A new discovery about pancreatic cancer sheds light on how the cancer fuels its growth and may help explain how promising cancer drugs work—and for whom they will fail.
The finding one day could help doctors determine which treatments will be most effective for patients, so that they get the best outcomes.
“Pancreatic cancer is a very difficult problem.
It has been a very difficult problem for a long time.
The survival for pancreatic cancer patients is very low compared to other tumors,” said researcher David F. Kashatus, Ph.D., of the University of Virginia School of Medicine and the UVA Cancer Center.
“We’re really trying to understand the biology so that scientists and drug developers can be more informed as they try to tackle this disease.
Any progress we can make, no matter how small, is going to be an improvement over the current state of affairs.”
A Big Mystery of Pancreatic Cancer
The new discovery represents the fulfillment of years of work for Kashatus, who first proposed the research project while interviewing at UVA in 2012.
It also helps to answer an 80-year-old mystery: Why and how do cancers rewire cells to fuel themselves using a much more inefficient process?
Scientists previously have noted strange changes in the shape of mitochondria, the powerhouses of cells, in cancers driven by mutations in the RAS gene.
Kashatus wanted to understand what was occurring and how it affected pancreatic cancer’s growth.
Kashatus found that when the mutated RAS gene gets activated, it causes the mitochondria to fragment.
This fragmentation supports the earliest shifts toward the cancer’s new fueling process.
This was quite surprising, because suddenly the mitochondria were playing a very unusual role. Their division was actually helping the cancer establish itself.
But there’s good news: This process could prove to be a weakness for the cancer that doctors could exploit to help patients. Kashatus found that blocking mitochondrial division in tumor samples largely prevented the tumors from growing.
And when they did grow, the cancer cells gradually lost mitochondrial function.
This was bad for the cancer, and the loss of mitochondria represents another weakness doctors could exploit.
“This mitochondrial fragmentation is really playing two distinct roles: On the one hand, it’s promoting this shift in metabolism. But it’s also promoting mitochondrial health,” Kashatus said.
“These two things are combining to drive the pancreatic tumor growth process. So I think this is something that could be therapeutically valuable. But it also really teaches us about pancreatic tumor growth in general.”
Explaining How Cancer Drugs Work
The finding also may help explain the workings of several drugs in development, and it could help doctors understand which patients they will benefit, said Kashatus, of UVA’s Department of Microbiology, Immunology and Cancer Biology.
“Inhibiting [mitochondrial division in patients’ cancer cells] would be a nice future goal for us.
However, the drugs targeting this process are really very early in development, and so it’s not something that will really be ready for the clinic anytime soon,” he said.
“But this work can really help us understand how some of these other drugs that are a little bit further along in the process may be acting, so that we can better understand which patients may or may not benefit.”
Pancreatic ductal adenocarcinoma (PDAC) relies on energy produced by oxidative phosphorylation (OXPHOS) from the mitochondria to grow and metastasize (1). Mitochondria within a cell can collectively alter their structures to optimize their metabolic functions in response to cellular insults (2). For instance, mitochondria join together in a process called mitochondrial fusion, which is critical for organelle quality control (3). In other cases, mitochondria fragment into smaller organelles through a controlled process called mitochondrial fission, which is often a response to oxidative stress (4). The homeostatic processes of fission and fusion in response to cellular demands is often referred to as mitochondrial dynamics.
The morphology of these networked mitochondria is regulated by a few key proteins. The mitofusin family of proteins (MFN1 and MFN2) positively regulate the fusion of mitochondria by bringing together the outer mitochondrial membranes. Dysfunction or deficits in MFN expression can lead to unopposed mitochondrial fission and possibly disease, such as Charcot-Marie-Tooth type 2A syndrome, which is caused by an autosomal dominant mutation in MFN2 (5). On the other hand, the GTPase dynamin-related protein-1 (DNM1L/DRP1) regulates mitochondrial fission, and its dysfunction may promote unregulated mitochondrial fusion (6).
