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Journal of Lipid Research, Vol. 48, 1846-1856, August 2007
Copyright © 2007 by American Society for Biochemistry and Molecular Biology


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* ProMPT Genito Urinary Cancer Research Group, Paterson Institute for Cancer Research, The University of Manchester, Manchester, UK
School of Chemical Engineering and Analytical Science, The University of Manchester, Manchester, UK
Department of Urology, Hope Hospital, NHS Trust, Manchester, UK
** Department of Urology, Christie Hospital, NHS Trust, Manchester, UK
Published, JLR Papers in Press, May 11, 2007.
1 To whom correspondence should be addressed. e-mail: EGazi{at}picr.man.ac.uk
| ABSTRACT |
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as+s(C-D)2+3 (D31-PA):
as+s(C-H)2+3 (lipid hydrocarbon) signal. In addition, larger adipocytes were found to consist of a higher percentage of D31-PA of the total lipid found within the adipocyte. Following background subtraction, the
as(C-D)2+3 signal illuminated starved PC-3 cells cocultured with D31-PA-loaded adipocytes, indicating translocation of the labeled FA. This study demonstrates lipid-specific translocation between adipocytes and tumor cells and the use of FTIR microspectroscopy to characterize various biomolecular features of a single adipocyte without the requirement for cell isolation and lipid extraction.
Supplementary key words tumor fatty acids Fourier transform infrared spectroscopy deuterated palmitic acid
Abbreviations: BMS, bone marrow stroma; CaP, prostate cancer; CPD, critical-point drying; D31-PA, deuterated palmitic acid; FFA, free FA; FTIR, Fourier transform infrared; IL-6, interleukin-6; IR, infrared; MSC, mesenchymal stem cell; PF, paraformaldehyde; SR, synchrotron; TAG, triacylglyceride
| INTRODUCTION |
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The linkage between lipids and advanced CaP may also be reflected in the natural history of this disease, because it is well documented that circulating metastatic CaP cells in the peripheral blood possess a predilection for deposition into the bone marrow stroma (BMS) (8), which harbors a rich source of lipids stored within adipocytes. Brown et al. (9) have shown that PC-3 cell (human CaP cell line derived from bone metastases) invasion toward BMS was significantly reduced when the experiment was repeated with BMS grown in the absence of hydrocortisone so that adipocyte formation was not stimulated.
Adipocytes are specialized for the synthesis and storage of FAs as triacylglycerides (TAGs) (10) and for FA mobilization through lipolysis (11) in response to hormones, cytokines, and other factors involved in energy metabolism (12). In recent years, there has been growing interest in understanding the interactions of CaP cells with adipocytes at the molecular level (9, 13, 14). Tokuda et al. (13) have shown that adipocytes cocultured with PC-3 cells influenced not only the differentiation growth patterns of the PC-3 cells but also their proliferative rate. Oil Red O staining for neutral lipids and transmission electron microscopy revealed larger and greater frequency of lipid droplets within the cytoplasm of cocultured PC-3 cells compared with the control (PC-3 monoculture) (13). Previously, we have demonstrated through Fourier transform infrared (FTIR) microspectroscopic imaging that metastatic CaP cells in bone marrow tissue, with close proximity to adipocytes, give rise to higher lipid hydrocarbon signals relative to distant CaP cells (9). FTIR microspectroscopy is an optically based technique that can measure the transitions in vibrational modes (mainly stretching and bending) of the functional groups of biomolecular constituents within cells, as a result of absorption and subsequent excitation by infrared (IR) radiation. The functional group vibrations are representative of biomolecular constituents such as lipids, proteins, carbohydrates, and a variety of phosphorylated molecules. The FTIR spectrum derived from the cell comprises a series of peaks as a function of wave number, which depict the state of chemical bonding (intra- and intermolecular hydrogen bonding, van der Waals interactions, and steric factors) and the relative intensities of the above-mentioned species within the scrutinized area of the cell.
