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[Cancer Research 62, 1394-1400, March 1, 2002]
© 2002 American Association for Cancer Research

Experimental Therapeutics

Nuclear Magnetic Resonance-visible Lipids Induced by Cationic Lipophilic Chemotherapeutic Agents Are Accompanied by Increased Lipid Droplet Formation and Damaged Mitochondria1

Edward J. Delikatny2, Wendy A. Cooper, Susan Brammah, Nalayini Sathasivam and Darryl C. Rideout

Department of Cancer Medicine, The University of Sydney, New South Wales 2006, Australia [E. J. D., W. A. C., N. S.]; Central Sydney Area Health Service Electron Microscopy Unit, Concord Hospital, Concord, New South Wales 2139, Australia [S. B.]; and Darryl Rideout Structural Bioinformatics, Inc., San Diego, California [D. C. R.]


   ABSTRACT
Top
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Proton nuclear magnetic resonance (NMR) spectroscopy, histological lipid staining, and electron microscopy were used to assess the biochemical and structural changes induced by treating the cultured human breast cell line HBL-100 with the cationic lipophilic phosphonium salts p-(triphenylphosphoniummethyl) benzaldehyde chloride (drug A) and [4-(hydrazinocarboxy)-1-butyl] tris-(4-dimethylaminophenyl) phosphonium chloride (drug B). The major biochemical change detected by 1H NMR in drug-treated cells was a significant time- and concentration-dependent increase in lipid acyl chain resonances arising from mobile lipids. The amount of NMR-visible lipid strongly correlated with morphometric measurements of oil red O-staining lipid detected in the cytoplasm by light microscopy. Ultrastructural investigations revealed substantial damage to mitochondria and the progressive development of lipid droplets accompanied by end-stage autophagic vacuoles, in the form of densely staining myelinoid bodies, after treatment of HBL-100 cells with drug B at the IC50. No apparent increase in acid phosphatase activity was observed using electron microscopy, indicating that the accumulation of phospholipids in myelinoid bodies may result from substrate inundation of the lysosome, rather than increased lysosomal activity. These results indicate a potential role for lysosomal lipid catabolism in the formation of NMR-visible lipids in models of cytotoxic insult.


   INTRODUCTION
Top
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Tetraphenylphosphonium-based CLPS3 are novel anticancer agents with potential utility in the treatment of neoplasia. TPP is the parent compound of a series of CLPS, including drug A and the cationic acylhydrazine drug B, which have been shown to selectively inhibit the growth of cell lines derived from a wide variety of carcinomas (breast, colon, pancreas, bladder, and hypopharynx) relative to untransformed cell lines in vitro [CV-1 monkey kidney epithelial cells; IEC-18 rat ileal epithelial cells (1 , 2) ]. These drugs are thought to accumulate intracellularly as a function of membrane potential because collapsing the membrane potential with either valinomycin or high extracellular potassium concentrations reduces both the accumulation and the cytotoxicity of these agents (1 , 2) . The high negative plasma membrane potentials characteristic of neoplastic cells are believed to account for the selective accumulation and toxicity of CLPS and other cationic lipophilic compounds against malignant cells (3, 4, 5) .

Studies of various cationic lipophilic compounds suggest that mitochondrial toxicity provides the basis for the cytotoxicity of these compounds. Inhibition of mitochondrial respiration has been demonstrated in studies of TPP (2) and other lipophilic cationic compounds, such as MKT-077 (5 , 6) and rhodamine 123 (4) . Loss of ATP and phosphocreatine and decreases in intracellular pH have also been shown to accompany treatment with cationic lipophilic compounds (7 , 8) . It is thought that CLPS such as drugs A, B, and TPP as well as rhodamine 123 all have a similar mechanism of action because sensitivity profiles determined with a fixed set of cultured cells are closely similar for these agents (2) . Although loss of respiratory control has been suggested as the main mechanism of toxicity of lipophilic cationic agents, this may not be the sole cause for cell death. Damage to the mitochondria can cause changes in mitochondrial permeability and the release of apoptotic factors that would ultimately result in cell death (9 , 10) . Indeed, the full range of biochemical effects these drugs have on cancer cells has not been fully elucidated, and, consequently, it is currently not possible to unequivocally identify the final mediators of CLPS-induced cell death.

