retinyl esters directly into the tumor cells, since this may result in higher internal levels of retinoids than those achieved by giving more retinol to the cells.

One of our most striking observations is that LRAT was undetectable in all of the carcinoma cell lines examined so far by western analysis, but present in all of the normal epithelial cell strains examined. We have now examined normal human epithelial cell strains and carcinoma lines from the oral cavity, breast and skin. Our results strongly indicate that this major reduction in LRAT protein is a common feature of human carcinoma cells. Loss of LRAT protein may therefore be a good marker for carcinoma cells. We are currently in the process of examining biopsies of tumors taken directly from patients to assess the levels of LRAT protein in the tumor samples. It will also be important to determine at what point during the process of carcinogenesis the LRAT protein levels decline.


    Notes
 
5 To whom correspondence should be addressed Email: ljgudas{at}mail.med.cornell.edu Back


    Acknowledgments
 
We thank members of the Gudas laboratory for scientific discussions, Drs J.Rheinwald and M.Stampfer for some of the normal cell strains, and T.Resnick for editorial assistance. This research was supported by NIH grant R01DE10389 (L.J.G.); in part by a fellowship from the New York State Department of Health (X.G.); by NIH grant R01EY04096 (R.R.R.); and by NIH grants R01EY00444, R01EY00331, and the Dolly Green Endowed Chair (D.B.).


    References
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 

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Received February 16, 2000; accepted July 27, 2000.