Although CaP cells in the presence of adipocytes exhibit higher lipid signals (9, 13), there have been no data in the literature that unequivocally establishes whether this increase is the result of adipocyte-to-CaP cell lipid transposition or is due to adipocyte-secreted cytokine stimulation of CaP cells, such as with interleukin-6 (IL-6) (14, 15), which may elicit de novo lipid synthesis. IL-6 has been reported to augment de novo lipid synthesis of FAs in hepatocytes of mice (16). Furthermore, accepting that lipid molecules are transferred from adipocytes into CaP cells, it is not known whether this process is selective toward specific FAs that are required by the CaP cell. Here, we use imaging FTIR microspectroscopy as an analytical methodology for i) the biomolecular characterization of the adipocyte following loading with deuterated FA and ii) studying the translocation of deuterated FA between adipocytes and PC-3 cells, following fixation.
| MATERIALS AND METHODS |
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Cell fixation
Cultures on MirrIR plates were washed twice with PBS and then fixed in 4% paraformaldehyde (PF) in PBS for 25 min. Following this, the cells were washed three times (5 min each wash) in Sorensen's buffer (0.15 M, pH 7.4) to remove free PF. The cells were postfixed in 1% osmium tetroxide (OsO4) for 1 h and washed a further three times (5 min each wash) with Sorensen's buffer prior to dehydration using increasing concentrations of ethanol-water (30:70, 50:50, 70:30, 90:10) for 5 min. The cells were placed twice in 100% ethanol for 5 min. The cells were then transferred to Arklone (112 trichloro-122 trifluroethane; TAA Laboratories, UK) and placed into the precooled (9°C) chamber of the critical-point drier (Bal-tec CPD-030). The Arklone was substituted for liquid CO2 following five Arklone-CO2 exchanges, and then a 5 min immersion (this procedure was repeated twice again). Phase-transition was induced by heating the chamber to 40°C and at 80 bar pressure. Preservation was assessed by a high-power optical microscope. The cells were stored in a desiccator until FTIR analysis. A separate set of adipocytes on MirrIR was fixed in 4% formalin in PBS for 20 min. These cells were then briefly washed in distilled water for 3 s to remove residual PBS from the surface of the cells (17). The cells were dried under ambient conditions and stored in a desiccator until FTIR analysis.
FTIR microspectroscopy
A TAG mix [containing triacetin (C2:0), tributyrin (C4:0), tricaproin (C6:0), tricaprylin (C8:0), and tricaprin (C10:0)] (Sigma-Aldrich) was analyzed in reflection mode by FTIR after smearing a small amount onto an MirrIR slide. The FTIR spectrum was collected using a Nicolet Magna 550 spectrometer, coupled to a Nicplan microscope equipped with a liquid nitrogen-cooled mercury cadmium telluride detector, using a sampling aperture size of 60 µm x 60 µm. The FTIR spectrum represents an average of 512 scans in the wave number range 750–4,000 cm–1 with a resolution of 4 cm–1. The sample spectrum was ratioed to a background spectrum that was recorded on a blank area of the plate to correct for the instrument function plus atmospheric CO2 and water vapor absorptions. FTIR spectra of the subcellular lipid deposit of a formalin-fixed, water-rinsed, and air-dried adipocyte and a PF-OsO4-CPD-fixed adipocyte were collected at the Daresbury Laboratory (Beamline 11.1) using synchrotron (SR)-based FTIR microspectroscopy. The SR source provides high signal-to-noise spectra with aperture sizes close to the diffraction limit of IR light. For this, a Nicolet Nexus FTIR spectrometer equipped with a liquid nitrogen-cooled mercury cadmium telluride detector and a KBr beam splitter coupled to a Nicolet Continum microscope was used in reflectance mode with a 10 µm x 10 µm sampling aperture. The sample spectrum was collected at 4 cm–1 spectral resolution with 128 scans and was ratioed to a background spectrum that was recorded on a blank area of the plate. High-definition FTIR microspectroscopic maps of PF-OsO4-CPD-fixed adipocyte-PC-3 cell cocultures were collected in reflectance mode at 6.25 µm pixel resolution in rapid-scan mode using a Perkin Elmer Spotlight spectrometer with a 16 x 1 MCT linear array detector. The background scan was recorded at 8 cm–1 spectral resolution with 75 scans, whereas the sample scan was recorded at 8 cm–1 spectral resolution with 64 scans. A charged-coupled detector camera, integrated into the Spotlight spectrometer, was used to obtain optical images of the mapping area.