NMR studies of human breast cancer cells treated with TPP have revealed increases in NMR-visible or mobile lipids that are not readily explained by mitochondrial respiratory enzyme inhibition (11, 12, 13) . The drug-induced 1H NMR spectral changes were independent of cellular phenotype because similar concentration-dependent increases in lipid resonances were demonstrated in both the highly tumourigenic DU4475 breast carcinoma and the transformed HBL-100 human breast cell lines. However, the early and persistent detection of these lipid signals in response to TPP suggests that these alterations in lipid pools may reflect some of the primary cellular responses prior to or leading to cell death. Consequently, the aim of the current studies was to use histochemical staining and electron microscopy to identify subcellular alterations that give rise to the 1H NMR-visible lipids. These observations shed light on the ongoing controversy over the origins of NMR-visible lipid signals by demonstrating cytoplasmic inclusions of neutral lipids accompanied by evidence of lysosomal activity in this model system of cytotoxic insult.


   MATERIALS AND METHODS
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Cell Culture.
Human HBL-100 breast cells were grown as described previously (11 , 13) in RPMI-1640 supplemented with 10% (v/v) fetal bovine serum (batch number 81012053; Trace Biosciences), 2 mM L-glutamine, gentamicin (0.1% v/v; 40,000 units/ml), and 250 units/liter human insulin and buffered with 26.2 mM sodium bicarbonate using standard culture conditions of 37°C and 5% CO2 in air. HBL-100 cells grew as an adherent monolayer with a doubling time of approximately 40 h. In all experiments, HBL-100 cells were inoculated at a concentration of 2.5 x 105 cells/ml and treated with drug A or B (or equal volumes of PBS for controls) 5 h after seeding. Cells were treated at concentrations of drug A (300 or 10 µM) or drug B (10 and 2 µM) that represents the IC50 and IC10 for each drug, respectively (11) . Cells were harvested and examined using various techniques after 24, 48, or 72 h of drug exposure.

Drugs.
Two CLPS were used in this study, a cationic aldehyde, drug A (Mr = 417) and a cationic acylhydrazine, drug B (Mr = 542). Before each experiment, fresh stock at concentrations of 10 mM were prepared in PBS.

NMR.
HBL-100 cells grown in 175-cm2 tissue culture flasks were harvested by trypsinization, counted, and washed three times in 1 ml of PBS/deuterated water (D2O; pH 7.2). The pellet was resuspended in PBS/D2O to a volume of 400–500 µl and transferred to a 5-mm-diameter NMR tube. PABA (60 µl at 10 mM) was placed in a 2-mm coaxial capillary insert as an external standard. Cell counts and viability were assessed by trypan blue exclusion during sample preparation. An average of 5.9 ± 0.6 x 107 cells were used for the NMR dose-response experiments, and an average of 4.9 ± 0.4 x 107 cells for the NMR time-course experiments. One-dimensional NMR spectra were obtained at 37°C on samples spun at 20 Hz on a Bruker AM 360 NMR spectrometer as described previously (12 , 13) . Briefly, the standard acquisition parameters used were a spectral width of 10 ppm, relaxation delay of 2 s including a 1-s presaturation of the residual water signal, a total repetition time of 3.4 s (T1 of lipid methylene resonance ~500–650 ms), a 90° pulse length of 5.5–6.1 µs, and 128 8K transients acquired. Before Fourier transformation, a 3-Hz line broadening was applied, the spectra were phased, and the baseline was corrected using a fourth-order polynomial. Changes in lipid resonances were quantified by measuring peak heights or areas relative to the aromatic resonances of the external standard PABA. Areas were measured using standard Bruker integration routines or by fitting the methyl, methylene, and PABA regions of the spectra to a sum of Gaussian lines using MacNUTS (Acorn NMR, Inc., Livermore, CA). Peak heights or areas were corrected for minor variations in the volume contained within the NMR tube as well as for differences in cell number. For each condition, at least three independent experiments were performed.

