Fumarate Hydratase

Author: Annalisa Ghiglia
Date: 06/11/2009



Fumarase (or fumarate hydratase) is an enzyme that catalyzes the reversible hydration/dehydration of Fumarate to S-malate.

Two classes of fumarases are described:
• class I: Fumarase enzyme catalyses the reversible hydration of fumarate to L-malate. This class comprises thermolabile dimeric enzymes
• class II: Fumarase enzyme comprises thermostable and tetrameric enzymes, which are found both in prokaryotes and in eukaryotes (human).
The sequences of the two classes are not closely related.

The enzyme is widespread in both foetal and adult tissues with most abundant expression in skin, parathyroid, lymph and colon (four highest NCBI expression profiles). In addition it is expressed in malignant tumours such as leiomyosarcoma and papillary (type II) renal cell carcinoma.

Enzyme Structure
It is a homotetrameric enzyme composed by four identical subunits. In the structure three active DNA binding sites (site A) and one lower affinity substrate (site B) have been identified. The protein has been purified, and the crystal structure has been resolved.

The enzyme exists in a cytosolic and a mitochondrial form (2 isoforms produced by alternative initiation). Isoenzyme products have nearly identical amino acid sequences, but vary at the amino terminus and for electrophoretic mobility.
The Isoform Mitochondrial (identifier: P07954-1¸ Length 510¸ Mass 54,637 Da) has an extended N-terminal, differing only in the translation start site used. Once in the mitochondrion, the extension is removed, generating the same form as in the cytoplasm.
The sequence of the Isoform Cytoplasmic (identifier: P07954-2, Length 467¸ Mass 50,213 Da) differs from the mitochondrial as follows: 1-43 (MYRALRLLAR SRPLVRAPAA ALASAPGLGG AAVPSFWPPN AAR): Missing. Note: Initiator Met-1 is removed. Contains N-acetylalanine at position 2.

Fumarase (FH) gene (10 exons; 22,152 bp) has been cloned and sequenced (Gene map locus 1q42.1).

Mutations cause fumarase-deficiency and lead to progressive encephalopathy as well as uterine fibroids, skin leiomyomata and papillary renal cell cancer.
Germline mutations are associated with two distinct conditions:
• Homozygous and compound heterozygous mutations (e.g., missense and in-frame deletions) of the 3' end result in fumarate hydratase deficiency (FHD).
• Heterozygous 5' mutations (e.g., nonsense, missense and deletions ranging from one base pair to whole gene) predispose individuals to somatic mutations in the normal allele leading to hereditary leiomyomatosis and renal cell carcinoma/multiple cutaneous and uterine leiomyomatosis (HLRCC/MCUL1).
Somatic mutations: loss-of-heterozygosity of the wt allele results in functional nullizygosity for fumarate hydratase. Malignant uterine and kidney tumors characteristic of HLRCC can subsequently develop.

Entrez GeneURL



Protein Aminoacids Percentage


mRNA synthesis
1,1790 bp. Multiple RNA transcripts encode two FH gene products- one with a mitochondrial signal protein and the other lacking the signal sequence.

protein synthesis
Doonan et al. (1984) cited evidence suggesting that the isoenzymes of fumarase are translated in \precursor form from 2 different mRNA molecules, these mRNAs in turn arising from alternative splicing of a single gene transcript.

post-translational modifications
Isoform Cytoplasmic is acetylated at position 2 (Acetylation)


cellular localization,
Mitochondrial and cytosolic.
Subcellular localization is determined by presence or absence of a signal sequence at the amino terminus (Transit peptide). Presence of the signal generates the mitochondrial-targeted form while absence of the signal results in the cytosolic form.

biological function

  • Enzymes
    Fumarase plays a key enzymatic role in fundamental metabolic pathways. The mitochondrial isoenzyme catalyzes conversion of fumarate to malate in the Krebs, or tricarboxylic acid (TCA) cycle, in which acetyl-CoA produces CO2, reduced electron carriers (FADH2 and NADH) and ATP.

The cytosolic isoenzyme is involved with amino acid metabolism (see figure below).

  • Cell signaling and Ligand transport
  • Structural proteins


There are 2 substrate binding sites: the catalytic A site, and the non-catalytic B site that may play a role in the transfer of substrate or product between the active site and the solvent. Alternatively, the B site may bind allosteric effectors.