Pancreatic cancer cells exhibit highly fragmented mitochondria (7), which suggests basal mitochondrial dynamics that favor mitochondrial fission. We hypothesized that shifting the balance of mitochondrial dynamics toward mitochondrial fusion in PDAC cells would normalize their function and reduce oncogenicity, as has been suggested in previous cell culture studies (8). We also reasoned that such an approach may have a favorable therapeutic ratio, since mitochondrial fusion is more frequently observed in normal cells (9). Indeed, we found that the genetic or pharmacologic activation of mitochondrial fusion reduces PDAC growth and improves survival in mouse models of pancreatic cancer. Moreover, we found that the induction of mitochondrial fusion correlates with less mitochondrial mass and reduced OXPHOS compared with controls. We provide evidence that mitochondrial fusion induces mitophagy in pancreatic cancer cells, which may selectively reduce the functional mitochondrial mass in tumors. These proof-of-principle experiments demonstrate the critical nature of mitochondrial dynamics and how they could potentially be exploited therapeutically against PDAC.
Genetic or pharmacological inhibition of mitochondrial fission promotes mitochondrial fusion and suppresses OXPHOS. Pancreatic cancer is frequently driven by oncogenic KRAS, the downstream signaling of which activates DRP1 and promotes mitochondrial fission (10). Whether disrupting mitochondrial fission would have a therapeutic effect in pancreatic cancer was unknown. Toward this end, we used CRISPR/Cas9 methodology to edit the endogenous locus of Dnm1l/Drp1 to ablate expression of DRP1 in murine KPC cells syngeneic to C57BL/6 that also expressed a luciferase transgene. We confirmed a nearly quantitative abrogation of DRP1 by Western blot (Figure 1A). This loss of DRP1 expression led to unopposed mitochondrial fusion, as observed with confocal microscopy with MitoTracker staining, which selectively stains live mitochondria in cells (Figure 1B; for other clones see Supplemental Figure 1A; supplemental material available online with this article; https://doi.org/10.1172/jci.insight.126915DS1). Control KPC cells edited with guide RNAs to GFP (sgGFP) retained highly fragmented morphology (Figure 1B), but sgDrp1 cells normalized the mitochondria in more than 70% of the cells (Figure 1B and Supplemental Figure 1A). These data were confirmed by transmission electron microscopy (TEM), where we found that the average mitochondrial length after Drp1 knockout doubled compared with sgGFP controls (Figure 1C; sgGFP, 0.53 μm vs. sgDrp1, 1.26 μm, P < 0.0001).
The induction of mitochondrial fusion by DRP1 knockout decreased oxygen consumption rates (OCR), as determined by extracellular flux assay (Figure 1D). In addition, we observed lower basal respiration, spare respiration capacity, and ATP production in sgDrp1 cells compared with controls (Figure 1E; repeated in other clones in Supplemental Figure 1, B and C). This reduced OCR was correlated with reduced growth in vitro (Supplemental Figure 1D) and increased G1 arrest (Supplemental Figure 1E). To determine the significance of these metabolic changes, we orthotopically implanted Drp1-knockout cells into the pancreata of recipient C57BL/6 mice and observed impaired in vivo growth (Figure 1F), as estimated by luciferase signal. The loss of DRP1 more than tripled the median survival from 17 days to 55 days (Figure 1G; sgGFP vs. sgDrp1, P = 0.0074). These data were confirmed in one other sgDrp1 clone and sgGFP control (Supplemental Figure 1F). Knockout of DRP1 was retained in orthotopic tumors, as we verified via immunoblots (Supplemental Figure 1G). We also assessed the metastatic burden in these animals and found that sgDrp1 tumors were approximately 5 times less likely to develop macrometastases compared with sgGFP controls (Figure 1H; P = 0.0186).