Lipid Quantification by Light Microscopy.
Cells were seeded into 8-chamber tissue culture microscope slides, treated with drug A or B at the IC10 or IC50, and incubated under standard culture conditions for 48 h. The medium was removed, and the slides were air-fixed for 2–3 h, stained with hematoxylin and oil red O, and mounted with an aqueous mountant. Random fields from four independently prepared microscopic slides were photographed at x1000 magnification, and the accumulation of lipid droplets was quantified morphometrically (14) . A point-counting grid was randomly superimposed on projected 35 mm photographic transparencies of hematoxylin and oil red O-stained cells. The spacing of grid points (d), was chosen so that d2 was greater than the area of the largest profile (in this case a cell) to ensure that no more than one point could fall on the same profile. Due to the substantial size difference between whole cells and individual lipid droplets, two lattices of differing point density were superimposed to create a coherent double lattice system with 16 fine points to each coarse grid point. For each sample, 6–10 test areas were counted (>=200 lipid droplet point counts), and fractional areas were determined by point count ratios. Statistical significance for NMR and morphometric data was determined using a two-sided unpaired Student’s t test. All data are reported as mean ± SEM.

Transmission Electron Microscopy.
Cells were seeded, treated, and incubated as described above. They were removed from the culture flasks using a cell scraper and centrifuged at 3000 rpm for 15 min. The resultant pellet was resuspended in 2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer (pH 7.4) and fixed at 4°C for 1 h. The cells were then pelleted using 10% BSA and gelled with 2.5% glutaraldehyde for 4–5 h at 4°C (15) . They were then processed routinely, as tissue blocks, for transmission electron microscopy. Ultrathin sections were stained with uranyl acetate followed by lead citrate and examined in a Philips 410LS transmission electron microscope. For the identification of acid phosphatase activity, cells were fixed as described before and stained using a modified Gomori method (16) . The cell pellet was placed in 0.1 M sodium cacodylate containing 7% (w/v) sucrose, resuspended, and stored overnight at 4°C. After centrifugation, the cells were rinsed in 50 mM sodium acetate buffer with 7% sucrose and incubated in 50 mM sodium acetate buffer containing 10 mM disodium-ß-glycerophosphate and 3.3 mM lead nitrate for 30 min at 37°C. After centrifugation, the cells were rinsed in 0.1 M sodium cacodylate with 7% sucrose, pelleted as described before in BSA, and processed for transmission electron microscopy. The ultrathin sections were viewed unstained.


   RESULTS
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Drug-induced Increase in NMR-visible Lipid.
HBL-100 cells were treated with drugs A and B at the IC50 or at the IC10, and 1H NMR spectra were obtained on harvested cells 48 h after treatment. The 1H NMR spectra of untreated cells (Fig. 1a)Citation typically exhibit a number of broad signal bands from proteins superimposed with a number of narrower resonances from amino acids, creatines, carbohydrates, lactate, and choline-containing metabolites, but essentially contain no lipid signals. Spectra of HBL-100 cells treated with drug A or B at the IC50 (300 and 10 µM, respectively; Fig. 1, b and cCitation ) demonstrate substantial increases in NMR-visible lipids, particularly the lipid acyl chain methylene (CH2) resonance at 1.3 ppm, lipid methyl groups (CH3) at 0.9 ppm, fatty acyl chain ß-methylene at 1.7 ppm, and olefinic groups (CH-CH) at 5.35 ppm. The contribution from lactate and threonine methyl groups to the resonance at 1.3 ppm was minimized by the three washes of the cells during harvest. Detailed resonance assignments for human breast cells have been presented previously (12 , 13) .



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  		Fig. 1. One-dimensional proton NMR spectra of HBL-100 human breast cells treated with (a) PBS (control cells), (b) 300 µM drug A (IC50) for 48 h, and (c) 10 µM drug B (IC50) for 48 h.

 
To quantify drug-induced changes, the intensities or areas of lipid resonances were measured relative to signals arising from the external standard, PABA, at 6.8 and 7.8 ppm. The peak height and area ratios of the lipid methylene (1.3 ppm) and methyl (0.9 ppm) relative to PABA increased significantly in HBL-100 cells treated with drug A or B at the IC50 compared with control cells (Fig. 2)Citation . As expected, the increase in the lipid methylene signal was far greater than the corresponding increase in the methyl signal. In fatty acids, the ratio of methylene:methyl protons contributing to lipid signals is approximately constant (e.g., 22:3 and 18:3 for palmitic and oleic acids, respectively); the greater increase in methylene resonance occurs as these lipid resonances emerge from the background amino acid signals. No statistically significant changes were observed after treatment with drug A or B at the IC10 (10 µM for drug A and 2 µM for drug B).



