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The genetic basis for the Hereditary Leiomyomatosis and Renal Cell Cancer (HLRCC) syndrome is germline inactivating mutation in the gene for the Krebs/tricarboxylic acid (TCA) cycle enzyme, fumarate hydratase (FH), the enzyme that converts fumarate to malate. These individuals are predisposed to development of leiomyomas of the skin and uterus as well as highly aggressive kidney cancers. Inhibition of FH should result in significant decrease in oxidative phosphorylation (OXPHOS) necessitating that glycolysis followed by fermentation of pyruvate to lactate will be required to provide adequate ATP as well as to regenerate NAD+. Moreover, FH deficiency is known to upregulate expression of hypoxia inducible factor (HIF)1 α by enhancing the stability of HIF transcript. This leads to activation of various HIF regulated genes including vascular endothelial growth factor (VEGF), glucose transporter GLUT1, and increased expression of several glycolytic enzymes. Since lactate dehydrogenase-A (LDH-A), also a HIF1 α target, promotes fermentative glycolysis (conversion of pyruvate to lactate), a step essential for regenerating NAD+, we asked whether FH deficient cells would be exquisitely sensitive to LDH-A blockade. Here we report that HLRCC tumors indeed overexpress LDH-A; that LDH-A inhibition results in increased apoptosis in a cell with FH deficiency and that this effect is reactive oxygen species (ROS) mediated; and that LDH-A knockdown in the background of FH knockdown results in significant reduction in tumor growth in a xenograft mouse model.
Many cancer cells show enhanced glycolysis (1). In recent years, the molecular basis for this effect has started to be elucidated. In particular, upregulation of hypoxia inducible factor (HIF), which occurs as a consequence of hypoxia as well as from alterations in certain oncogenes or mutations in tumor suppressor genes, results in the increased transcription of genes involved in glucose transport, glucose metabolism, lactate formation and lactate export from cells (2–6). Moreover, HIF activation results in decreased activity of the pyruvate dehydrogenase complex (7, 8). Collectively, these studies provide a partial explanation for the Warburg effect, which states that even under aerobic conditions, cancer cells preferentially undergo glycolysis followed by fermentation (9). Fumarate hydratase (FH) plays an important role in the mitochondrial TCA cycle by catalyzing the conversion of fumarate to malate. The primary function of the Krebs/tricarboxylic acid (TCA) cycle is oxidation of pyruvate to generate ATP. The genetic basis for the HLRCC syndrome is germline inactivating mutation in the FH gene (10–12). Inhibition of FH and presumably of OXPHOS should necessitate that fermentative glycolysis will be required to provide ATP as well as regenerate NAD+ for glycolysis. The pseudohypoxic drive in HLRCC involving fumarate-dependent HIF activation as a result of competitive HIF prolyl hydroxylase (HPH) inhibition by accumulated fumarate is likely to lock FH deficient cells into a glycolytic pathway for sustaining the metabolic energy requirements (13).
In the present study, we wanted to ask whether FH deficient cells do rely on glycolysis for growth and/or proliferation and whether they would be sensitive to inhibition of fermentative glycolysis. The fermentative reaction is regulated by lactate dehydrogenase, which in most tissue occurs in two major isoforms LDH-A and LDH-B. LDH-B is a homotetramer of LDH-H and favors the conversion of lactate to pyruvate and is also known to be upregulated in many cancers (14). LDH-A is a heterotetramer of LDH-M and favors conversion of pyruvate to lactate and is upregulated in many solid tumors that have high HIF expression (3, 15). LDH-A is an attractive target for cancer therapy since its expression is largely relegated to skeletal muscle. It is not present in red cells, in which glycolysis followed by fermentation is an obligatory process for energy generation. Moreover, it is well known that humans with LDH-A deficiency only show myoglobinuria under intense anaerobic exercise (16, 17) and individuals with complete lack of LDH-A or B subunit have been documented with no apparent increase in hemolysis (16, 18, 19). Therefore it is likely that potential LDH-A inhibitors might show relatively modest systemic toxicity.
Patients with suspected or confirmed HLRCC and VHL were evaluated in the Urologic Oncology Branch of the National Cancer Institute, where they underwent a comprehensive clinical evaluation for phenotype assessment. Each of the ten patients with HLRCC-associated kidney cancer came from a family with confirmed fumarate hydratase germline mutation; six had surgery at the NCI, the other four had surgery at outside institutions. The clear cell kidney cancers came from von Hippel Lindau patients with confirmed germline mutation of the VHL gene; all six had kidney cancer surgery performed at the National Cancer Institute. The LDH-A staining in VHL and HLRCC tumor samples was compared with nonmalignant kidney tissue. The pathology of all of the renal tumors was reviewed by co-author MM.