Fumarate hydratase as a mitochondrial tumor suppressor gene

The neoplastic transformation and mitochondrial tumor suppressor genes
The cancer phenotype is influenced by both cell-autonomous and environment-dependent characteristics. Despite the apparent complexity of this phenotype, almost all types of human cancers share six essential alterations, which influence cell physiology and dictate malignant growth. These alterations include self-sufficiency in growth signals, insensitivity to growth-inhibitory signals, evasion from programmed cell death (apoptosis), limitless replicative potential, sustained angiogenesis and tissue invasion and metastasis (fig.1) (1).

Fig.1 Acquired capabilities of cancer cells Hanahan D. and Weinberg A. Cell, Vol. 100, pag. 58 January 7 2000.

More recently, changes in bio-energetic processes have been recognized as an additional cancer-hallmark (2). Indeed, the survival and proliferation of cancer cells depend predominantly on glycolysis more than on respiration. This effect, known as “Warburg effect”, consists in an important increase of glycolysis and lactate production in cancer cells, even in the presence of oxygen and without an accompanied increase in respiratory chain production of energy.
The switch from respiration to glycolysis is usually considered a consequence rather a cause of cancer. However, the recent discovery of mitochondrial tumor suppressors has changed this viewpoint. Mitochondria are traditionally considered as energy-producing organelles, as they generate approximately 90% of the cellular ATP through the respiratory chain and the tricarboxylic-acid (TCA) cycle (Krebs cycle). Therefore, mitochondrial defects result in severe disease affecting the entire organism, or at least the organs that have the highest energy consumption (central nervous system and heart). Interestingly, it has been discovered that mutations in mitochondrial enzymes are directly implicated in hereditary neoplasias. Mutations in the fumarate hydratase (FH, also known as fumarase) gene lead a predisposition to the onset of multiple cutaneous and uterine leiomyomata (MCUL) and to the variant syndrome in which leiomyomata are associated to kidney cancer (HLRCC) (7-10,11). Moreover, germline heterozygous mutations in three nuclear genes encoding subunits of succinate dehydrogenase (SDHB, SDHC, SDHD) cause the inherited syndromic phaeochromocytoma and paraganglioma (3-6). Both of the enzymes encoded by these genes are part of the Krebs cycle, which connects glucose metabolism in the cytosol to oxidative phosphorylation in the mitochondria.
Because of their key role in cellular metabolism, the genes encoding SDH and FH were considered ‘housekeeping genes’, which are ubiquitously expressed at high level in all tissues, making it difficult to understand how they can be tumor suppressors. This aspect (for the FH gene) will be treated in detail in the next paragraphs.

Fumarate hydratase as a tumor suppressor gene

Germline mutations in FH gene are associated with two distinct conditions. In the homozygous state, bi-allelic FH mutations cause the autosomal recessive fumarase deficiency syndrome (also known as fumaric aciduria). This is characterized by gross developmental delay, severe neurological impairment and death in the first decade. The affected individuals show a very low FH enzymatic activity (12-14). In the heterozygous state, FH acts as a tumor suppressor gene (9), with germline mutations causing an autosomal dominant syndrome of multiple leiomyomata (smooth muscle tumors) of the skin and uterus (Multiple Cutaneous and Uterine Leiomyomatosis, MCUL). MCUL has also been associated with an increased risk of type II papillary renal cancer and uterine leiomyosarcoma (7, 15). This variant has been termed Hereditary Leiomyomatosis and Renal Cell Cancer (HLRCC). In MCUL patients, smooth muscle tumors consist in multiple and early-onset benign lesions arising from either the erector pili muscles of the skin or the myometrium of the uterus. The renal cell cancer reported in MCUL/HLRCC families occurs with minor frequency than uterine and skin leiomyomas and shows variable morphology, including type II papillary, and in some cases collecting duct and clear cell lesions. These renal tumors are early onset lesions usually found as solitary and unilateral tumors, which invade locally and metastasize to distant organs. Uterine leiomyosarcomas also occur infrequently in MCUL/HLRCC but are highly malignant and occur at young age.
The observation that homozygous germline mutations in FH lead to neurodegeneration, without cancer, whereas heterozygous germline mutations in FH lead to cancer susceptibility is at first puzzling. This may be explained by the role of FH in metabolism. Indeed, FH plays an essential role in TCA cycle, by catalyzing the conversion of fumarate to malate. At the organismal and organ level, homozygous mutation in FH leads to complete or near-complete loss of activity in every cell of the body throughout development and shortly after birth, crucial times in neurological development. Given to particularly high oxygen and energy demands in the developing nervous system, it becomes plausible that the chronic lack of energy due to fumarase inactivation would lead to severe neurological dysfunction. By contrast, heterozygous germline mutations only reduce dosage, which however remains adequate for normal neurological development. A following somatic mutation in residual wild type allele leads to a near complete loss of enzymatic function in the transforming cells that comprise the tumor. LOH (loss of heterozygosity) on chromosome 1q42, where the FH gene lies, as well as several acquired somatic mutations have been observed in uterine and skin leiomyomas, and in papillary RCC from patients with HLRCC (7-9, 15, 17-18).
The spectrum of FH mutations associated to MCUL/HLRCC and FH deficiency includes missense mutations, insertion/deletion of highly conserved amino acids localized in or around the active site of the enzyme, nonsense mutations that are predicted to truncate the protein, along with several whole-gene deletions (fig.2).