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  		Fig. 2. Ratio of methylene (CH2, 1.3 ppm) and methyl (CH3, 0.9 ppm) intensities relative to the external standard PABA at 6.8 and 7.8 ppm. Data represent the mean ± SEM of at least three independent experiments.

 
HBL-100 cells were treated with drug B at the IC50 over a 3-day period, and NMR spectra were obtained every 24 h. Proton NMR spectra of HBL-100 cells demonstrated a time-dependent increase in both the peak height and the peak area of the methylene resonance at 1.3 ppm relative to either the methyl resonance at 0.9 ppm (data not shown) or the PABA resonance at 6.8 and 7.8 ppm (Fig. 3)Citation . At 48 and 72 h, these ratios were significantly greater in the drug-treated cells relative to control (P < 0.05). Moreover, the time-dependent increase in the methylene signal ratios in drug B-treated cells was significant between 24 and 72 h (P < 0.05). These changes were also significant when the methylene and PABA resonances were fit to a sum of Gaussian lines. No significant changes in the linewidth of the methyl resonance were observed under any treatment conditions for either dose-response or time course conditions.



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  		Fig. 3. Time dependence of the CH2:PABA ratio for control ({blacksquare}) and HBL-100 cells treated with drug B at the IC50 ({square}). Data represent the mean ± SEM of at least four independent experiments. Statistical significance is denoted by ** (P < 0.01) and * (P < 0.05, Student’s t test).

 
Drug-induced Accumulation of Cytoplasmic Lipid.
Light microscopic lipid staining techniques were undertaken to further investigate the nature and cellular location of the mobile lipid detected by NMR. HBL-100 cells treated with drugs A or B displayed a concentration-dependent increase in oil red O lipid staining in cytoplasmic droplets of varying sizes (Fig. 4, A and B)Citation . In contrast, control HBL-100 cells displayed very few small scattered cytoplasmic lipid droplets that were also not detectable in all cells (Fig. 4C)Citation .



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  		Fig. 4. Light micrographs of HBL-100 cells treated with (A) 300 µM drug A (IC50), (B) 10 µM drug B (IC50), and (C) PBS (control) photographed at a magnification of x1000. Cells were grown on multichamber tissue culture microscope slides and stained with oil red O and hematoxylin.

 
Morphometric point counting techniques were used to quantify accumulations of oil red O-staining lipid observed by light microscopy. The volume fraction of lipid droplets present in either whole cells or the cytoplasm (Fig. 5)Citation was significantly greater in HBL-100 cells treated with drugs A and B at the IC50 (P < 0.01). At the IC10, the lipid volume fraction increased to a lesser extent but was still significantly greater than controls in both the cytoplasmic and whole cell measurement (P < 0.01). HBL-100 cells treated with either drug A or B at the IC50 accumulated oil red O-staining lipids to fill 25% and 13% of the cytoplasm, respectively, compared with only 5% lipid in the cytoplasm of controls (P < 0.01). A comparison of morphometric and spectroscopic data demonstrated that the concentration-dependent increase in cytoplasmic lipids after treatment with either drug A or B correlated strongly with lipid detected by one-dimensional 1H NMR (Fig. 6)Citation . Positive controls prepared simultaneously from frozen sections of rat fat tissue were stained intensely red, confirming the lipidic nature of substances taking up the stain. Negative controls prepared by placing slides of rat fat tissue or HBL-100 cells in a delipidizing solution did not show any staining with oil red O (data not shown).



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  		Fig. 5. Volume fraction of oil red O-positive lipid in the whole cell and in the cytoplasm of HBL-100 cells treated with drugs A and B (IC50 and IC10). Error bars, SEM. Statistical significance is denoted by ** (P < 0.01, Student’s t test).

 


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  		Fig. 6. Correlation of the two methods of measuring lipid accumulation in HBL-100 cells: point counting of oil red O droplets to determine the volume fraction of lipid versus the CH2:PABA peak height ratio from one-dimensional 1H NMR. The correlation constant, R, = 0.99.