Five micron slides form formalin-fixed paraffin embedded tissue samples were used for immunohistochemistry. Slides were deparafinized in 3 changes of xylene for 5 min each followed by rehydration in graded alcohols. Antigen retrieval was achieved by heating the slides in Tris-EDTA buffer pH 9.0 in a microwave oven at 95°C for 20 min. Endogenous peroxidase activity was inhibited by incubation in 3 per cent hydrogen peroxide in methanol for 10 min. Sections were then incubated for 1 hr. at room temperature with sheep anti-human LDH-A antibody (1:30,000, Abcam, Cambridge, MA) or with isotype-matched IgG as negative controls. After several washes in Tris buffer, samples were further incubated with rabbit anti-sheep IgG (1:1000, Millipore, Danvers, MA) as a secondary antibody for 30 min at room temperature. Slides were then incubated with goat anti-rabbit HRP polymer (Envision™ PO System; Dako, Carpinteria, CA) for 30 min followed by 3,3-diaminobenzidine as chromogen for 5 min. The sections were counterstained with Mayer’s haematoxylin and permanently mounted. Staining for LDH-A in HLRCC and VHL null tumors was evaluated in 4 high-power fields and samples were scored as strongly positive (3+), moderately positive (2+), weakly positive (1+) or negative (0). Staining pattern (membranous, cytoplasmic or nuclear) was also recorded.
Cell culture medium was purchased from American Type Culture Collection (ATCC; Manassas, VA). Lentiviral shRNA (Short hairpin RNA) constructs were purchased from Sigma Aldrich (St. Louis, MO) and the viral power kit and packaging cell line 293FT were purchased from Invitrogen (Carlsbad, CA). Sheep polyclonal LDH-A antibodies were purchased from Abcam (Cambridge, MA), and FH mouse monoclonal from Novus (Littleton, CO). Rabbit polyclonal for activated caspase-3 and activated PARP were purchased from Cell Signaling (Cambridge, MA). The assay kit for measuring ATP was purchased from Roche (Indianapolis, IN). Lactate measurement kit was purchased from CMA microdialysis (N. Chelmsford, MA). Cell cycle reagents and annexin-V assay kits were purchased from Guava Technologies (Hayward, CA). Athymic mice for tumor implantation were purchased from Charles River Laboratories (Wilmington, MA). A549 cells were maintained in Ham’s F12 medium with glutamine, and sodium pyruvate with 10% fetal calf serum (Sigma). RCC4 and RCC4+VHL cells were gift from Celeste Simon at University of Pennsylvania School of Medicine. Sequences of shRNA used in this study; LDH-A: CCACCATGATTAAGGGTCTTT LDH-A#1: GATCTGTGATTAAAGCAGTAA, FH63: CGCTGAAGTAAACCAGGATTA; FH65: CCCAACGATCATGTTAATAAA.
A549 cells were infected separately with empty shRNA vector control, three different FH shRNAs and LDH-A lentiviral particles as described (20). Briefly, Recombinant lentiviral particles were produced by transient transfection of 293T cells according to standard protocol. Briefly, subconfluent 293FT cells were cotransfected with 3μg of a shRNA plasmid, and 9μg viral power packaging mix (an optimized proprietary mix of three plasmids, pLP1, pLP2, and pLP/VSVG from Invitrogen) using lipofectamine 2000 (Invitrogen). After 16 h culture medium was switched to regular growth medium and cells were allowed to incubate for additional 48 hours. Conditioned cell culture media containing recombinant lentiviral particles were harvested and frozen. A549 cells were treated with above cell culture supernatant containing lentiviral particles for 24 hours. These cells were then selected in puromycin (Sigma Aldrich) to generate stable cell lines encoding empty vector shRNA, FH shRNA, and LDH-A shRNA. The selected cell lines were validated for diminished LDH-A and FH expression by western blot analysis. Briefly, total cellular proteins were separated by SDS-PAGE and electro transferred to PDVF membranes and immunoblotted with anti-LDH-A (Abcam) or anti-FH (Novus) antibody for overnight at 4°C. After washing with TBS-T, the membrane was incubated with secondary antibody of choice for 30 min. The protein bands were detected using SuperSignal West Pico Chemiluminescent substrate (Pierce). For generation of FH/LDH-A clones, the A549 cells were infected with FH and LDH-A shRNA’s and selected with puromycin and validated with western blot analysis with FH and LDH-A antibodies for double knockdown. For tumor implantation double stable FH/LDH-A cells were generated by larger scale infection and tested for the lack of FH/LDH-A expression before implantation. LDH-A deficient RCC4 cells were generated in a similar manner as described above and were validated for LDH-A inhibition by western blot analysis.