We complemented these genetic studies with pharmacologic inhibition of DRP1 by the small-molecule DRP1 inhibitor, Mdivi-1 (11). Similar to DRP1 knockouts, Mdivi-1 promoted mitochondrial fusion, as confirmed by confocal microscopy with MitoTracker staining (Figure 2A). These treated KPC cells also exhibited decreased OCR (Figure 2B) as well as basal respiration, spare respiration capacity, and ATP production in a dose-dependent manner (Figure 2C). Furthermore, these metabolic changes corresponded to a dose-dependent decrease of in vitro cell growth (Figure 2D) and increased apoptosis, as determined by altered sub G0/G1 population on cell cycle analysis (Supplemental Figure 2A) and TUNEL staining (Supplemental Figure 2, B and C). We also observed reduced in vivo pancreatic cancer growth in a syngeneic flank model (Figure 2E).
Direct expression of MFN2 promotes mitochondrial fusion and decreases OXPHOS. To ensure that this phenotype was due specifically to the induction of mitochondrial fusion and not a pleiotropic alteration caused by DRP1 knockout or Mdivi-1 (12), we overexpressed MFN2 in KPC cells in a doxycycline-dependent manner (Tet-On-Mfn2 KPC, Figure 3A). After 48 hours, mitochondria were more elongated by MitoTracker staining compared with PBS controls (Figure 3B and Supplemental Figure 3A). To verify that these changes in mitochondrial morphology could also be observed in vivo, we implanted these syngeneic Tet-On-Mfn2 KPC cells into the flanks of C57BL/6 mice and analyzed the resultant tumors by electron microscopy (Figure 3C). Mitochondrial fusion was readily apparent in MFN2-overexpressing tumors, with a doubling of the average length of mitochondria (Figure 3C; Mfn2 OFF, 0.39 μm vs. Mfn2 ON, 0.84 μm, P = 0.0001). These data suggest that MFN2 expression directly regulates mitochondrial dynamics in vivo and are not limited to in vitro studies.
Direct mitochondrial fusion by MFN2 expression reduced OCR (Figure 3D), basal respiration, and ATP production (Figure 3E) compared with controls. This reduction in OXPHOS by MFN2 overexpression correlated with decreased cell proliferation in vitro (Supplemental Figure 3B) and enhanced G1 arrest (Supplemental Figure 3C). Furthermore, MFN2 overexpression reduced tumor volume (Figure 3F) and improved survival in a syngeneic orthotopic model (Figure 3G). Doxycycline alone exhibited no significant effects on OCR and did not decrease KPC tumor growth over a range of concentrations (Supplemental Figure 3, D–F). MFN2 expression in implanted tumors was confirmed by Western blot (Supplemental Figure 3G). MFN2 expression also decreased metastatic lung colonization after tail vein injection by more than 4-fold using 2 different Tet-On-Mfn2 clones (Figure 3H; representative H&E in Figure 3I).
Leflunomide activates MFN2 expression and improves survival in multiple mouse models of pancreatic cancer. Recently, the FDA-approved anti-arthritis drug, leflunomide, was found to also enhance the expression of MFN2 and promote mitochondrial fusion in HeLa cells (13). To determine whether leflunomide had similar activity in pancreatic cancer, we treated KPC cells with the drug and observed a 2-fold enhancement of Mfn2 mRNA (Figure 4A) and MFN2 protein levels (Figure 4B). Interestingly, we did not observe any changes in dihydroorotate dehydrogenase (DHODH) expression by leflunomide treatment (Supplemental Figure 4A). DHODH expression was also unchanged in sgDrp1 (Supplemental Figure 4B) and Tet-On-Mfn2 cells compared with controls (Supplemental Figure 4C). Fused mitochondria were more frequently observed in KPC cells (Figure 4C) and in implanted syngeneic KPC tumors (Figure 4D) after leflunomide treatment, which led to a doubling of the average mitochondrial length observed compared to vehicle (Figure 4D; vehicle, 0.46 μm vs. leflunomide, 1.14 μm, P = 0.007). This increased mitochondrial fusion was associated with reduced OCR (Figure 4E) and decreased basal respiration, spare respiratory capacity, and ATP production (Figure 4F). Leflunomide also suppressed in vitro growth of KPC cells and correlated with G2 cell cycle arrest (Supplemental Figure 5, A and B). Interestingly, we did not detect any changes in apoptosis by TUNEL staining (Supplemental Figure 5, C and D).