Fig.2 The FH gene and disease-associated mutations: The yellow bars represent the structure of FH gene, with the numbers denoting the exons. Heterozygous mutations, which give rise a predisposition to cancer are noted in the upper side of FH gene structure, whereas mutations associated with FH deficiency are shown in the lower side. In addition to the germline changes, the figure includes somatic mutations (s) detected in tumors.

FH mutations for HLRCC are found throughout the entire gene with no particular genotype-phenotype correlations, while the mutations leading to FH deficiency tend to be located in the 3’ end of the gene. More truncating and whole-gene deletion mutations are present in HLRCC than in fumarase deficiency (9, 15). It is possible that some type of mutations with severe effect (such as truncations/deletions or N-terminal changes) are under represented in fumarate hydratase deficiency, because they are lethal in uterus.
Interestingly, some MCUL patients with missense germline mutations of FH present more severely reduced enzymatic activity than patients with truncating mutations, including a whole gene deletion (9). The most plausible explanation is a dominant negative activity of the missense mutant since fumarase functions as a homotetramer. It is conceivable that mutations in FH might affect multimerization, resulting in defective catalytic activity and affecting other putative functions of FH. This hypothesis has been confirmed by Lorenzato and colleagues in 2007 (16).

Hypothesis that link FH loss-of-function mutations to tumorigenesis

Several mechanisms have been proposed to explain how loss of fumarase function leads to tumorigenesis. These include:
* the activation of hypoxia-like pathway under normoxic conditions (pseudo-hypoxia),
* the decrease in programmed cell death (apoptosis),
* the increase in the production of reactive oxygen species (ROS).

These mechanisms may be not mutually exclusive but rather they can overlap and interact with each other. At present, the main part of reported data support a role of pseudohypoxic drive in tumor development.