Control, FH deficient, and FH/LDH-A deficient cell lines were plated in 60-mm dish at a density of 1 × 105 cells/dish in HAM’s F12 medium supplemented with 10% FBS for 24 h at 37°C in a 5% CO2 incubator. After 24, 48, 72, 96 and 120 hours of initial plating, cells were scraped, washed with PBS and re-suspended in 1 ml of Hanks’ buffer and counted in presence of trypan blue. All samples were assayed in duplicate to generate proliferation curves as described (21).
We measured lactate accumulation in control and LDH-A deficient cell lines as described (22). Briefly, the cell culture conditioned supernatant was used to measure lactate levels by a simple calorimetric analysis based on the reduction of the tetrazolium salt INT in a NADH-coupled enzymatic reaction to formazan, which is water-soluble and exhibits an absorption maximum at 492 nm. The intensity of the red color formed is proportional to the lactate concentration in the conditioned medium. All samples were assayed in three independent experiments in triplicate. The average value of the absorption readings were used for graphical representation.
LDH-A activity was measured in the direction of reduction of pyruvate to lactate by monitoring the changes in the absorbance of NADH at 340 nm, 25°C in potassium phosphate buffer, pH 7.4. Briefly, the reaction mixture contained 50 mM potassium phosphate buffer (pH 7.4) and pyruvate (200 μM). The reaction was initiated by addition of cell extract prepared in potassium phosphate buffer, and NADH (250 μM), and decrease in absorbance at 340 nm was monitored every 4 minutes.
Intracellular ATP levels in control, FH, and FH/LDH-A deficient cells were measured according to manufacturer’s instructions and as described before (7). In brief, cell lysates were collected and luminescence was measured using a luminescence reader (Molecular Devices), and normalized for protein concentration.
Apoptosis was measured by Guava PCA-96 Nexin (Guava Technologies) as per the manufacture’s protocol. Briefly, cells were harvested and re-suspended in 1X Nexin buffer with Annexin-V-PE, and Nexin 7-AAD. The cells were allowed to incubate for 15 minutes and analyzed in the Guava flowcytometer.
The invasiveness of cells was analyzed using Falcon BioCoat Matrigel chambers (Becton Dickinson) with 6.4-mm diameter and 8-μm pore size. Invasiveness of cells expressing control shRNA, FH shRNA, and FH/LDH-A shRNA was assayed as described before (23, 24). Briefly, 4 × 104 cells were seeded on the upper side of a Matrigel invasion chamber. After 24 h, the cells on the upper surface of the filter were removed using cotton swabs, and the invaded cells were fixed with 20% solution of 4% para-formaldehyde and stained with crystal violet. Incorporated dye was extracted with 0.05M sodium phosphate (pH4.5) in 50% ethanol and the absorbance of released dye was read at 550 nm. The data represents average of two independent experiments performed in triplicate.
The XF24 Extracellular Flux analyzer (Seahorse Biosciences, Billerica, MA) is a fully integrated 24-well instrument that measures in real time the uptake and release of metabolic endproducts. Each XF24-assay well contains a disposable sensor cartridge, embedded with 24 pairs of fluorescent biosensors (oxygen and pH), coupled to fiber-optic waveguides. The wave guides delivers light ray at various excitation wavelengths (oxygen= 532 nm, pH= 470 nm) and transmits a fluorescent signal (oxygen= 650 nm, pH= 530 nm) to a set of highly sensitive photodetectors. This technology was used to measure oxygen consumption (OCR) expressed in pMoles/min and extra cellular acidification rate (ECAR), expressed in mpH/min in control, FH deficient and FH/LDH-A deficient cell lines as described (25).
Intracellular ROS production was measured by staining with dichlorodihydrofluorescein diacetate (CM-H2DCFDA, Invitrogen). CM-H2DCFDA is a cell-permeant indicator for reactive oxygen species that is nonfluorescent until removal of the acetate groups by intracellular esterases and oxidation occurs within the cell. The procedure for measuring ROS was carried out as described earlier (7), with minor modification. Briefly, cells were loaded with 5 μM H2DCFDA for 1 hr, washed in PBS, and incubated in fresh media for 30 min. The cells were then subjected to FACS analysis to visualize the fluorescence.