 
Drug-induced Changes Observed using Electron Microscopy.
In conjunction with the NMR time course experiments, HBL-100 cells were treated with drug B at 10 µM (the IC50 at 48 h) over a 3-day period, and electron microscopy specimens were obtained every 24 h. In treated cells at 24 h (Fig. 7A)Citation , mitochondrial changes were observed, including overall swelling and dense matrix condensations. Mitochondrial cristae were lost or disordered, and some were pushed to the edges forming rings or doughnut shapes, consistent with previous observations on cationic lipophilic compounds (2 , 17) . Some autophagic vacuoles were observed, and occasionally the presence of myelinoid bodies was noted. Increased numbers of lipid droplets were present, with groups of small droplets often observed. By 48 h (Fig. 7B)Citation , mitochondrial swelling and matrix condensation had increased, as had the number of lipid droplets that were often found grouped with damaged mitochondria. Small- to medium-sized myelinoid bodies were seen. In some cells, mitochondrial damage was extensive, with only mitochondrial double membrane ghost remnants present along with the presence of numerous perinuclear myelinoid bodies (Fig. 7C)Citation . Evidence of necrosis was present with cells showing damaged or missing plasma membranes, swollen endoplasmic reticulum, and proteinaceous debris. By 72 h, all cells appeared to be necrotic. Myelinoid bodies and especially lipid droplets still appeared to be present.



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  		Fig. 7. A, electron micrograph of HBL-100 cells treated with drug B (10 µM, IC50) for 24 h. Mitochondria show swelling, disordered cristae (including one doughnut form), and flocculent condensations in the matrix. A lipid droplet is also present. Bar, 1 µM. x17,000. B, electron micrograph of HBL-100 cells treated with drug B (10 µM, IC50) for 48 h. Mitochondrial damage is increased; marked swelling, dense condensations in the matrix, and disorganization of cristae are evident. Numerous lipid droplets are present in the same region of the cell as the damaged mitochondria. x17,000. C, electron micrograph of HBL-100 cells treated with drug B (10 µM, IC50) for 48 h. Numerous myelinoid bodies are seen in one cell, and extensively damaged mitochondria are seen in another. x7,425. D, electron micrograph of control HBL-100 cells at 48 h. Viable elongated cells with numerous normal mitochondria. x7,425.

 
Ultrastructural observations of control HBL-100 cells showed viable elongated cells with no significant abnormalities (Fig. 7D)Citation . Numerous mitochondria were present with readily discernible cristae. Although some lipid droplets were present in the control at 24 h, by 48 h these had virtually disappeared. At 72 h, the mitochondria appeared smaller, and some accumulation of debris was present, but the cells were generally viable and healthy.

In control cells, staining for acid phosphatase, a lysosomal enzyme marker, showed activity mostly in lysosomes as well as some activity in the rough endoplasmic reticulum and Golgi zones (data not shown). Lysosomal activity was variable from cell to cell. In drug B-treated cells, activity was observed in primary lysosomes and also in autophagic vacuoles at 24 h that persisted until 72 hours. Myelinoid bodies showed little activity. There was no specific association of lysosomal activity with the damaged mitochondria or lipid droplets, but all three structures appeared in the same region of the cells. There was no apparent increase in overall acid phosphatase activity in the treated cells when compared with controls (data not shown).


   DISCUSSION
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
The results presented in this study demonstrate that the cytotoxic cationic lipophilic compounds, drugs A and B, induced an increase in NMR-visible lipids in human HBL-100 breast cells that was also seen as an accumulation of oil red O-positive lipid droplets. Concurrently, electron microscopy demonstrated the formation of lipid droplets and myelinoid bodies accompanying mitochondrial damage and destruction.

A strong correlation was demonstrated between the increase in NMR-visible lipid and the accumulation of oil red O-positive cytoplasmic lipid (Fig. 6)Citation . Oil red O is predominantly a neutral lipid dye, staining nonpolar lipids red and some phospholipids pink (18) . These results suggest that the cytoplasmic droplets that accumulate in CLPS-treated cells are composed of triglyceride and other neutral lipids. This observation is confirmed by the appearance of cytoplasmic neutral lipid droplets on electron micrographs. It is possible that the myelinoid bodies may contribute to the oil red O-positive droplets observed in drug A- and drug B-treated cells. Myelinoid bodies are lysosomal bodies containing markedly osmiophilic membranes representing tertiary (end-stage) autophagic vacuoles and show a stacked, reticulated, or whorled arrangement of multilamellar lipid surrounded by a single membrane enclosure (19) . Myelinoid bodies observed in the hereditary lysosomal storage disorders Tay-Sachs and Niemann-Pick disease stain positively with oil red O (20) , as do those caused by lysosomal storage induced by 4,4'-diethylaminoethoxyhexestrol, an amphiphilic aromatic base (21) . However, the observation of a time-dependent increase in the NMR-visible lipid signal accompanied by an increasing number of lipid droplets in electron micrographs indicates that the lipid droplets are the primary source of the NMR-visible lipids.