The pseudo-hypoxia model implies a link between inactivation of FH and the induction of a hypoxic response under normoxic conditions. This response is mediated by the oxygen-regulated HIF transcription factor.
Hypoxia-inducible factor (HIF) is a heterodimeric complex made of HIF-α and HIF-β subunits (19). Three genes encode the three different α-subunits of HIF-α (HIF-1α, HIF-2α, and HIF-3α) and two genes encode the HIF-β subunits (HIF-1β/ARNT1 and ARNT2) (20). HIF-α subunit, together with HIF-β, forms an active transcription complex that up-regulates numerous target genes. The physiological function of HIF is to promote adaptation of cells to low oxygen conditions (hypoxia). This is achieved by inducing glycolysis as an anaerobic alternative to oxidative phosphorylation and by inducing blood vessels growth to facilitate oxygen and nutrient supply into hypoxic tissues (20). The adaptation to hypoxia is mainly mediated by the transcription of HIF-1α regulated genes, while tumor growth and survival are mainly regulated by HIF-2α.
HIF-1α (or in some cases HIF-2α) is overexpressed in a majority of primary tumors, cancer cell lines, and metastases (21-23). While the HIF-β subunits are constitutively expressed, the HIF-α subunits are labile under normoxic conditions, due to proteasomal degradation following their oxygen-dependent ubiquitination by an ubiquitin ligase complex targeted by the von Hippel-Lindau (VHL) protein (24-25). VHL-mediated recognition of HIF requires the enzymatic hydroxylation of two conserved proline residues. This reaction is mediated by HIF-α prolyl hydroxylases (1-3-PHDs, also known as EglNs 2, 1, 3 respectively, or HPHs 3-1) (26-28). PHD activity requires ascorbate and iron as cofactors and α-ketoglutarate and molecular oxygen as co-substrates. This model explains the basis for HIF stabilization under hypoxia. PDH function and following VHL recognition of hypo-hydroxylated HIF is compromised in the absence of oxygen. Once stabilized, HIF-α translocates from the cytosol to the nucleus, where it can heterodimerize with HIF-β to form an active transcription factor, leading the expression of target genes, that play a role in glycolysis, angiogenesis, metastasis and survival (29).
Several lines of evidence connect loss of fumarase to HIF accumulation and activation in both leiomyomas and renal tumors from HLRCC patients. Pollard and colleagues examined HIF-1α expression in kidney tumors from HLRCC patients (30). In both papillary and collecting duct tumors, the authors noted a strong nuclear HIF-1α staining. This data was also confirmed by Western blot, as HIF-1α was easily detected in tumor cell lysate, with no detectable HIF-1α in normal kidney lysate [78]. Moreover, Isaacs and coworkers also examined HIF-1α expression in kidney tumors from HLRCC patients (31). Both HIF-1α and HIF-2α, evaluated by immunohistochemistry, were found to be significantly increased in HLRCC renal tumors, but HIF-1α expression seemed to be preferentially increased compared to HIF-2α. Altogether these findings were compelling for activation of hypoxic pathways in kidney tumors. Further supportive evidence comes from observation of the upregulated downstream HIF targets in tumor tissues isolated from patients with HLRCC.
In addition, Pollard et al. (32) examined microvessel density in HLRCC uterine leiomyomas. Vascular density of the vascular endothelial markers determined by immunohistochemistry was significantly higher in HLRCC uterine leiomyomas as compared to non-leiomyomatous myometrium from HLRCC women. Furthermore, there was a statistically significant higher vascular density in HLRCC leiomyomas compared to sporadic leiomyomas or normal myometrium from women without HLRCC. Accordingly to these results, in situ hybridisation studies revealed up-regulation of VEGF transcripts in HLRCC uterine leiomyomas with respect to normal myometrium (from HLRCC and non-HLRCC women) as well as sporadic uterine leiomyomas. These findings were confirmed by quantitative real-time PCR data that revealed enhanced expression of VEGF (1.4-3.5 folds) as compared to normal myometrium. In addition, other hypoxia-responsive gene changes were also found in HLRCC leiomyomas including down-regulation of TSP1, a known anti-angiogenesis factor (33). Further investigation revealed moderate HIF-1α expression in HLRCC fibroids and a remarkable VEGF increase (30). The concomitant up-regulation of these proteins suggests that pseudohypoxic activation contributes to the genesis of uterine leiomyomas in HLRCC patients.
There is also supportive evidence for activation of hypoxic pathways in kidney tumors from HLRCC patients. Isaacs et al. identified enhanced GLUT1 expression by immunohistochemistry in multiple HLRCC kidney tumors as compared to normal renal tissue (31).
Although these works provided evidence for activation of pseudo-hypoxia, demonstrated by high HIF activity under normoxic conditions in HLRCC tumors, they did not provide the molecular mechanism for this phenomenon.
A different work reported the biochemical explanation for the pseudoxypoxic drive induced by SDH enzyme dysfunction, indicating a possible parallel mechanism for HIF stabilization due to FH inactivation (34). The authors demonstrated that succinate, the substrate of the SDH enzyme, can cause HIF stabilization in the test tube, by interfering with PHD activity. Succinate is a dicarboxylic acid, capable of crossing the mitochondrial inner membrane (the only barrier to small metabolites in the mitochondria) via the dicarboxylate carrier. Selak et al. showed that SDH dysfunction in cells raised the levels of succinate and, that succinate serves as an intracellular messenger between the mitochondria and the cytosol. Succinate that accumulates in the mitochondrial matrix owing to SDH dysfunction, leaks out in the cytosol where it inhibits the activity of HIFPHD. As mentioned before, HIF-α prolyl hydroxylation by PHD is the first step in tagging HIF-α for degradation, enabling its interaction with pVHL. To hydroxylate HIF-α , prolyl hydroxilase (PHD), requires oxygen and α -ketoglutarate as substrate, and ferrous iron (Fe2+) and ascorbate as co-factors. PHD catalyses two coupled reactions: the HIF-α prolyl hydroxylation and the decarboxylation of α-ketoglutarate to succinate (fig.3).

