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Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
Mol Nutr Food Res. Author manuscript; available in PMC 2010 August 25.
Published in final edited form as:
PMCID: PMC2928140

From delocalized lipophilic cations to hypoxia: Blocking tumor cell mitochondrial function leads to therapeutic gain with glycolytic inhibitors


An unexpected similarity between cancer and cardiac muscle cells in their sensitivity to anthracyclines and delocalized lipophilic cations (DLC) prompted a series of studies in which it was shown that the positive charge of these compounds is central to their selective accumulation and toxicity in these two distinct cell types. An initial finding to explain this phenomenon was that cancer and cardiac muscle cells exhibit high negative plasma membrane potentials resulting in increased uptake of these agents. However, the p-glycoprotein efflux pump was shown to be another factor underlying differential accumulation of these compounds, since it recognizes positively charged drugs and thereby actively reduces their intracellular concentrations. The delocalized positive charge and lipophilicity of DLCs leads to their retention and inhibition of ATP synthesis in mitochondria. Years later it was realized that cancer cells in the hypoxic portions of solid tumors were similar to those treated with DLCs in relying mainly on anaerobic metabolism for survival and could thus be targeted with a glycolytic inhibitor, 2-deoxy-d-glucose (2-DG). This hypothesis has lead to a Phase I clinical trial in which 2-DG is used to selectively kill the hypoxic tumor cell population which are resistant to standard chemotherapy or radiation.

Keywords: Anthracycline, Cancer, Delocalized lipohilic cation, Hypoxia, Mitochondria

1 Introduction

The impetus for this review dates back to the early 1970s when it became known that the antitumor agent, doxorubicin (Dox) a member of the anthracycline family of compounds, caused serious and sometimes fatal cardiomyopathy in a significant number of treated cancer patients [1, 2]. A fundamental question which arose was, why should a drug that affects actively dividing cancer cells also affect the heart which is comprised mainly of nondividing cells? Thus, it followed that the toxicity of Dox might be reflecting similarities in these very different cell types which could be investigated in vitro by measuring the relative accumulation and potency of this drug. In initial studies it was found that Dox (which is naturally fluorescent and thus easy to detect in live cells under microscopy) accumulated preferentially in the nucleus of cardiac muscle cells versus cocultured cardiac fibroblasts [3, 4]. Testing a number of other anthracyclines it was observed that those analogs which were positively charged at physiologic pH, daunorubicin, detorubicin, and rubidazone were shown to display similar preferential accumulation in cardiac muscle versus nonmuscle cells [5]. In contrast, a neutral anthracycline analog, AD32, accumulated equally in cardiac muscle versus nonmuscle cells [5]. Thus, a connection was made between the positive charge of anthracyclines and their preferential accumulation in cardiac-muscle cells (Fig. 1). It is of interest that a charged molecule can permeate the plasma membrane, however the mechanism for the diffusion of anthracyclines remains unclear. A possibility is that the charge is weak enough so that it does not interfere with the lipid solubility of these compounds but further investigation is required to clarify this issue.

Figure 1
Note that the NH2 group (A) on the sugar moiety of Dox, as well as daunorubicin, detorubicin, rubidazone is protonated at physiologic pH. On the other hand, Rho123 and other DLCs carry a delocalized positive charge (B) which does not interfere with their ...

This connection was later fortified by the finding that Rhodamine 123 (Rho123), a delocalized lipophilic cation (DLC), also preferentially accumulated and was retained in cardiac muscle versus nonmuscle cells [68]. Although Dox is an anthracycline and Rho123 is a member of the DLC family, the commonality of their preferential accumulation in cardiac cells led to the realization that the positive charge of these compounds at physiologic pH was a key chemical component responsible for this phenomenon (Fig. 1).

2 Rhodamine 123 and other DLCs preferentially accumulate in mitochondria of living cells

In the late 1970s, while investigating the staining of a surface antigen with a rhodamine-conjugated antibody in living cells, it was observed that so-called “snakelike organelles” were staining which were later shown to be mitochondria [9]. In a following report, it was found that a contaminant in the rhodamine preparation, identified as rhodamine 3B, was responsible for mitochondrial staining of living cells when used at a concentration of 0.5 μg/mL for 15 min [10]. Furthermore, two other rhodamines (123 and 6G) designated by Kodak as laser dyes, were observed to be supravital stains for mitochondria, whereas rhodamine B, 110, 116, and tetramethylrhodamine B were not [11]. Structure function analysis with these rhodamine analogs led to the observation that only those rhodamines that carried a delocalized positive charge were accumulating specifically in mitochondria [10, 11]. These rhodamine analogs are a subset of DLCs which contain an esterified carboxyl group and a positive charge over the molecule that does not interfere with the lipophilic nature of the dye [12]. They differ from the neutral rhodamine analogs which carry a free unesterified carboxyl group and are therefore amphoteric. Thus, the delocalized positive charge of esterified rhodamines allows these compounds to permeate through plasma as well as mitochondrial membranes.