3.0 × 106 control and FH65/LDH-A deficient cells were subcutaneously implanted in 8 athymic mice. The implantation was done such that each mouse had FH65 cells on the right flank and FH65/LDH-A deficient cells on the left flank. The control shRNA cells were implanted in separate set of athymic mice. Tumors were measured every five days and tumor volume was calculated as described before (26). Tumor lysates were prepared by homogenization of tumor tissues in lysis buffer and were separated by SDS-PAGE and electro transferred to PDVF membranes and immunoblotted with anti-LDH-A and anti-FH antibody and normalized by actin as a loading control.
The Student’s t test was used to evaluate the statistical significance of the results.
Since FH deficiency up regulates HIF1 α (13), and LDH-A is a known HIF1 α target, we asked whether expression of LDH-A is significantly enhanced in HLRCC tumors. Previous microarray profiling data in FH deficient uterine fibroids suggest that LDH-A expression may be increased compared to normal myomatrium (27). As shown in Fig. 1, LDH-A expression is abundant in HLRCC tumors, and almost negligible in surrounding normal tissue. This expression pattern largely overlaps with HIF expression in the corresponding section from the same tumor (Fig. 1B). This result is almost universal in HLRCC tumor samples that were sequence validated for FH mutations (Fig. 1C and D). Based on these results, we wanted to ask whether inhibition of LDH-A in an FH deficient background would have significant impact on the survival or the rate of proliferation of FH deficient cells in vitro.
Since there is no established HLRCC cell line that can be used to study metabolic alterations and signaling events associated with FH deficiency, we used the A549 lung cancer cell line as a surrogate model system to generate FH deficient cell lines. Our choice of the cell line to create HLRCC surrogate model system in cell culture was based on the previous publication that suggested these cells carry wild type VHL gene, as is the case in HLRCC patients and inhibition of endogenous FH expression in these cells results in enhanced glycolysis (13). Western blot analysis with anti-FH antibody demonstrates that cell lines FH63, and FH65, which express shRNAs targeting two different regions within the FH mRNA, have diminished FH protein expression compared to the cell line expressing control shRNA (Fig. 2A). The FH deficient cells compared to control cells have comparable rates of proliferation (Fig. 2B). However, in agreement with the study by Isaacs et al (14) FH deficient cells compared to control cells have increased lactate levels (Fig. 2C), enhanced HIF1 α expression in normoxic conditions (Fig. 2F) and showed increased expression of two HIF1 α target genes, GLUT 1 and VEGF (Fig. 2D and E). Since FH deficiency mimics the pseudohypoxic state that should favor fermentative glycolysis regulated by LDH-A, a HIF1 α target, we also found increased HIF1 α expression in FH deficient cells and a 2.0 fold increase in LDH-A enzyme activity in FH deficient cells as compared to empty vector control or wild type cells as measured by loss of NADH in the presence of pyruvate (Fig. 2G).
To test whether inhibition of LDH-A in FH deficient cells would result in reduced proliferation, we generated LDH-A deficient cells in an FH deficient background and will refer these cells as FH/LDH-A deficient cells in this manuscript. Western blot analysis with anti-FH and anti LDH-A antibodies demonstrates that cell line FH63/LDH-A, and FH65/LDH-A have diminished FH and LDH-A protein expression compared to the control cell line or FH only deficient cell lines (Figs. 3A and B). As expected, FH65 shRNA cells show increased lactate formation as measured by lactate accumulation and real time measurements in an XF-24 metabolite analyzer as compared to control cells but FH/LDH-A deficient cells show reduced lactate formation (Figs. 3C and D).
Next, we asked whether cell lines deficient in FH/LDH-A expression would show altered cell survival/growth. We first characterized these stable cell lines in various in vitro assays. As shown in Fig. 4A, all FH/LDH-A deficient cell lines display a significant proliferation defect compared to cells with LDH-A knockdown in the background of wild type FH gene. The FH/LDH-A deficient cells also demonstrate significantly higher levels of apoptosis as compared to control shRNA cells (Fig. 4B). Moreover, induction of apoptosis is also demonstrated by western blot analysis of caspase-3 cleavage and PARP cleavage in FH/LDH-A deficient cell lines (Fig. 4C). This data could also be extended to growth in 3-dimensions: we have noted that in a soft agar growth assay, LDH-A deficient cells on an FH deficient background generate smaller colonies compared to FH deficient cells suggesting lower tumorigenic potential (data not shown).