Fig.3 Pseudo-hypoxic drive: the pseudo-hypoxia model entails a link between inactivation of fumarase and the induction of hypoxic response under normoxic conditions.This model implies that accumulated succinate and fumarate, due to FH loss of function, could leave mitochondria and inhibit the activity of a prolyl hydroxylase in the cytosol, that is responsible for HIF-α hydroxylation. In normoxic conditions HIF-α, after the modification given by this enzyme, can bind to pVHL protein, became polyubiquitylated and degradated. However, if this enzyme is inhibited by fumarate, HIF-α is not hydroxylated, and so can escape degradation even under normoxic conditions.Thus HIFα can accumulate and traslocate into the nucleus where together with HIF-β, forms an active complex that induces the expression of genes that support tumor growth and spreading.

Thus succinate is not only a substrate for SDH in the mitochondria, but also a product of PHD, in the cytosol. Therefore, the accumulated succinate, by feedback inhibition of PDH, leads to HIF-α stabilization and activation of HIF complex.

By analogy, the absence of FH could presumably result in chronically elevated levels of fumarate and altered levels of other TCA intermediates, which could drive cells to a pseudo-hypoxic state. The direct link between FH-deficiency and HIF activation was described subsequently (31). It was shown that, like succinate (and potentially even better), fumarate, whether induced pharmacologically or by molecular knockdown of FH gene with small interfering RNA (siRNA), is able to inhibit PHD activity, by competing with the cosubstrate α-ketoglutarate, and so cause HIF accumulation and activation. Fumarate is not a product of PHD, but is chemically similar to succinate. Furthermore, it has been reported that also other Krebs cycle intermediates, as oxaloacetate and pyruvate stabilize HIF-1α in cultured cancer cell lines and inactivate HIF-PHDs in a way that is reversible by ascorbate (35, 36). A recent work also demonstrated in vitro, that all three Krebs cycle intermediates fumarate, succinate and oxalacetate inhibit the three HIF-PHDs, while citrate is an effective inhibitor of only PHD-3. Nevertheless, citrate inhibits the HIF-α asparaginyl hydroxylase FIH, which blocks HIF interaction with the transcriptional coactivator p300. However, in this work they did not find PHDs inhibition by pyruvate (37).
Further support for the biochemical role of succinate and fumarate in FH deficient cells and tumors came from studies of these tumors (30). It was clearly demonstrated that both succinate and fumarate levels are high in HLRCC tumors and FH deficient cells. The increase in these metabolites coincided with high levels of HIF-α protein and HIF-α targets.
The experimental evidence that pseudo-hypoxic drive, resulting from HIF-1α and HIF-2α overexpression, is a direct consequence of FH inactivation came from a recent in vivo work (38). In this study, Pollard et al. showed that the conditional inactivation of mouse Fh1 in the kidney, leads to the development of multiple clonal renal cysts that overexpressed Hif-1α and Hif-2α, and Hif target genes, such as Glut1 and Vegf. Notably these cysts did not undergo progression to malignancy.
All these works suggest that fumarase deficiency causes HIF stabilization and activation in HLRCC tumors; it is conceivable that activation of pseudo-hypoxia confers a selective advantage to HLRCC tumor cells.
It is well known that hypoxic regions of tumors provide a favorable environment for the selection of aggressive cells (39). In addition, the ability of cells to survive and evolve under hypoxic conditions is largely dependent on HIF. Nevertheless, there is insufficient evidence to indicate that the induction of HIF under normoxic conditions can, by itself, support tumor development. However, it is likely that HIF induction, in combination with other lesions, will facilitate or enable tumor progression. Mechanisms that might account for HIF-mediated tumorigenesis include the induction of genes that facilitate neovascularization, stimulate aerobic glycolysis, or directly block apoptosis. While an increased blood supply provides sufficient nutrients to the tumor, aerobic glycolysis in the absence of faulty oxidative phosphorylation can increase energy production, enabling cells to proliferate and survive in stressful environments (40).
Blockade of the TCA cycle by genetic inactivation of key enzymes such as FH and SDH, should enhance cellular reliance upon glycolysis and give a selective advantage to cells demonstrating up-regulation of this pathway. A direct experimental evidence supporting this hypothesis was provide by Isaacs and colleagues, who demonstrated that glucose and lactate levels raise in cells following FH siRNA-dependent doubling of intracellular fumarate (31).
The role of HIF in directly regulating apoptosis is less clear. Although in some cases HIF was shown to induce pro-apoptotic genes such as BNIP3 (41), it has also been reported that HIF can reduce apoptotic sensitivity by directly repressing transcription of another pro-apoptotic gene, BID (BH3-interacting domain death agonist) (42). HIF-regulated apoptosis is probably cell-type specific and might also be dependent on the microenvironment of the tumor (43). At this regard, even if it was reported an HIF-stabilization and -activation in FH-deficient cells and in HLRCC tumors, any experimental evidence till now, demonstrated a role of HIF in regulating apoptosis of cells where FH function was loss. Although fumarate-mediated HIF up-regulation, coupled with inactivated FH-driven adaptation to glycolysis, together create an environment permissive for tumorigenesis, further experimentation is needed to fully explore the link between deregulation of the TCA cycle and tumorigenesis and to more thoroughly elucidate the roles of HIF and PHD in this process.