Their intracellular localization is governed by the high negative transmembrane potential of mitochondria which attracts positively charged agents such as DLC, in accordance with the Nernst equation [13]. Staining of mitochondria by Rho123 was shown to be inhibited by dissipation of mitochondrial membrane potential further indicating that the negative charge in the mitochondrial inner membrane acts as an attractive force for positively charged agents [10, 11, 14]. In contrast, pretreatment of cells with nigericin which increases mitochondrial transmembrane potential resulted in increased Rho123 uptake and staining of this organelle [10, 15]. These data suggested that the uptake of lipophilic cations such as Rho123, could be used in living cells as a relative measure of their mitochondrial transmembrane potentials (Δψm) [10].

The fact that one group of compounds (the DLCs) localizes intracellularly in mitochondria while the anthracyclines accumulate in nuclei is most likely due to the overall size, lipophilicity, and nature of the DNA binding properties of each group, respectively. The free amino group in the glycosidic portion of the anthracyclines appears to be a key determinant for actual covalent bonding to nuclear DNA while the delocalized positive charge of the DLCs does not favor this kind of bonding to DNA. Thus, it appears that the nature of the positive charge (localized vs. delocalized) plays an important role in determining nuclear versus mitochondrial localization although the degree of lipophilicity has also been shown to affect the intracellular localization of analogs within each family of compounds [12].

3 Positively charged anthracyclines and delocalized lipophilic cations accumulate preferentially in carcinoma versus normal epithelial cells

The finding that preferential accumulation and toxicity of anthracyclines and DLCs in cardiac cells was related to the positive charge of these compounds, prompted experiments in which it was shown that these drugs selectively accumulate in, and are toxic to, a number of different carcinoma cell lines as compared to normal epithelial cell types [16]. These early studies led to a series of publications in which it was shown that Rho123 and other DLCs, exhibited anticancer activity both in vitro [17, 18] and in vivo [19]. Although the mechanism of toxicity induced by anthracyclines was different from that by Rho123 [18, 20], the data in these reports established that positive charge for both anthracyclines as well as DLCs was an important chemical component in the preferential accumulation of these different families of compounds in cardiac muscle and carcinoma cells as compared to cardiac nonmuscle and normal epithelial cells.

3.1 Increased plasma and mitochondrial membrane potentials are mechanisms for selective accumulation of anthracyclines and DLCs

A biophysical parameter that could account for increased uptake of positively charged compounds is increased transmembrane potentials at both plasma (Δψp) and mitochondrial (Δψm) membranes. Since all cells generate an electrical gradient which is negative on the inside of the cell, Δψp could act as an attractive force for intracellular accumulation of agents of this nature. The evidence obtained suggested that cardiac muscle as well as some carcinoma and leukemic cells exhibit higher Δψp as compared to cardiac nonmuscle, normal epithelial, and other cell types which influences their preferential accumulation of positively charged compounds [21]. In contrast to positively charged anthracyclines which are mainly affected by Δψp, intracellular accumulation of DLCs are influenced by the magnitude of both Δψp and Δψm. Using valinomycin to depolarize Δψm and high K to lower Δψp, it was shown that Δψm had a more profound effect than Δψp in determining the overall accumulation and retention of DLCs [22]. Thus, the difference in accumulation of Rho123 between carcinoma and normal cell types has been related to a more negative transmembrane potential of mitochondria extracted from carcinoma cells which results in the higher retention of DLCs in these cells [22].

Moreover, it has been suggested that increased Δψm appears to correlate with aggressive features of carcinoma cells in a model of gastric cancer [23]. In a recent study using several isogenic carcinoma cell lines which have differences in their Δψm, it was found that those cell lines with higher intrinsic Δψm exhibited enhanced tumorogenic properties, i.e., survival in low oxygen conditions by inhibiting hypoxia-induced apoptotic pathways, increased anchorage-independent growth ability as well as invasion of the basement membrane [23]. This study provides a rationale for the possible clinical application of DLCs since cancer cells with a more aggressive character may have higher Δψm and thereby preferentially accumulate and be more sensitive to these positively charged agents.

The mechanisms underlying the differences in Δψm between carcinoma and normal cells, however, remain unclear. It was suggested that the deficiencies in ATP synthase observed in cancer, as compared to normal cells, correlate with their higher Δψm [2431]. However, in reports where defects in mitochondrial ATP synthase was detected, electron transport deficiencies were also observed and furthermore Δψm was not directly measured [2531]. It is reasonable to presume, a priori, that reduction in electron transport chain (ETC) should lead to decreased Δψm. Thus, the effects of reduced ATP synthase activity in increasing Δψm would be negated by ETC interruption. This contention is supported by the findings that mutations in cancer mitochondria cause electrons to be retained in complexes I and III. This, in turn, facilitates the direct transfer of electrons onto molecular oxygen and thereby higher production of reactive oxygen species (ROS) which are shown to be involved with oncogenic transformation [32, 33]. Thus, it appears that mitochondrial deficiencies in cancer cells correlate with high ROS levels, but not necessarily with high proton gradient or increased Δψm. Moreover, the studies, in which Δψm was found to be higher in cancer cells, used DLC accumulation as a measure of Δψm rather than direct measurement of this parameter [2531]. Taken together, to date there is direct evidence supporting a link between the retention of positively charged compounds in tumor cells and increased Δψp [21] while the role of Δψm in this phenomenon remains to be further investigated.