Data analysis
The FTIR spectra of the TAG reference standard and of the lipid deposit of a formalin-fixed, water-rinsed, and air-dried adipocyte were recorded using the OMNIC v.5.1a software, which also included the Atlµs video capturing program. FTIR spectral maps of PF-OsO4-CPD-fixed adipocyte-PC-3 cell cocultures were processed with Spotlight version 1.0.1. This software permitted the intensity of baseline corrected areas of selected peaks as well as ratio of peaks to be mapped across the map. Adipocyte size in micrometers was determined with ImageJ (version 1.36b) software using optical images captured at x320 by the charged-coupled detector camera attached to the SR-FTIR Nicolet Nicplan microscope.
| RESULTS |
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s(C=O) peak at 1,744 cm–1 of the same frequency of absorption as the lipid ester
s(C=O) peak in the reference TAG spectrum (Fig. 1Di). Note: the lipid ester
s(C=O) peak of glycerol-bound FAs is at higher frequency compared with the lipid
s(C=O) peak of free FAs (1,725–1,690 cm–1) (19–21). Several other characteristic peaks of the glycerol moiety of TAG are also observed in the lipid deposit of the formalin-fixed adipocyte at frequencies >1,500 cm–1, and these are identified in Fig. 1Dii as peaks 8–12. However, the peaks in this spectrum (Fig. 1Dii) are broader, compared with the same peaks in the reference TAG spectrum (Fig. 1Di). This is due to the collapse of the lipid droplets, which gives rise to a range of bonding strengths with neighboring molecular species for those functional groups absorbing at frequencies >1,500 cm–1.
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s(=C-H) signal from unsaturated hydrocarbons is present in the lipid deposit spectrum of the formalin-fixed, water-rinsed, air-dried adipocyte (Fig. 1Dii), but is not observed in the lipid droplet spectrum of the PF-OsO4-CPD adipocyte (Fig. 1Diii). Additionally, both methods of fixation (formalin-fixed, water-rinsed, air-dried and PF-OsO4-CPD) result in a decrease in peak resolution of the
as(CH)2 and
as(CH)3 modes, which is also the case for the
s(CH)2 and
s(CH)3 modes.
FTIR biomolecular characterization of adipocytes
Figure 2Ai
shows an optical photomicrograph of a cluster of adipocytes, loaded with isotopically labeled palmitic acid (D31-PA), following PF-OsO4-CPD fixation. The dark features within this optical image represent the lipid droplets retained within the fully differentiated adipocytes. A negative image (Fig. 2Aii) of Fig. 2Ai provides better visualization of these lipid droplets within the adipocytes. These adipocytes are surrounded by a potentially heterogeneous population of stroma cells. Typical raw FTIR spectra extracted from a fully differentiated adipocyte and from a stroma cell location [points 1 and 2, respectively, in the optical photomicrograph (Fig. 2A)] are shown in Fig. 2B. The incorporation of D31-PA into the adipocyte lipid droplets is confirmed by the presence of the C-D peaks that are clearly observed within the spectral region 2,250–2,000 cm–1 (Fig. 2B). This spectral region does not overlap with IR absorbencies from endogenous FAs or other endogenous biomolecular functional group vibrations arising from the adipocyte, as seen in the adipocyte spectrum incubated with no D31-PA (Fig. 1Diii).
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s(C=O) peak, glycerol and lipid hydrocarbon signals, and relatively lower amide I and II signals. The area shown in Fig. 2A was analyzed with imaging FTIR microspectroscopy, and the resulting FTIR peak extracted images are shown in Fig. 2C–F. The method of baseline integration (blue line) and peak areas (in gray) used to generate the images in Fig. 2C–F are shown in Fig. 2B.