Decrease in the programmed cell death (apoptosis):

Another theory for the development of tumors as a result of FH mutations is a decreased tendency of cells to undergo apoptosis, caused by some form of mitochondrial dysfunction.
Deficiency in apoptosis, the cellular intrinsic mechanism of self destruction, has been demonstrated as a crucial step in tumorigenesis. Indeed, a defective suicide program endows nascent neoplastic cells with multiple selective advantages. The cells can persist in hostile niches (for example where cytokines or oxygen are limiting), escape the death, which is often imposed as a fail-safe mechanism by other oncogenic changes, and evolve into more-aggressive derivatives. Finally, defective apoptosis facilitates metastasis, because the cells can ignore restraining signals from neighbors and survive detachment from the extracellular matrix. So, neoplastic progression largely requires loss of normal apoptotic mechanism. Impaired apoptosis is also a significant impediment to cytotoxic therapy (44). The mutations that favored tumor development dampen the response to chemotherapy and radiation, and treatment might select more refractory clones.
Apoptosis can be initiated by two alternative mechanisms: one is mediated by death receptor on the cell surface, sometimes referred to as “extrinsic pathway”; the other is referred to as the “intrinsic pathway”, and is mediated by mitochondria. In addition to their crucial role in bioenergetics, mitochondria are in fact widely recognized as essential subcellular organelles in the regulation of apoptosis. Several apoptogenic proteins normally reside in the mitochondria, such as cytochrome c, apoptosis inducing factors (AIF), endonuclease G, DIABLO. In response to an appropriate signal, these apoptogenic factors are released from the mitochondria to activate apoptosis.
In the intrinsic apoptosis pathway, mitochondria play a central role since they integrate and propagate the death signal originating inside the cell, such as DNA damage, oxidative stress, starvation as well as those induced by chemotherapeutic drugs (45). In this pathway the members of the Bcl-2 family are important regulators. These are a group of homologous proteins that act either to promote or suppress apoptosis. In addition, they are implicated in the control of outer mitochondrial membrane (OMM) permeability, which in turn controls the release of proapoptotic factors from the mitochondrion into the cytoplasm (46). Thus, pro-apoptotic BCL2 family proteins (BAX, BID, BAD and BIM) are important mediators of death signals, and their activation, following apoptotic stimuli, leads to the release of apoptogenic factors as cythocrome c (Cyt c) from the intermembrane space into the cytosol. Concomitantly, the mitochondrial transmembrane potential drops.
In the cytosol, cytochrome c binds apoptotic protease activating factor 1 (APAF1), ATP and the inactive initiator caspase pro-caspase-9 to form the apoptosome (fig.4).

Fig.4 Apoptosis signaling through mitochondria: DNA damage, oxidative stress, starvation and other stimuli can initiate apoptosis through the mitochondrial (intrinsic) pathway. Pro-apoptotic BCL2 family proteins are important mediators of these signals. Activation of mitochondria leads to the release of Cyt c into the cytosol, where it binds apoptotic protease activating factor 1 (APAF1) to form the apoptosome. At the apoptosome, the initiator caspase-9 is activated. Apoptosis through mitochondria can be inhibited at different levels by antiapoptotic proteins, including the anti-apoptotic proteins BCL2 family members and inhibitors of apoptosis proteins (IAPS), which are regulated by SMAC/DIABLO. Another way is through survival signals, that activate the phosphatidylinositol 3-kinase (PI3K) pathway. PI3K activates AKT, which phosphorylates and inactivates the pro-apoptotic BCL2-family member BAD.