3.2 p-Glycoprotein (p-gp) mediated multidrug resistance is an underlying mechanism for selective accumulation of DLCs

Another explanation for increased DLC retention in carcinoma mitochondria comes from the observation that most of the compounds recognized by cancer cells expressing the clinically important mechanism of p-gp mediated multi-drug resistance (MDR), are also positively charged. Juliano and Ling have shown that tumor cells which develop the MDR phenotype, earlier described by Riehm and Biedler [34], and are typified by lower drug accumulation, have increased levels of a 170-kDa plasma membrane glycoprotein [35]. Extensive work after this initial finding revealed specific MDR genes encoding this glycoprotein which conferred cellular drug resistance [36]. It is known that the p-gp acts as an efflux pump for a variety of structurally unrelated compounds, which accounts for their lower accumulation in MDR+ cells [34, 37]. To reiterate, our observations and results [21, 38] confirmed by others [3946] indicate that most compounds effluxed by MDR+ cells are positively charged.

Further evidence to support this idea comes from studies in which a single negatively charged amino acid in the MDR transporter, Mdfa, of Escherichia coli was found to account for recognition of positively charged compounds [47]. Additionally, it was demonstrated that when this amino acid was substituted with a positively charged one, recognition of positively charged drugs was abolished but the transporter still functioned for other uncharged compounds, i.e., chloramphenicol [47]. Although this result was reported in prokaryotic cells, the authors speculated that similar recognition for p-gp mediated transporters in eukaryotic cells may exist due to single amino acid residues. This work is particularly pertinent for supporting the hypothesis that the positive charge of a number of different families of compounds, including DLCs, is an important chemical characterestic for determining whether a drug will be recognized by p-gp mediated MDR in a variety of human and mouse cells [47].

Moreover, using a series of nine anilinoacridines, Baguley and Ferguson [48] have found that strongly basic or positively charged analogs of this group of compounds showed the greatest degree of crossresistance in P388/ADR resistant cells, which further supports this concept. It was also reported that the higher the pKa of the amino sugar of a selected number of anthracyclines studied, the greater intracellular accumulation in drug sensitive HL-60 leukemia cells [49]. Beck and coworkers, in a series of papers have shown the necessity of cationic charge for compounds to act as MDR modulators [40, 41]. Lampidis et al. using a selected series of anthracycline analogs in which lipophilicity and charge were altered found the following: (i) positively charged anthracyclines as compared to their neutral counterparts are better recognized by MDR+ cells; (ii) with increasing lipophilicity charge becomes less important for MDR recognition; (iii) resistance to anthracylines can be reduced >1000-fold with an analog that does not contain a protonatable nitrogen and is highly lipophilic; and (iv) highly lipophilic analogs regardless of charge can act as modulators of p-gp [50]. This latter finding suggests that at a high enough lipophilicity either drug transport overcomes the speed of the efflux pump and or that highly lipophilic compounds nonspecifically bind to or interfere with the effluxing function of p-gp and thereby act as self-modulators.

To define more precisely the influence that charge and lipophilicity have on MDR recognition, a series of simple aromatic (pyridinium) and nonaromatic (guanidinium) cations were designed and synthesized which differ in lipophilicity by stepwise addition of single alkyl groups [51]. Using these simple compounds it was found that an aromatic ring and a minimal degree of lipophilicity (log -P > 1 = at least five alkyl groups) in addition to positive charge, were required for p-gp mediated MDR recognition [51]. With increasing lipophilicity (above five alkyl groups) the resistance ratio between MDR- and MDR+ cells increased as a function of increasing chain length [51]. This study, which is further supported by other labs, emphasizes the importance of positive charge but also defines a minimal degree of lipophilicity necessary for MDR recognition [5155].

It is clear that the chemical structure of Rho123 and other DLCs is suitable for recognition by the p-gp efflux pump [56] which suggested that the selectivity we had originally observed between carcinoma versus normal epithelial cells with Rho123 [6, 16] could, at least in part, be due to the presence of the MDR1 gene product. Indeed, the normal epithelial cell line, CV-1, which was originally used as a model for cationic drug selectivity between normal and tumor cell lines, was later found to intrinsically express MDR1 [57] and display reduced uptake of a number of “MDR recognizable drugs,” i.e., Dox, vinblastine, taxol, as well as Rho 123 [57]. In this report, it was also shown that the classic MDR modulator, verapamil, reverses the resistance to each of these agents in CV-1 cells [57]. However, not all normal cells are known to express MDR1 and therefore, may not be resistant to DLC treatment. In fact, when one of the DLCs, MKT-077, was tested in phase 1 clinical trial, it was found to be nephrotoxic [58]. It appears that the use of DLCs for cancer treatment is limited due to the absence of p-gp in various normal tissues [59, 60].