Neutral lipid accumulations are a well-known indicator of cell stress (20) and can be induced in tumor cells cultured to high density (22, 23, 24) , exposed to experimental conditions of hypoxia (25) , or treated with chemotherapeutic agents (11, 12, 13 , 26, 27, 28, 29) . In models of cytotoxic insult, the accumulation of mobile lipids is dose dependent, generally occurring at concentrations below the IC50, with the absolute concentration depending on the drug and cell line. The location of NMR-visible triglyceride accumulations has been controversial, with some groups reporting a correlation between cytoplasmic lipid droplets and NMR-visible lipid (23 , 26 , 30 , 31) and other groups attributing the origins of mobile lipids to plasma membrane domains (reviewed in Ref. 29 ; see also Refs. 32 and 33 ). A recent study by Ferretti et al. (34) has proposed that both origins are possible, with minor lipid accumulations being plasma membrane related and larger lipid accumulations accumulating as cytoplasmic lipid droplets. The present study shows that the relatively large increase in methylene resonances observed accompanying cytotoxic insult is strongly linearly correlated to the increase in cytoplasmic lipid droplets. Moreover, for the first time, a possible functional source of mobile lipids is indicated by demonstrating the presence of autophagic vacuoles and thus implicating lysosomal metabolism in the formation of NMR-visible lipids in models of cytotoxic insult.

It is interesting to note that electron microscopy of HBL-100 cells showed none of the morphological changes characteristic of apoptosis, such as chromatin condensation or cell shrinkage, and instead showed death by necrosis. Whereas this evidence does not preclude the possibility of apoptotic cell death arising from CLPS, it does indicate that mobile lipid production may not be limited or specific to the apoptotic process, as has been proposed by others (35) , but may exist as a more general indicator of cell stress and death. This is in keeping with the work of Kuesel et al. (36) , who reported increased mobile lipids accompanying necrosis in human brain tissues.

The structural mitochondrial changes reported here confirm other reports of mitochondrial damage by cationic lipophilic compounds. Human pancreatic carcinoma cells treated with the cationic lipophilic agent MKT-077 developed enlarged mitochondria with abnormal cristae and dense matrix inclusions, whereas normal epithelial cells did not (5) . FaDu human hypopharyngeal carcinoma cells treated with TPP for 6 days (2) and L1210 murine leukemic cells treated with rhodamine-123 (17) also displayed damaged mitochondria with disrupted cristae, membrane blebbing (2) , distension, and the formation of doughnut-shaped structures (17) . Interestingly, the micrographs presented in these studies show structures resembling myelinoid bodies and intermediate myelinoid structures. Hypoxia, which also inhibits mitochondrial aerobic respiration, can induce structural damage in cellular organelles, including mitochondria, similar to that induced by CLPS (25) .

Many conditions causing sufficient increases in lysosomal substrates or decreases in lysosomal degradative functions can result in lysosomal phospholipid storage in the form of myelinoid bodies (37) . The formation of myelinoid bodies observed with CLPS treatment probably results from an oversupply of substrate from damaged mitochondria (19) because no overall increase in lysosomal function was observed using acid phosphatase staining. Substrate inundation of autophagic processes has also been used to explain myelinoid bodies observed in progressively degenerating Yoshida hepatomas (38) .