Within this complex caspase-9 is activated. Once the initiator caspases are activated, they cleave and activate “executioner” caspases, mainly caspase-3, 6 and 7. Then, the active caspases cleave each other beginning an amplifying proteolytic cascade of caspase activation is (47). Eventually, the active caspases cleave cellular substrates, the “death substrates”, which leads to characteristic biochemical and morphological changes (47). The cleavage of ICAD (the inhibitor of the DNase CAD, caspase-activated deoxyribonuclease) causes the release of endonuclease, which travels to the nucleus to fragment DNA. The cleavage of cytoskeletal proteins such as actin, plectin, Rho kinase 1 (ROCK1) and gelsolin lead to cell fragmentation, blebbing and formation of apoptotic bodies. After exposure of “eat me signals” (for example, exposure of phosphatidyl serine and changes in surface sugars), the remains of the dying cells are engulfed by phagocytes (48).
Apoptosis through mitochondria can be inhibited at different levels by anti-apoptotic proteins, including the anti-apoptotic BCL2 family members BCL2 and BCL-XL and inhibitors of apoptosis proteins (IAPS), which are regulated by SMAC/DIABLO. Survival signals, such as growth factor and cytokines, function by activating the phosphatidylinositol 3-kinase (PI3K) pathway. PI3K activates AKT, which phosphorylates and inactivates the pro-apoptotic BCL2-family member BAD.
Besides playing a central regulator function in the intrinsic apoptosis pathways, mitochondria in some type of cells can also mediate extrinsic apoptotic pathways. In these cells the signal coming from the activated receptor does not generate a caspase activation strong enough for execution of cell death on its own. Thus, the apoptotic signal is amplified via mitochondria-dependent apoptotic pathways. As mitochondria have a central role in orchestrating many apoptotic processes, and also mitochondrial physiology is affected by several master regulators of apoptosis (49-50) it is conceivable that TCA cycle dysfunction might give rise to apoptosis-resistant cells, thereby contributing to tumor development.
Mitochondrial-mediated apoptosis, which is regulated by BCL2 proteins and involves the activity of caspases, is also known as energy-dependent apoptosis. Pro-caspases are indeed cleaved to caspases in order to mediate cell death, in an energy-dependent reaction.
It is possible that partial or complete loss of FH function leads to energy depletion that could cause an inefficient energy-dependent apoptosis (51). Mitochondrial-mediated apoptosis can be also energy-independent. This process is mediated by oxygen free radicals. When mitochondrial function is impaired, severe energy deficits occur and large amounts of oxygen free radicals are generated. When mitochondria sense the presence of oxygen free radicals, hypoxia inducible factors (HIFs), such as HIF-1α, are activated and translocated to the nucleus where they induce gene expression. HIF-1α gene targets encode proteins that promote cellular proliferation or can prevent apoptosis, leading to neoplasia (51).
Another possible explanation for a hypothetic apoptosis-inhibiting effect of mitochondrial dysfunction might be the upregulation of glycolysis as an alternative energy producing metabolic pathway. Because of the inactivation of the TCA cycle, oxidative phosphorylation will occur at sub-optimal levels. This is a problem because tumor cells have increased metabolic requirements. When oxidative phosphorylation is impaired, glycolytically generated ATP becomes the only other source of energy in tumor cells (52). Once induced, glycolytic enzymes could regulate other cellular processes that include the blocking of apoptosis (53, 54). For example mitochondria-associated hexokinases, key glycolytic enzymes, have been identified playing a role in the control of mitochondrial phase of apoptosis (55, 56). Indeed, some hexokinase isoenzymes (HKI and HKII) present porin-binding domain and are able to bind to the mitochondrial PTPC (permeability transition pore complex) member, VDAC (voltage dependent anion channel), and so inhibiting its apoptotic function (54, 57). This mitochondrial translocation is finely modulated by the anti-apoptotic serine/threonine kinase Akt that is required to maintain hexokinase association with mitochondria (54, 58). This suggests that, in cancer cells, elevated levels of mitochondria-bound isoforms of hexokinase can result in apoptotic evasion, allowing the cells to proliferate. Therefore, by enabling cells to increase glucose metabolism, it is plausible that inactivation of mitochondrial tumor-suppressors might contribute to a persistent anti-apoptotic effect. It is also plausible that beyond its traditional enzymatic functions in Krebs cycle, FH might have other independent functions. A good model that might shed some light on this possibility comes from studies of the apoptosis-inducing factor (AIF). AIF is believed to induce nuclear apoptosis independent of the caspases. On apoptosis induction, AIF is released from the mitochondrial inter-membrane and translocates to the nucleus (59), where it causes peripheral-chromatin condensation and DNA fragmentation. In addition to these apoptotic function, AIF can independently catalyse the reduction of cytochrome c in the presence of NADH (60), and stably bind FAD. Thus, AIF is a flavoprotein that has oxydoreductase activity. It is also possible that FH can act independently of its Krebs cycle functions to mediate apoptosis directly.
An alternative mechanism, is that blockage of the TCA cycle at FH may cause accumulation of metabolites from previous step of the cycle and other substances such as the amino acids glutamate and glutamine, derived from a-ketoglutarate. Both glutamate and glutamine play an important role in the resistance of cells to apoptosis and also in promoting cell proliferation (61, 62).
More direct evidence for the role of mitochondrial tumor suppressors in apoptosis has recently emerged from studies of VHL (63). Lee et al. found that apoptosis in phaecromocytoma cells is mediated by c-Jun, which activates PHD3 (EglN3). To activate c-Jun, pVHL eliminates atypical-PKC activity that induces the transcription of JunB, the upstream inhibitor of c-Jun. They also showed that PHD3 is an important target for succinate-mediated inhibition in SDH-deficient cells and the accumulation of succinate impairs PDH3-induced apoptosis. However, it remains unclear which PHD3 substrates cause the apoptotic response. It is possible that several targets of PHDs, including HIF-a, and apoptotic inducing factors, contribute to tumor progression owing to loss of mitochondrial tumor suppressor genes.
To date, no evidence of abnormal mitochondrial number, structure or functions have been reported in MCUL/HLRCC (64). In addition, no experimental evidence has linked mitochondrial dysfunction due to FH loss of function and impaired apoptosis. Only in a recent work, Wortham and colleagues (64) observe an alteration of the expression of the Bcl2 proteins, which regulate apoptosis at mitochondrial level, in tumors caused by FH mutations. They revealed by immunohistochemistry an increase in expression of antiapoptotic Bcl-2 in both sporadic and HLRCC uterine leiomyomata. Furthermore, they observe an increase in antiapoptotic Bcl-XL and a concurrent decrease of in proapoptotic Bak solely in HLRCC leiomyomas. These alterations in expression of apoptosis-related proteins indicate a shift in both HLRCC and sporadic leiomyomas to increased resistance to apoptosis compared with myometrium. Moreover, these data also show that in HLRCC leiomyomas the survival signals may be stronger than in sporadic leiomyomas.