In summary, the significance of these findings is that cancer cells that do not express MDR and contain high Δψp, are most likely to retain and be sensitive to Dox as well as DLCs. The same can be said of cardiac muscle cells, i.e., their unusual sensitivity to Dox may, at least in part, be explained by their high negative Δψp, whose electrical force would contribute to the intracellular attraction of positively charged compounds, and the absence of MDR to rid itself of these types of agents.

4 Targeting hypoxic tumor cells with glycolytic inhibitors: Lessons from DLCs

Most anticancer agents, which are currently used clinically, target the aerobic rapidly dividing cells of a tumor. Therefore, slow-growing anaerobic (hypoxic) cells found in necrotic centers and at the inner core of most solid tumors, will be resistant to these chemotherapeutic drugs. Thus, slow-growth may be considered another form of MDR. The fact that the most common toxicities of currently used chemotherapeutic agents in normal cells are found in the fastest dividing tissues, i.e., bone marrow, gut, and hair, provides further evidence that the selectivity of anticancer drugs in general lies not as much between tumor and normal cells as it does between rapidly dividing and slow, or nondividing cells.

The hypothesis on how to overcome these slow-growing drug-resistant tumor cells found in the hypoxic areas of tumors derives from work in which Rho123 was shown to inhibit mitochondrial oxidative phosphorylation (OxPhos) [61, 62]. Consequently, tumor cells treated with this drug have to rely solely on glycolysis for ATP production and thus become hypersensitized to inhibitors of glycolysis, such as 2-deoxy-d-glucose (2-DG). In fact it was shown that cotreating human breast carcinoma cells, MCF-7, with Rho 123 and 2-DG, 100% of the colony forming units was inhibited whereas similar treatment in normal epithelial cells showed little or no toxicity [17]. This concept was carried over to in vivo studies in which it was found that tumor bearing animals treated in combination with 2-DG and Rho123 were cured whereas when treated with either drug alone, only partial or no responses were obtained [18]. This latter result provides evidence that manipulation of OxPhos and glycolysis simultaneously can cure tumors in animals. Furthermore, these in vivo data also demonstrate that 2-DG can be administered safely to animals, at doses which are effective for antitumor activity in combination with an OxPhos inhibitor. In this regard, several reports have shown that low levels of 2-DG can be safely administered to animals for various reasons including hypersensitization of tumors to irradiation [63].

Since hypoxia, similar to Rho123 treatment, forces cells to rely mainly on anaerobic metabolism of glucose for survival, hypoxic tumor cells could be selectively targeted with inhibitors of glycolysis such as 2-DG [64]. The switch from aerobic to anaerobic metabolism in cells that lie in hypoxic regions of tumors creates two windows of selectivity that we believe will ultimately prove beneficial to cancer patients when 2-DG is used in combination with cytotoxic agents. The first is that tumor cells under hypoxia upregulate both glucose transporters and glycolytic enzymes [64], which favors increased uptake of 2-DG in these cells as compared to normal aerobic cells. The second is based on the principle that even if enough 2-DG is accumulated in normal cells to block glycolysis, they can survive by using oxygen to burn fats and proteins through their mitochondria to produce ATP. In contrast, when glycolysis is blocked in hypoxic tumor cells they die, since their mitochondria at these oxygen levels (8–57 μM) [64, 65] are less efficient in converting these alternative energy sources to ATP. These two windows of selectivity are based on fundamental principles of biochemistry and as such provide the basis for using glycolytic inhibitors to raise the efficacy of current chemotherapy by targeting the slow-growing hypoxic cell population found in most, if not all, solid tumors.

In order to investigate the mechanisms involved with hypersensitivity to glycolytic inhibitors, we developed three distinct models of simulated hypoxia which are referred to as chemical model A, genetic model B, and environmental model C [6668]. Model A approximates hypoxia by using chemicals such as Rho123, rotenone, antimycin A, and oligomycin to interfere with mitochondrial function thereby rendering the cell unable to produce ATP via OxPhos [66]. Model B uses tumor cells that have been permanently genetically altered by depletion of their mitochondrial DNA and cannot undergo OxPhos [66]. Since models A and B are growing under oxygen, but are unable to produce ATP via their mitochondria, they simulate hypoxic cells by relying exclusively on glycolysis for this function, whereas, model C are actually tumor cells grown under decreased atmospheric oxygen [68]. In all three models, glycolysis is increased (as measured by lactic acid) and most importantly, they are all found to be hypersensitive to glycolytic inhibitors as compared to their aerobic counterparts [66, 68]. However, model C is found to be less sensitive to glycolytic inhibitors than models A and B, which suggests that certain variables influencing the cellular response to glycolytic inhibitors differ between these models. One such variable is hypoxia-inducible factor-1 (HIF-1), which is expressed only in model C. HIF-1 is known to be a key regulator of a wide range of cellular responses to lowered oxygen tension [69]. Among the numerous genes activated by HIF-1, are glucose transporters and glycolytic enzymes [7073]. Since HIF-1 is activated in model C [69], but not in other models, it seems likely that HIF-1 induction is contributing to the increased cellular resistance to glycolytic inhibition by 2-DG found in this model. In this regard, a recent publication demonstrated that HIF-1 induced hexokinase 2 expression confers resistance to glycolytic inhibition by 2-DG, suggesting that combining inhibitors of HIF with 2-DG may be a more effective strategy than either agent alone, particularly for targeting the slow-growing hypoxic cell populations found in most solid tumors.