Lysosomal processing of damaged mitochondria would lead to the shunting of a large amount of lipid through the lysosome. Lysosomes contain a number of lipid catabolic enzymes including sphingomyelinase and phospholipases A1, A2, and C that would result in production of fatty acids, glycerol, and phosphodiesters (39) . Whereas fatty acids and glycerol are known to be exported from the lysosome, the fate of phosphodiesters such as glycerophosphocholine remains less certain (39) . Fatty acids would be available for triglyceride production; the release of glycerophosphocholine could result in the increases in this metabolite observed in breast cells treated with the parent compound TPP but not its derivatives, drugs A and B (11, 12, 13) . Autophagy of phagocytosed organelles has been shown to result in the formation of lipid droplets after periods of 24 h (40) . The hypothesis that triglyceride formation is at least partially mediated by lysosomal processing of lipids from damaged mitochondria in CLPS-treated cells is consistent with all the data and is further supported by preliminary data from our laboratory whereby TPP-induced lipid production can be inhibited by the lysosomal inhibitor chlorpromazine (41) .

It is possible that the fatty acid chains necessary for the production of neutral lipid droplets arise from uptake of external lipid sources, such as serum, or from decreased oxidation or de novo synthesis of fatty acids. The contribution of serum lipids or exogenous fatty acids has been shown to be important in the generation of NMR-visible lipids in activating immune cells and in myeloma cells (30 , 42 , 43) . On the other hand, Henke et al. (44) showed using 13C NMR that fatty acid synthesis was increased within 2 h of treatment in KB epithelial cells undergoing miltefosine-induced apoptosis. Similarly, treatment of the prostate cancer cell line LNCaP with androgens or retinoids resulted in a sustained increased level of fatty acid synthase and other lipogenic enzymes that was at least partially responsible for the accumulation of neutral lipid droplets (45 , 46) . This response was shown to be relatively specific because androgens cause accumulations of both triglycerides and cholesterol esters, whereas retinoic acid treatment leads only to the formation of triglycerides. In CLPS-treated cells, the relative contributions of these various pathways to the formation of NMR-visible lipids remain unknown, but investigations in our laboratory are currently under way to address these questions.

In the long term, it is important to know the degree to which these changes contribute to the overall toxicity—whether lipid accumulations contribute significantly to cytotoxicity or are solely a consequence of cellular degradation. Many malignant cells rely primarily on glycolysis as an energy source, and it has been shown that mitochondria are not necessary for tumorigenicity in animal models (47) . If this is indeed the case, then it is important to determine whether CLPS-induced mitochondrial destruction is a sufficient condition for cell death, and what contribution the metabolism of lipids and other breakdown material makes to the cytotoxic process. Furthermore, lysosomal and cytoplasmic lipid accumulations that are simply a secondary result of drug action may hamper the efficacy of these agents and lead to side effects in vivo. Finally, it is possible that the observed gross metabolic changes could be translated into the clinical setting, where drug efficacy within tumors could be monitored and predicted using in vivo 1H NMR.


   ACKNOWLEDGMENTS
 
We acknowledge the critical assessments of Dr. Thomas M. Jeitner, Wendy A. Bartier, and Prof. Martin Tattersall. We thank Songul Bekir (Department of Pathology, University of Sydney) and Prof. Peter Russell (Royal Prince Alfred Hospital Department of Anatomical Pathology) for histopathological advice and assistance and Dr. Edward J. Wills (Central Sydney Area Health Service Electron Microscopy Unit, Concord Hospital) for helping to interpret the electron microscopic findings. In addition, we thank Dr. Jillian Kril (Centre for Education and Research on Ageing, the University of Sydney) for advice on morphometry.


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

1 Supported by Australian National Health and Medical Research Council Grant 940630, the Leo and Jenny Leukaemia and Cancer Foundation, and NIH Grant R21 CA79718. This article is dedicated to the memory of Josephine Vandeleur. Back

2 To whom requests for reprints should be addressed. Present address: Department of Radiology, University of Pennsylvania Medical Center, Room B1, Stellar-Chance Building, 422 Curie Boulevard, Philadelphia, PA 19104-6140. Phone: (215) 898-1805; Fax: (215) 573-2113; E-mail: delikatn{at}oasis.rad.upenn.edu. Back

3 The abbreviations used are: CLPS, cationic lipophilic phosphonium salts; drug A, p-(triphenylphosphoniummethyl) benzaldehyde chloride; drug B, [4-(hydrazinocarboxy)-1-butyl] tris-(4-dimethylaminophenyl) phosphonium chloride; NMR, nuclear magnetic resonance; PABA, p-aminobenzoic acid; TPP, tetraphenylphosphonium chloride. Back

Received 7/27/01. Accepted 1/ 3/02.


   REFERENCES
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

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