Oral leflunomide can be used to treat rheumatoid arthritis, but its effects on pancreatic tumors have not been studied extensively. Thus, we used leflunomide to treat pancreatic cancer in 3 different cancer models. First, we heterotopically implanted KPC cells into the flanks of recipient C57BL/6 mice and gavaged mice with leflunomide at a dose of 20 mg/kg per day and found that it decreased tumor growth and weight (Supplemental Figure 5, E–G) and improved median survival by almost 50% (Figure 4G; 25 vs. 36 days, vehicle vs. leflunomide, P = 0.004). None of the leflunomide animals exhibited overt toxicity to treatment, which is in contradistinction to previous studies that observed significant toxicity with intravenous leflunomide (14). We then tested leflunomide efficacy in a syngeneic orthotopic model, where the drug doubled the median survival of the treatment cohort compared to vehicle (Figure 4H; 28 vs. 56 days, vehicle vs. leflunomide, P = 0.012). Finally, orally administered leflunomide also demonstrated single-agent efficacy in an aggressive autochthonous KPC model of pancreatic cancer (Figure 4I), increasing the median survival by 50% (22 vs. 33 days, vehicle vs. leflunomide, P = 0.036).
Mitochondrial dynamics are dependent on Kras activity. Oncogenic KRAS plays a critical role in the maintenance of pancreatic tumor progression (15). To gain more insights into whether mitochondrial dynamics are regulated by the RAS/MAPK pathway, we used a doxycycline-inducible Kras (iKras) cell line (AK192), derived from iKras tumors (15). Induced Kras expression (Kras ON) caused a 4-fold increase in fragmented mitochondria (Figure 5A, 13.3% in Kras OFF vs. 56.7% in Kras ON, P = 0.0001). Similarly, treatment of KPC cells with MEK inhibitor (MEKi), PD0325901, yielded a 13-fold increase in elongated mitochondria compared with controls (3.7% vs. 51.3%; vehicle vs. MEKi, Figure 5B, P = 0.0002). MEKi treatment also decreased OCR (Figure 5C) as well as basal respiration and ATP production compared with vehicle controls (Figure 5D).
Kras-dependent response of leflunomide in patient-derived pancreatic cancer cells. To determine whether human pancreatic cancer patients also exhibited the KRAS-dependent mitochondrial morphology change we observed in murine KPC cells, we characterized 7 different patient-derived pancreatic cell lines (PATC), which were generated from resected pancreatic tumors at the MD Anderson Cancer Center, as has been described (16–18). We performed sequencing on these PATC lines and described the mutations in Supplemental Figure 6A. We noted that nearly all the PATC lines had an expected KRAS mutation. However, one cell line, MDA-PATC153, was KRAS WT with alternative driver mutations in TP53 and PIK3CA.
We performed MitoTracker staining on these PATC lines and used human pancreatic epithelial cells (HPNE) as a control for the morphology of normal tissues. We found that, as expected, normal HPNE cells exhibited fused, elongated mitochondria (Figure 6A). We stratified the rest of our analysis by KRAS mutation status. Interestingly, the KRAS WT line, MDA-PATC153, exhibited mitochondrial morphology similar to the normal HPNE cells and was not statistically different after quantification (Figure 6A and Supplemental Figure 6, B and C). The other PATC lines that harbored mutations in KRAS, however, exhibited marked mitochondrial fragmentation (Figure 6A, MDA-PATC 118, 50, 102, 53, 124, and 148).
To determine whether PATC cell lines respond to leflunomide treatment in a KRAS-dependent fashion, we first treated KRAS mutant MDA-PATC53 with leflunomide and found that the drug induced mitochondrial fusion, with 5 times more tubular mitochondria by MitoTracker staining in leflunomide-treated cells compared to vehicle controls (Figure 6B, 8.86% vs. 43.61%, vehicle vs. leflunomide, P = 0.02). Consistent with our previous findings, this stimulation of mitochondrial fusion decreased OCR (Figure 6C) and basal and spare respiration in addition to ATP production (Figure 6D) When we interrogated the KRAS WT cell line, MDA-PATC153, with leflunomide, we observed no significant changes in mitochondrial morphology (Figure 6E). By the same token, we did not observe any significant changes in OCR, although ATP production increased by about 25% (Figure 6, F and G).