5 Conclusions

The studies which began by investigating the reasons why Dox seemed to preferentially affect cardiac and cancer cells has lead to a number of findings which have resulted in clinical applications. Notably, nonpositively charged analogs of Dox such as annamycin and AD 32 are shown to cause little or no cardiotoxicity and are able to overcome MDR in tumor cells expressing this mechanism of drug resistance [74, 75]. Similarly, a member of the DLC family, MKT-077, has been used in clinical trials as an antitumor drug based on its selectivity for accumulating in cancer mitochondria [58]. However, this approach has not yielded successful clinical results which is most likely due to a variety of factors including, the presence of MDR in many tumor types, the ineffectiveness of mitochondrial inhibition as a sole means of killing a cell and the toxicity in various normal tissues due to lack of MDR expression. On the other hand, hypoxic regions in solid tumors are shown to be hypersensitive to glycolytic inhibitors such as 2-DG [76]. Thus, combined with chemotherapeutic agents, 2-DG is currently in a Phase I clinical trial to selectively target the hypoxic cell population found in most solid tumors [77]. The biochemical differences in glucose metabolism between slow-growing hypoxic tumor cells and slow-growing aerobic normal cells provides a natural window of selectivity that can be exploited for therapeutic gain using glycolytic inhibitors.


delocalized lipophilic cation
hypoxia-inducible factor
multidrug resistance
oxidative phosphorylation
rhodamine 123


The authors have declared no conflict of interest.