As metastatic potential is one of the hallmarks of HLRCC tumors (28, 29), we asked whether a cell line deficient in FH would display a more invasive phenotype as assessed by a matrigel invasion assay. We found that FH deficient cells have significantly enhanced invasive potential in this assay and this invasive behavior can be dialed back to the same level as cells expressing control shRNA (Fig. 3E) by LDH-A inhibition. Moreover, this invasive ability correlated with lactic acid generation (Fig. 3C and D).
To determine impact on ATP generation of LDH-A inhibition in FH deficient cells, ATP levels were measured. In comparison to the control cells, or FH alone deficient cells, FH/LDH-A deficient cells show significant ATP depletion (~50–60% reduction) (Fig. 5B). This result is in agreement with the substantially reduced proliferation rate observed in our various growth assays. We were unable to show increased NADH/NAD+ ratio (data not shown) in FH/LDH-A deficient cells. Furthermore, we also noted an increased rate of oxygen consumption in FH/LDH-A deficient cells suggesting that LDH-A inhibition in FH deficient background may be forcing the cells to increase OXPHOS (Fig 5B).
VHL deficiency is a common feature of clear cell renal carcinoma (30, 31). It also results in HIF1 α upregulation (32). We would therefore expect enhanced LDH-A expression in VHL deficient renal cancer tumor tissue samples. This was indeed found to be the case (Fig. 6A). Moreover, we have also shown that inhibition of LDH-A by two different shRNAs in the VHL deficient cell line RCC4, results in increased apoptosis (Fig. 6B and C). RCC4 cells reconstituted with wild type VHL lacks LDH-A expression as a result of diminished HIF1 α expression (33), and hence these cells cannot be used to generate LDH-A deficient cell lines for comparison studies between VHL deficient and VHL reconstituted RCC4 cells. Our analysis of VHL deficient tumors and in vitro data suggests that LDH-A is overexpressed in VHL deficient cells and its inhibition results in apoptosis.
Based on our previous results, we asked whether enhanced OXPHOS due to LDH-A knockdown in FH deficient cells might result in increased formation of reactive oxygen species. 2′,7′-dichlorodihydrofluorescein diacetate (H2DCFDA) is a fluorogenic probe commonly used to detect cellular ROS. We found enhanced ROS levels as measured by H2DCFDA in FH/LDH-A deficient cells compared to control or FH deficient cells (Fig. 5C). Furthermore, this ROS generation could be blocked by the antioxidant n-acetyl cysteine (NAC), and resulted in partial rescues from apoptosis as measured by cleaved PARP (Fig. 5D). This data suggests that ROS generation may be partly implicated in the increased apoptosis seen in FH/LDH-A deficient cells.
To ask whether FH/LDH-A deficient cell lines have significantly diminished potential for tumor formation, we implanted these cells into the flanks of athymic mice. We found that FH65/LDH-A knockdown cells show marked growth inhibition (Fig. 7A) compared to cells expressing control shRNA or FHshRNA. We also show representative tumor tissue from control, FH deficient, and FH/LDH-A deficient tumors assayed for FH and LDH-A expression (Fig. 7B) to confirm that tumor cells in vivo displayed diminished expression of FH and LDH-A as they had in vitro. These results are similar to those recently published by Leder et al (34).
The major findings in this report are: a) HLRCC tumors over express LDH-A; b) LDH-A inhibition results in increased apoptosis via ROS production in an A549 surrogate FH knockdown cell line; and c) inhibition of fermentative glycolysis in this cell line results in significant reduction in growth in a xenograft mouse model.
The common feature of tumor cells is their reliance upon fermentative glycolysis, a phenomenon coined as the Warburg effect (1). Although the mechanism that may control this metabolic shift is not clearly defined, many lines of investigations suggest that HIF expression may be central to Warburg’s effect both by induction of fermentative glycolysis and by diminishing the activity of PDH (7, 8). FH inactivation should mimic this state, as it results in increased expression of HIF1 α and upregulaiton of HIF-dependent glycolysis (13, 35). This pseudohypoxic drive is similar to that described in VHL deficient cells as a result of the HIF accumulation in the absence of the appropriate ubiquitination process that targets HIF. The switch to the fermentative glycolysis should therefore be a common feature in tumors with VHL, FH, and SDH (succinate dehydrogenase) deficiency, resulting from HIF1 α stabilization, and we would expect to see increased LDH-A expression in these tumors. Indeed, we have shown this to be the case in two of these tumor types, namely VHL and FH deficient tumors.