Figure 2C shows the lipid hydrocarbon:amide I peak area intensity distribution across this imaging field. As expected, the lipid hydrocarbon signal intensity (Fig. 2C) is greatest in the lipid droplets contained within the adipocytes and is lower in the stroma cells. This signal distribution is also reflected by the lipid ester
s(C=O) intensity map shown in Fig. 2D, which provides better contrast between adipocytes and stroma cells, compared with the lipid hydrocarbon signal. The intensity distribution of the
as(C-D)2+3 (Fig. 2E) also demonstrates localization with high intensity at all adipocytes in the imaging field, indicating that all adipocytes have incorporated D31-PA. The between-adipocyte differences in D31-PA incorporation are further investigated below. We also find that the
as(C-D)2+3 signal is located at reduced intensity within the surrounding stroma cells.
In Fig. 2F, we demonstrate how FTIR technology can be applied to determine the subcellular deposits of highly concentrated D31-PA, among other lipids, in a single adipocyte. Figure 2Fi shows a high-magnification FTIR image of adipocyte 1 (indicated in the corresponding optical image in Fig. 2A) using the lipid ester
s(C=O):amide I intensity distribution, because it provides good contrast between the adipocyte and surrounding stromal cells. Figure 2Fii shows the
as(C-D)2+3:lipid hydrocarbon signal distribution for adipocyte 1 and is used to identify regions of the most highly concentrated D31-PA relative to nonisotopically labeled lipids. For this, the
as(C-D)2+3 signal was ratioed against the lipid hydrocarbon signal in preference to the lipid ester
s(C=O) signal, because the latter originates from both nonisotopically labeled lipids and D31-PA. A merge of Fig. 2Fi (color coded with red channel) and Fig. 2Fii (color coded with green channel) gives rise to Fig. 2Fiii, which shows the subcellular localizations of highly concentrated D31-PA molecules (yellow pixels), designated by white arrows.
We further investigated the lipid composition within adipocyte 1 by determining the subcellular locations of the most-concentrated TAG store. As mentioned earlier, although the frequency of absorption for the lipid
s(C=O) of FFA and TAG overlap, the peak maxima occur at well-separated wavenumbers. The peak maximum for TAG arises at 1,741 cm–1, whereas the lipid
(C = O) peak maximum for the FFA standard absorbs at 1,714 cm–1 (21). However, these wavenumber values may shift in biological samples, depending on the bonding environment of the lipid
s(C=O) moiety. Gomez-Fernandez et al. (19) report that the protonated carboxyl (COOH) group of FFAs appears at
1,725 cm–1, and Chalmers (20) has assigned a wave number range of 1,690–1,710 cm–1. We use the upper limit of absorption for the lipid
s(C=O) moiety as 1,725 cm–1, because one can observe in our spectra, reported in Fig. 3A
, a shoulder on the lipid
s(C=O) peak at this wave number due to the FFA, which also conforms to Gomez-Fernandez et al. (18). In addition, we have previously shown that the intensity of absorption of wavenumbers below 1,725 cm–1 increases as the relative ratio of FFA to glycerol-bound lipids increases (21).
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Adipocyte 1, shown in Fig. 3Ai, was magnified and presented in Fig. 3Bi. Figure 3Bii shows baseline corrected FTIR spectra for the lipid
s(C=O) spectral region, extracted from three different locations in adipocyte 1 (marked 1–3 in Fig. 3Bi). These spectra show that there is an increase in absorption from TAG at >1,725 cm–1, between the spectrum collected at the cell periphery and spectra collected at internal locations. Accompanying this, there is an increase in cell thickness between the cell periphery and location 3, as depicted by the lipid intensity image (inset in Fig. 3Bi). This confirms that the storage of TAG molecules in the adipocyte is the dominating factor influencing its volume. In contrast to the intracellular TAG concentration, there appears to be little change in the FFA absorption at <1,725 cm–1, between spectra collected at the periphery (location 1) of the adipocyte and those collected at internal locations (locations 2 and 3). Edens et al. (22) report that in adipose tissue, stored TAG undergoes continuous, simultaneous synthesis and breakdown. The data reported here (Fig. 3) suggest that within a single adipocyte, the FFAs are broken down or are in the process of being synthesized into TAG around an internalized TAG store.