1. Bonadonna G, Monfardini S. Cardiac toxicity of daunorubicin. Lancet. 1969;1:837. [PubMed]
2. Cortes EP, Gupta M, Chou C, Amin VC, Folkers K. Adriamycin cardiotoxicity: Early detection by systolic time interval and possible prevention by coenzyme Q1. Cancer Treat Rep. 1978;62:887–891. [PubMed]
3. Lampidis TJ, Henderson IC, Israel M, Canellos GP. Structural and functional effects of adriamycin on cardiac cells in vitro. Cancer Res. 1980;40:3901–3909. [PubMed]
4. Lampidis TJ, Johnson LV, Israel M. Effects of adriamycin on rat heart cells in culture: Increased accumulation and nucleoli fragmentation in cardiac muscle v. nonmuscle cells. J Mol Cell Cardiol. 1981;10:913–924. [PubMed]
5. Lampidis TJ. An in vitro model of anthracycline cardiopathogenesis. In: Mathe G, Maral R, De Jager M, editors. Anthracyclines 1981: Current Status and Future Development. Chapter 6 Masson Publication; New York: 1983.
6. Summerhayes IC, Lampidis TJ, Bernal SD, Nadakavukaren JJ, et al. Unusual retention of rhodamine 123 by mitochondria in muscle and carcinoma cells. Proc Natl Acad Sci USA. 1982;79:5292–5296. [PubMed]
7. Lampidis TJ, Savaraj N, Valet GK, Trevorrow K, et al. Relationship of chemical charge of anticancer agents to increased accumulation and cytotoxicity in cardiac and tumor cells: Relevance to multidrug resistance. J Cell Pharm. 1989;89:16–22.
8. Lampidis TJ, Salet C, Moreno G, Chen LB. Effects of the mitochondrial probe rhodamine 123 and related analogs on the function and viability of pulsating myocardial cells in culture. Agents Actions. 1984;14:751–757. [PubMed]
9. Johnson LV, Walsh ML, Bockus BJ, Chen LB. Monitoring of relative mitochondrial membrane potential in living cells by fluorescence microscopy. J Cell Biol. 1981;88:526–535. [PMC free article] [PubMed]
10. Chen LB, Summerhayes IC, Johnson LV, Walsh ML, et al. Probing mitochondria in living cells with rhodamine 123. Cold Spring Harb Symp Quant Biol. 1982;46:141–155. [PubMed]
11. Johnson LV, Walsh ML, Chen LB. Localization of mitochondria in living cells with rhodamine 123. Proc Natl Acad Sci USA. 1980;77:990–994. [PubMed]
12. El Baraka M, Deumie M, Viallet P, Lampidis TJ. Fluoroscence properties and partitioning behaviour of esterified and unesterified rhodamines. J Photochem Photobiol A: Chem. 1991;17:234–251.
13. Chen LB. Mitochondrial membrane potential in living cells. Ann Rev Cell Biol. 1988;4:155–181. [PubMed]
14. Johnson LV, Summerhayes IC, Chen LB. Decreased uptake and retention of rhodamine 123 by mitochondria in feline sarcoma virus-transformed mink cells. Cell. 1982;28:7–14. [PubMed]
15. Reed PW. Ionophores. Meth Enzymol. 1979;31:435. [PubMed]
16. Lampidis TJ, Bernal SD, Summerhayes IC, Chen LB. Rhodamine-123 is selectively toxic and preferentially retained in carcinoma cells in vitro. Ann NY Acad Sci. 1981;14:299–303.
17. Lampidis TJ, Bernal SD, Summerhayes IC, Chen LB. Selective toxicity of rhodamine 123 in carcinoma cells in vitro. Cancer Res. 1983;43:716–720. [PubMed]
18. Lampidis TJ, Hasin Y, Weiss MJ, Chen LB. Selective killing of carcinoma cells in vitro by lipohilic-cationic compounds: Cellular basis. Biomed Pharmacother. 1985;39:220–226. [PubMed]
19. Bernal SD, Lampidis TJ, McIsaac RM, Chen LB. Anticarcinoma activity in vivo of rhodamine 123, a mitochondrial-specific dye. Science. 1983;222:169–172. [PubMed]
20. Modica-Napolitano JS, Weiss MJ, Chen LB, Aprille JR. Rhodamine 123 inhibits bioenergetic function in isolated rat liver mitochondria. Biochem Biophys Res Commun. 1984;118:717–723. [PubMed]
21. Lampidis TJ, Savaraj N, Valet GK, Trevorrow K, et al. Relationship of chemical charge of anti-cancer agents to increased accumulation and cytotoxicity in cardiac and tumor cells: Relevance to multi-drug resistance. In: Tapiero H, Robert J, Lampidis TJ, editors. Anticancer Drugs, Collosque Inserm. John Libbey, Eurotext Ltd.; London: 1989. pp. 29–38.
22. Davis S, Weiss MJ, Wong JR, Lampidis TJ, Chen LB. Mitochondrial and plasma membrane potentials cause unusual accumulation and retention of rhodamine 123 by human breast adenocarcinoma-derived MCF-7 cells. J Biol Chem. 1985;260:13844–13850. [PubMed]
23. Kim HK, Park WS, Kang SH, Warda M, et al. Mitochondrial alterations in human gastric carcinoma cell line. Am J Cell Physiol. 2007;293:C761–771. [PubMed]
24. Modica-Napolitano JS, Aprille JR. Delocalized lipohilic cations selectively target the mitochondria of carcinoma cell. Adv Drug Del Rev. 2001;49:63–70. [PubMed]
25. Capuano F, Guerrieri F, Papa S. Oxidative phosphorylation enzymes in normal and neoplastic cell growth. J Bioenerg Biomembr. 1997;29:379–384. [PubMed]
26. Capuano F, Barone M, D'Eri N, Russo E, et al. Ursodeoxycholate promotes protein phosphorylation in the cytosol of rat hepatocytes. Biochem Mol Biol Int. 1997;41:329–337. [PubMed]