Mitochondrial fusion promotes mitophagy, which proportionally reduces functional mitochondrial mass. Mitochondrial fusion is a known mechanism of organelle quality control (3); thus, we hypothesized that our treatments might be inhibiting OXPHOS simply by reducing mitochondrial mass. We calculated the total mitochondria present by electron microscopy and found that the average number of mitochondria decreased by at least 50% when we induced mitochondrial fusion. For instance, the mitochondrial number decreased after DRP1 ablation (33.1 vs. 15.7 in sgGFP vs. sgDrp1, P < 0.0001), overexpression of MFN2 (39.2 vs. 18.4 in Mfn2 OFF vs. Mfn2 ON, P = 0.01), and pharmacological promotion of fusion by leflunomide (23.9 vs. 8.6 in vehicle vs. leflunomide treatment, P = 0.003, Figure 7A).
We complemented our microscopic data by estimating mitochondrial mass with mitochondrial DNA (mtDNA) copy number (19). This common approach uses a quantitative PCR assay to estimate total mtDNA by copy numbers of genes transcribed from mtDNA, such as mt-Nd1 and mt-Nd2, the gene products of with which reside in complex I (Figure 7B and Methods). Mitochondrial fusion was associated with a 39% reduction in the complex I marker mt-Nd1 (41.4 vs. 25.1, sgGFP vs. sgDrp1; P = 0.0008) and a 51% reduction in mt-Nd2 (56.4 vs. 28.5, sgGFP vs. sgDrp1, P = 0.00001). We confirmed this reduction in mitochondrial gene expression by Western blot for a mtDNA-encoded NADH dehydrogenase subunit of complex I called mt-Ndufb8, which exhibited a 48% decrease in expression (Figure 7, C and D; P = 0.007, sgGFP vs. sgDrp1). Importantly, mitochondrial fusion caused by direct MFN2 overexpression strongly suppressed complex I expression, as detected by immunoblot of the complex I protein, mt-Ndufb8 (Figure 7E).
We hypothesized that this observed loss of mitochondrial mass could have come from reduced mitochondrial biogenesis or altered turnover. To assess the former possibility, we measured expression of Ppargc1a, which is a critical regulator of this process (20). We found that Ppargc1a expression levels were unchanged by loss of Drp1 or overexpression of Mfn2 (Supplemental Figure 7, A and B). We additionally found no significant differences in expression of nuclear encoded genes involved in the electron transport chain, such as Ndufs3 and Tfam, which indicated that mitochondrial fusion was likely not inducing global problems with mitochondrial gene transcription (Supplemental Figure 7, C and D). Together, these data suggest that global defects in mitochondrial biogenesis do not explain the reduced mitochondrial mass after mitochondrial fusion.
Since mitochondrial biogenesis appeared to be unaffected, we investigated the possibility that mitochondrial fusion might stimulate mitochondrial turnover, possibly through mitophagy (21). We estimated mitophagy using an immunofluorescence assay, as has been described previously (20). Mitochondria from sgDrp1 or control sgGFP cells were stained with Tom20 and colocalized with an antibody for the autophagosome marker, microtubule-associated proteins 1A/1B light chain 3B (LC3). We found that enhancing mitochondrial fusion by loss of DRP1 was associated with a more than 2-fold increase in colocalization of Tom20/LC3 in sgDrp1 cells compared to controls (Figure 7, F and G; 12.8% vs. 29.5%, sgGFP vs. sgDrp1; P = 0.017). These data suggest that mitochondrial fusion promotes mitophagy, which then culls the excess mitochondria in pancreatic cancer cells (Figure 7H).
More information: Sarbajeet Nagdas et al, Drp1 Promotes KRas-Driven Metabolic Changes to Drive Pancreatic Tumor Growth, Cell Reports (2019). DOI: 10.1016/j.celrep.2019.07.031
Journal information: Cell Reports
Provided by University of Virginia