We next studied the amount of D31-PA incorporation, relative to other FAs from the adipogenic media, between adipocytes, and its association with cell size. SR-FTIR spectra were obtained from 20 different PF-OsO4-CPD adipocytes that varied in size. The baseline corrected lipid hydrocarbon
as+s(C-H)2+3 peak area was used to represent the amount of FAs other than the isotopically labeled D31-PA, which was represented by the intensity of the
as+s(C-D)2+3 peak area (Fig. 4A
). Note that the cross sections of the
as+s(C-H)2+3 and
as+s(C-D)2+3 signals are different, with the
as+s(C-D)2+3 signal being lower. For an isolated C-D bond, the intensity would be 1 /
2 lower than the equivalent C-H signal. However, for multiple bonds in a larger molecule, this relationship is complicated by neighboring molecules and difficult to determine. We have therefore simply used the band intensities uncorrected for cross section differences to provide a lower limit of the percent D31-PA. Figure 4B shows the association of percent D31-PA [
as+s(C-D)2+3 / [(
as+s(C-H)2+3 +
as+s(C-D)2+3)] of the total adipocyte lipids, incorporated into the adipocyte, with adipocyte size. A noticeable trend in the data presented in Fig. 4B is that there are no adipocytes with sizes greater than
7,000 µm2 that give rise to percent D31-PA values of less than 25%, whereas the inverse is true for adipocytes with sizes less than
7,000 µm2. Thus, these data suggest a general trend in which larger adipocytes contain higher concentrations of percent D31-PA, up to a limiting concentration of 30–35%.
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as(C-D)2+3 signal intensity distribution is shown in Fig. 5B. As expected, we observe localization of the
as(C-D)2+3 signal with high intensity at the adipocyte; however, we also find that this signal illuminates the PC-3 cells (Fig. 5B). Because the only source of
as(C-D)2+3 signal in the PC-3 cells is D31-PA released by the adipocytes, these data suggest translocation of D31-PA from the adipocytes into the PC-3 cells.
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as(C-D)2+3 signal in the PC-3 cells is due to background
as(C-D)2+3 absorption from stroma cells that may reside beneath the PC-3 cells. Baseline corrected FTIR spectra, in the frequency range comprising C-D absorbencies, were extracted from PC-3 cells (1–4) and extracellular locations (Sub 1–3) in close proximity to the PC-3 cells (Fig. 5C). The integrated areas of the C-D peaks are also displayed for these spectra in Fig. 5C. In Fig. 5Aii, we observe that the extracellular substrate adjacent to PC-3 cells 3 and 4 is optically absent of stroma cells. A spectrum was taken from this background location, as shown in Fig. 5B, and presented in Fig. 5C, Sub 1. This raw spectrum of the substrate reveals that the C-D peaks cannot be resolved from spectral noise (Fig. 5C). However, the spectrum of PC-3 cells 3 and 4 (Fig. 5C), which is in close proximity to this blank substrate (see optical image in Fig. 5A), gives rise to two prominent C-D peaks, demonstrating the translocation of D31-PA from the adipocyte into the PC-3 cell. A similar case is also found for PC-3 cell 1 (Fig. 5C), which displays two C-D peaks that are well resolved from spectral noise and whose adjacent background spectrum (Sub 3) displays no significant C-D absorbencies that can be resolved from spectral noise. For PC-3 cell 2, we find that this cell resides on a substrate/stroma cell background that gives rise to C-D peaks with appreciable absorbencies of 0.33 and 0.17 cm–1 (Fig. 5C). Nevertheless, even after subtraction of this background, the C-D peak areas in the spectrum of PC-3 cell 2 provide values of 0.64 and 0.33 cm–1. It is interesting to note that the intensity of C-D signal from the PC-3 cells is in the order of PC-3 cell 4 < cell 3 and cell 1 < cell 2, which correlates with their distances from the adipocytes (Fig. 5A).
| DISCUSSION |
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Prior to investigations concerning the uptake of FAs from adipocytes into PC-3 cells, issues concerning sample preparation were addressed. This was found to be a necessity, because delocalization/bleeding of lipid molecules from adipocytes in our adipocyte-PC-3 cell coculture system could arise in false-positive results concerning PC-3 uptake of adipocyte-derived D31-PA. Moreover, we found that in the absence of chemical fixation, the structured intracellular lipid droplets would collapse into a lipid deposit (Fig. 1B), and many adipocytes would lose a large number of lipid molecules (readily observed through optical microscopy), perhaps through air-oxidation of volatile unsaturated FAs. Fixation preserves not only the morphological information but also the spatial localization of the subcellular biomolecules of the adipocyte-PC-3 cell coculture system. Chemical fixation was chosen in preference to flash freezing and freeze drying for reasons outlined previously by our group (18).