27. Cuezva JM, Ostronoff LK, Ricart J, Lopez de Heredia M, et al. Mitochondrial biogenesis in the liver during development and oncogenesis. J Bioenerg Biomembr. 1997;29:365–377. [PubMed]
28. Sun AS, Sepkowitz K, Geller SA. A study of some mitochondrial and peroxisomal enzymes in human colonic adenocarcinoma. Lab Invest. 1981;44:13–17. [PubMed]
29. Heerdt BG, Halsey HK, Lipkin LH, Augenlicht LH. Expression of mitochondrial cytochrome c oxidase in human colonic cell differentiation, transformation, and risk for colonic cancer. Cancer Res. 1990;50:1596–1600. [PubMed]
30. Luciakova K, Kuzela S. Increased steady-state levels of several mitochondrial and nuclear gene transcripts in rat hepatoma with a low content of mitochondria. Eur J Biochem. 1992;205:1187–1193. [PubMed]
31. Sul HS, Shrago E, Goldfarb S, Rose F. Comparison of the adenine nucleotide translocase in hepatomas and rat liver mitochondria. Biochim Biophys Acta. 1979;551:148–156. [PubMed]
32. Wallace DC. A mitochondrial paradigm of metabolic and degenerative diseases, aging, and cancer: A dawn for evolutionary medicine. Annu Rev Genet. 2005;39:359–407. [PMC free article] [PubMed]
33. Brandon M, Baldi P, Wallace DC. Mitochondrial mutations in cancer. Oncogene. 2006;25:4657–4662.
34. Riehm H, Biedler JL. Cellular resistance to daunomycin in Chinese hamster cells in vitro. Cancer Res. 1973;13:409–412. [PubMed]
35. Juliano RL, Ling V. A surface glycoprotein modulating drug permeability in Chinese hamster ovary cell mutants. Biochem Biophys Acta. 1976;455:152–162. [PubMed]
36. Shen DW, Fojo A, Robinson IB, Chin R, et al. Multi-drug resistance of DNA-mediated transformants is linked to transfer of the human MDR1 gene. Mol Cell Biol. 1986;6:4039–4044. [PMC free article] [PubMed]
37. Dano K. Active outward transport of daunomycin in resistant ehrlich ascites tumor cells. Biochim Biophys Acta. 1973;323:466–483. [PubMed]
38. Lampidis TJ, Fourcade A, Tapiero H. Relationship of membrane potential to acquired and intrinsic multiple drug resistance. In: Jaquillat C, Weil M, Khayat D, editors. Neo-adjuvant Chemotherapy Inserm Colloquim. John Libbey Eurotext Ltd.; London: 1988. pp. 655–660.
39. Priebe W, Perez-Soler R. Design and tumor targeting of anthracyclines able to overcome multidrug resistance: A double-advantage approach. Pharmacol Ther. 1993;60:215–234. [PubMed]
40. Pearce HL, Safa AR, Bach NJ, Winter MA, et al. Essential features of the p-glycoprotein pharmacophore as defined by a series of reserpine analogs that modulate multi-drug resistance. Proc Natl Acad Sci USA. 1989;86:5128–5132. [PubMed]
41. Zamora JH, Pearce HL, Beck WT. Physical-chemical properties shared by compounds that modulate multidrug resistance in human leukemia cells. Mol Pharmacol. 1988;33:454–462. [PubMed]
42. Ramu A, Ramu N, Gorodetsky R. Reduced oubain-sensitive potassium entry as a possible mechanism of multidrug resistance in P388 cells. Biochem Pharmacol. 1991;42:1699–1704. [PubMed]
43. Ramu A, Ramu N. Resistance to lipohilic cationic compounds in multidrug resistant leukemia cells. Leuk Lymphoma. 1993;9:247–53. [PubMed]
44. Zou Y, Ling YH, Van NT, Priebe W, Perez-Soler R. Antitumor activity of free and liposome-entrapped annamycin, a lipophilic anthracycline antibiotic with noncross-resistance properties. Cancer Res. 1994;54:1479–1484. [PubMed]
45. Priebe W, Van NT, Perez-Soler R. Removal of the basic center from doxorubicin partially overcomes multidrug resistance and decreases cardiotoxicity. Anticancer Drugs. 1993;4:37–48. [PubMed]
46. Selassie CD, Hansch C, Khwaja TA. Structure-activity relationships of antineoplastic agents in multidrug resistance. J Med Chem. 1990;33:1914–1919. [PubMed]
47. Edgar R, Bibi E. A single membrane-embedded negative charge is critical for recognizing positively charged drugs by the Escheria coli multidrug resistance protein MdfA. EMBO J. 1999;18:822–832. [PubMed]
48. Baguley BD, Ferguson LC. Relationship between the structure of analogs of amsacrine and their degree of cross-resistance to adriamycin-resistant P388 leukemia cells. Eur J Cancer Clin Oncol. 1988;24:205–210. [PubMed]
49. Burke TG, Morin MJ, Sartorelli AC, Lane PE, Tritton TR. Function of the anthracycline amino group in cellular transport and cytotoxicity. Mol Pharmacol. 1987;31:552–556. [PubMed]
50. Lampidis TJ, Kolonias D, Podona T, Israel M, et al. Circumvention of P-GP MDR as a function of anthracycline lipophilicity and charge. Biochemistry. 1997;36:2679–2685. [PubMed]
51. Dellinger M, Pressman B, Higgenson C, Kolonias D, et al. Structural requirements of simple organic cations for recognition by multi-drug resistant cells. Cancer Res. 1992;52:6385–6389. [PubMed]
52. Hait WN, Aftab DT. Rational design and preclinical pharmacology of drugs reversing multidrug resistance. Biochem Pharmacol. 1992;43:103–107. [PubMed]