The functional consequences of inhibiting LDH-A have been explored both in vitro and in vivo. In the background of FH deficiency, LDH-A knockdown cells proliferate slower, undergo apoptosis and are less invasive. The underlying basis for these effects remains to be fully defined. FH/LDH-A deficient cells did show a ~50–60% reduction in intracellular ATP. We were surprised that the levels were not more profoundly depressed, since both fermentative glycolysis, and TCA cycle were expected to be compromised in FH/LDH-A cells. It is possible that intracellular ATP levels are being sustained either by leakiness in our surrogate system, by LDH-B overactivity or expression compensating for LDH-A inhibition, or by diminished ATP consumption. We are currently exploring whether inhibition of both isoforms of LDH may result in significant ATP depletion. This might be technically challenging, as it will involve a triple knockdown to create such a cell line.
A correlation between apoptosis, ROS, and mitochondrial respiration had been reported in several cancers (36). Studies have suggested that HIF1 α dependent mitochondrial repression may provide survival benefit by decreasing the risk associated with apoptotic cell death (37). Moreover, reduced ROS levels and decreased apoptosis results from forced expression of HIF1 α in oral squamous cell carcinoma (38). We therefore asked if increased ROS production was mediating apoptosis. This appears to be the case, at least partially, since NAC was able to decrease PARP cleavage. We also found that OXPHOS was enhanced in FH/LDH deficient cells – as evidenced by increased oxygen consumption. One effect of this might be to increase electron flow in OXPHOS and thereby account for increased ROS production. However, in preliminary observations, we did not find increased superoxide formation as measured by the Mitosox reagent (data not shown), so the source of increased ROS production remains to be defined. It is also possible that various anti-oxidant systems in these cells have also been affected accounting for increased ROS.
Another effect of the increased rate of oxygen consumption observed in FH/LDH-A deficient cells could be to maintain the NADH/NAD+ ratio. Since LDH-A catalyzes regeneration of NAD+ from NADH, we expected to observe an increased NADH/NAD+ ratio in FH/LDH deficient cells. The competitive phenomenon between mitochondrial NADH/NAD+ transport system and LDH-A for NADH consumption is driven by LDH-A as a result of its fast enzymatic kinetic properties and is thought to be a potential factor for decreased mitochondrial respiration (39). Our failure to note an increase in the NADH/NAD+ ratio may be partly due to a decrease in the LDH-A driven competitive reaction that drives more NADH into mitochondria.
Since hypoxia regulates transcription factors HIF1 and HIF2, and in general it is difficult to design inhibitors targeting transcription factors, we have investigated whether targeting downstream targets of HIF1 and HIF2 may be therapeutically advantageous. Indeed, VEGF, a HIF1 α induced gene product has been successfully targeted for renal cell carcinoma therapy and our in-vitro data showing enhanced VEGF expression in these cells supports the use of VEGF inhibitors in HLRCC. Our in vitro and in vivo data supports the notion that targeting LDH-A, another HIF1 α target may be a viable strategy, for treating this disease as well. Moreover, high lactate levels are associated with poor prognosis in advanced renal cell carcinoma (40) and in head and-neck cancer (41). It is likely that inhibition of LDH-A by increasing the extracellular pH (42, 43) may reduce the metastatic ability of these advanced cancer cells as extrapolated from our in vitro invasion assay results. Also, HLRCC patients may benefit from combined anti-angiogenic and LDH-A inhibitor therapy, as might patients with any tumors that have upregulated HIF.
Our findings add to a growing literature that suggest that metabolism plays a key role in tumorigenesis and is linked either directly or indirectly to hallmarks of cancer that are involved in initiation and proliferation of tumors. In conclusion, our data support the hypothesis that inhibition of fermentative glycolysis might serve as a therapeutic strategy for HLRCC. Therefore, the development of inhibitors of LDH-A makes eminent sense for the treatment of HLRCC and a potentially large group of cancers in which the Warburg effect is operational, perhaps as many as 60–80% of tumor types.
PS is partially supported by Temin award from NCI (K01 CA104700). PS, HX, AMA, SS, and VPS are partially supported by Dana-Farber/Harvard Cancer Center (DF/HCC) Kidney Cancer Specialized Program of Research Excellence (SPORE). VV, MJM, and WML supported by National Cancer Institute, National Institutes of Health under contract N01-C0-12400.