The use of OsO4 as a postfixative for preserving the localization of lipid droplets in adipocytes is well documented (26–28). However, a disadvantage of OsO4 fixation is that the OsO3 by-product following OsO4 reaction with the lipid (Fig. 1E) can dissociate to form OsO2, which is a black, insoluble precipitate (29). Unbound OsO3 should be washed away during the Sorensen's buffer washing step and dehydration steps before CPD. However, even following these extensive washing steps, we obtain some darkening of the adipocytes as a result of OsO2 precipitation, as observed for the adipocytes in Figs. 2A and 5A. Although the amount of OsO2 precipitate is different between these adipocytes, this does not influence our FTIR analysis, because OsO2 does not absorb IR light in the mid-IR spectral region.
Following our assessment of adipocyte fixation, we evaluated, for the first time, the molecular information that could be derived from the adipocyte through FTIR microspectroscopy. As expected, high lipid hydrocarbon and lipid ester
s(C = O) signals (Fig. 2C, D, respectively) were measured from the adipocytes, compared with the stroma cells, and an increase in contrast was observed between adipocytes and stroma cells using the lipid ester
s(C=O) signal. Together, these results highlight that a high proportion of the adipocyte volume is lipid; in fact, Frühbeck et al. (15) have reported it to be
90%. The high contrast observed between adipocytes and stroma cells, using the lipid ester
s(C = O) intensity distribution, arises because most of the adipocyte lipid is in the form of a concentrated store of TAG, with each TAG comprising three fatty acyl chains bonded via an ester moiety to a glycerol backbone.
The identification of intracellular D31-PA hot spots in the FTIR image of adipocyte 1 through the intensity distribution of
as(C-D)2+3:lipid hydrocarbon signal (Fig. 2Fiii) may have significance in relation to adipocyte metabolism. Free FA (FFA) reesterification in adipocytes refers to the cycle of TAG hydrolysis for FFA liberation, its release from the adipocyte into the extracellular environment, and then reabsorption into the adipocyte for reesterification and TAG synthesis (22). Edens, Leibel, and Hirsch (22) have reported that this loop requires functionalized compartmentalization of FFA within the adipocyte, which prevents access of lipolytically derived FFA to the enzymes of glycerolipid synthesis. Thus, the hot spots of high
as(C-D)2+3 signal could correspond to TAG as well as FFA-compartmentalized D31-PA in adipocyte 1 (Fig. 2Fiii). In fact, the FFA:TAG image in Fig. 3B for adipocyte 1 can provide information concerning the molecular state of these D31-PA-rich locations. The D31-PA hot spots in Fig. 2Fiii overlapping with the TAG-rich locations in Fig. 3Bi may correspond to D31-PA as TAG, whereas the D31-PA hot spots overlapping with FFA-rich locations may correspond to D31-PA in the form of FFAs. However, further processing of other adipocytes analyzed by FTIR microspectroscopy (data not shown) revealed that these discrete subcellular locations of highly concentrated D31-PA were not observed for all adipocytes.
The localization of an isotopically labeled FA in a single adipocyte was also studied by Kleinfeld, Kampf, and Lechene (30) through multi-imaging mass spectrometry (MIMS). However, the adipocytes used in that study (30) were removed from media and dried under argon, which may have negative effects on the subcellular localization and structural integrity of the delicate lipid droplets. Although relative quantifications of a labeled FA within the adipocyte cytosol and lipid droplets were approximated using partition coefficients, it is unclear from their work (30) how this information was obtained, because direct MIMS analysis of the adipocyte interior must be accessed through freeze fracture or depth profiling. In contrast, FTIR microspectroscopy is less invasive, because it does not require fracture to access intracellular information and fixation can be applied to preserve the structural and spatial integrity of the lipid droplets. Another system that can be used to image the incorporation of specific lipids into the droplets of adipocytes is polyene tagging of lipids with two-photon excitation fluorescence microscopy (31). Compared with FTIR microspectroscopy, this system can provide better spatial resolution for localizing tagged lipids to subcellular compartments; however, saturated lipids such as palmitic acid cannot be studied using the fluorescence technique.