53. Pawagi AB, Wang J, Silverman M, Reithmeier RAF, Deber CM. Transmembrane aromatic amino acid distribution in p-glycoprotein: A functional role in broad substrate specificity. J Mol Biol. 1994;235:554–564. [PubMed]
54. Burley SK, Petsko GA. Weakly polar interactions in proteins. Adv Protein Chem. 1988;39:125–189. [PubMed]
55. Burley SK, Petsko GA. Aromatic-aromatic interaction: A mechanism of protein structure stabilization. Science. 1985;229:23–28. [PubMed]
56. Tapiero H, Nguyen-Ba N, Lampidis TJ. Relevance of cross-resistance to the chemical structure of different anthracyclines in multi-drug resistant cell. Pathol Biol. 1994;42:328–337. [PubMed]
57. Brouty-Boye D, Kolonia D, Wu CJ, Savaraj N, Lampidis TJ. Relationship of multidrug resistance to rhodamine-123 selectivity between carcinoma and epithelial cells: Taxol and vinblastine modulate drug efflux. Cancer Res. 1995;55:1633–1638. [PubMed]
58. Britten CD, Rowinsky EK, Baker SD, Weiss GR, et al. A phase I clinical and pharmokinetic study of the mitoconhdrial specific rhodamine dye analog MKT 077. Clin Cancer Res. 2000;6:42–49. [PubMed]
59. Croop JM, Raymond M, Haber D, Devault A, et al. The three mouse multidrug resistance (mdr) genes are expressed in a tissue-specific manner in normal mouse tissues. Mol Cell Biol. 1989;9:1346–1350. [PMC free article] [PubMed]
60. Fojo AT, Ueda K, Slamon DJ, Poplack DG, Gottesman MM, Pastan I. Expression of a multidrug-resistance gene in human tumors and tissues. Proc Natl Acad Sci USA. 1987;84:265–269. [PubMed]
61. Abou-Khalil S, Abou-Khalil WH, Planas L, Tapiero H, Lampidis TJ. Effects of rhodamine 123 on mitochondria isolated from sensitive and resistant friend leukemia cell variants. Biochem Biophys Res Commun. 1985;127:1039–1044. [PubMed]
62. Modica-Napolitano JS, Weiss MJ, Chen LB, Aprille JR. Rhodamine 123 inhibits bioenergetic function in isolated rat liver mitochondria. Biochem Biophys Res Commun. 1984;118:717–723. [PubMed]
63. Mohanti BK, Rath GK, Anantha N, Kannan V, et al. Improving cancer radiotherapy with 2-deoxy-D-glucose: Phase I/II clinical trials on human cerebral gliomas. Int J Radiat Oncol Biol Phys. 1996;35:103–111. [PubMed]
64. Vaupel P, Kallinowski F, Okunieff P. Blood flow, oxygen and nutrient supply, and metabolic microenvironment of human tumors: A review. Cancer Res. 1989;49:6449–6465. [PubMed]
65. Rofstad EK, Halsør EF. Vascular endothelial growth factor, interleukin 8, platelet-derived endothelial cell growth factor, and basic fibroblast growth factor promote angiogenesis and metastasis in human melanoma xenografts. Cancer Res. 2000;60:4932–4938. [PubMed]
66. Liu H, Hu YP, Savaraj N, Priebe W, Lampidis TJ. Hypoxia increases tumor cell sensitivity to glycolytic inhibitors: A strategy for solid tumor therapy [model C] Biochem Pharmacol. 2002;64:1746–1751. [PubMed]
67. Liu H, Hu YP, Savaraj N, Priebe W, Lampidis TJ. Hypersensitization of tumor cells to glycolytic inhibitors. Biochemistry. 2001;40:5542–5547. [PubMed]
68. Maher JC, Krishan A, Lampidis TJ. Greater cell cycle inhibition and cytotoxicity induced by 2-deoxy-D-glucose in tumor cells treated under hypoxic vs aerobic conditions. Cancer Chemother Pharmacol. 2004;53:116–122. [PubMed]
69. Maher JC, Wangpaichitr M, Savaraj N, Kurtoglu M, Lampidis TJ. Hypoxic-induciblefactor-1 confers resistance to 2-deoxy-D-glucose in cells growing under hypoxia. Mol Cancer Ther. 2007;6:732–741. [PubMed]
70. Maxwell PH, Pugh CW, Ratcliffe PJ. Activation of the HIF pathway in cancer. Curr Opin Genet Dev. 2001;11:293–299. [PubMed]
71. Semenza GL, Roth PH, Fang H, Wang GL. Transcriptional regulation of genes encoding glycolytic enzymes by hypoxia-inducible factor 1. J Biol Chem. 1994;269:23757–23763. [PubMed]
72. Vaux EC, Metzen E, Yeates KM, Ratcliffe PJ. Regulation of hypoxia-inducible factor is preserved in the absence of a functioning mitochondrial respiratory chain. Blood. 2001;98:296–302. [PubMed]
73. Wang GL, Semenza GL. General involvement of hypoxia-inducible factor 1 in transcriptional response to hypoxia. Proc Natl Acad Sci USA. 1993;90:4304–4308. [PubMed]
74. Zou Y, Priebe W, Stephens LC, Perez-Soler P. Preclinical toxicity of liposome-incorporated annamycin: Selective bone marrow toxicity with lack of cardiotoxicity. Clin Cancer Res. 1995;1:1369–1374. [PubMed]
75. Lampidis TJ, Johnson LV, Israel M. Effects of adriamycin on rat heart cells in culture: increased accumulation and nucleoli fragmentation in cardiac muscle v. nonmuscle cells. J Mol Cell Cardiol. 1981;13:913–924. [PubMed]
76. Boutrid H, Cebulla CM, Pina Y, Jockovich ME, et al. Regional hypoxia in LHBetaTag murine retinoblastoma: Implication for novel treatments. Invest Ophthalmol Vis Sci. 2008;49:2799–2805. [PubMed]
77. Raez LE, Langmuir V, Tolba K, Rocha-Lima CM, et al. Responses to the combination of the glycolytic inhibitor 2-deoxy-glucose [2-DG] and docetaxel [DC] in patients with lung and head and neck [H/N] carcinomas. J Clin Oncol. 2007;25:14025.