The quantitative study of specific FA uptake into an adipocyte and its influence on cell size is of particular interest in oncology because it has been reported that the concentration of unsaturated FAs in adipose tissue in breast cancer patients was in negative correlation with adipocyte size, whereas this correlation was reversed in lung cancer patients (24). In the present in vitro study, a scatterplot (Fig. 4B) indicated that the larger bone marrow-derived adipocytes stored a greater amount of D31-PA relative to nonisotopically labeled FAs. The data presented in Fig. 4B demonstrate that the uptake of a specific FA as a function of adipocyte size can be carried out without lipid extraction or adipocyte isolation and size exclusion from whole single adipocytes, in vitro, using FTIR microspectroscopy. In the context of adipocyte-tumor cell associations, the FTIR methodology reported here can be used to correlate the percentage of incorporation of a specific FA by the adipocyte with its size when these adipocytes are cocultured with and without tumor cells.
However, a limitation of this technique is associated with FA chain modification as a result of desaturase activity. If a significant amount of incorporated D31-PA is desaturated and stored as TAG in the adipocytes, then this situation would cause a reduction in the
as+s(C-D)2 signal. Chain elongation will not affect the correlation between the amount of D31-PA incorporated into the adipocyte and its size, because the number of C-D2 moieties will remain the same.
The final part of our work studied the translocation of D31-PA from adipocytes into PC-3 cells. To date, there are only a few publications in the current literature that report on an upregulation of lipids in tumor cells in the presence of adipocytes (9, 13). However, one cannot assume, without lipid-tracing experiments, that the observed upregulation of these cytoplasmic lipid droplets in the tumor cell is due to lipid translocation from adipocytes into the tumor cell. This is because tumor cells are known to upregulate lipid biosynthesis de novo through the overexpression of lipogenic enzymes, which have been reported to be induced by growth factors and hormones (32, 33). It is known that adipocytes release a range of growth factors (15), and these may stimulate the tumor cell to express lipogenic enzymes for de novo lipid biosynthesis. The data presented in Fig. 5 provide the first direct experimental evidence that the upregulation of lipid droplets in PC-3 cells, when cocultured with adipocytes, is at least in part due to FA translocation between the adipocyte and the PC-3 cell.
Although the translocation of D31-PA from the adipocyte into the PC-3 cell has been unequivocally demonstrated in this study using isotopically labeled D31-PA as a tracer, it is not known whether this translocation is mediated through direct PC-3 cell-adipocyte contact or via uptake from the incubation medium following FFA release from the adipocyte. In light of the data presented here, the high C-D signal from PC-3 cell 2, which resides between two adipocytes (Adp. 2 and 3, Fig. 5Aii), may have capitalized on both processes for D31-PA uptake; whereas PC-3 cell 4, at 88 µm from adipocyte 1 and 75 µm from adipocyte 3, furthest away from all of the adipocytes within the field of view, may have only incorporated released D31-PA from the media, giving rise to a lower C-D signal than cell 2. These hypotheses are currently being investigated in our laboratory.
The in vitro system described here provides a foundation for further investigations aimed at determining the specific FAs that may be incorporated/preferred by tumor cells (as well as associated mechanisms of uptake) and their biological consequences. In a wider context, this molecular-based assay can complement and provide support for epidermiological findings correlating dietary fat with cancer risks; Dennis et al. (5) have highlighted inconsistencies in these studies with respect to measurement differences for estimating service sizes (definitions of food groups or instructions provided to responders) and reporting methods (questionnaire platforms).
| CONCLUSION |
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| ACKNOWLEDGMENTS |
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Manuscript received March 19, 2007 and in revised form April 24, 2007.
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