Increase in the production of reactive oxygen species (ROS):

Owing to TCA cycle blockage and a possible abnormal respiratory chain function, accumulation of oxygen free radicals may occur in MCUL/HLRCC tumors, leading to an increased of oxidative stress and hypermutation.
The reactive oxygen species (ROS) can be generated by electron transport chain (ETC), by the transfer of a single electron to molecular oxygen. The exact mechanism of ROS generation in the mitochondrial ETC is still discussed, although it is fairly well established that the important sites for ROS generation are complex I (NADH-ubiquinone oxidoreductase) and complex III (ubiquinone- cytochrome c oxidoreductase) (65).
The role of ROS in pathogenesis of HLRCC tumors is still being debated. It has been proposed that DNA mutations that result from oxidative damage could promote tumorigenesis given by FH mutations, but this remains unproven. In addition of mutagenesis, ROS might play another role in the pathology of tumor due to FH mutations. It has been reported that reactive oxygen species can inhibit PHD activity under normoxic conditions by oxidizing the PHD cofactors ferrous iron and ascorbate (66). So ROS could play a role in the pseudohypoxic response of FH-deficient tumors.
However, redox stress was not measured in FH deficient cells (31) and pseudo-hypoxia was found also in the presence of effective anti-oxidants (67). Recently, it has also been demonstrated (68) that increased HIF-1a in SDH and FH deficient tumors does not cause microsatellite instability and thus increased mutation rate.
Unlike SDH enzyme, which constitutes the complex II of the electron transport chain, FH does not play a role in the ETC and therefore there is not direct link with increased free radical production and oxidative damage of DNA.
Although probably ROS do not contribute to the tumorigenic effect of FH inactivation further studies are needed to clarify this point.

In conclusion several mechanisms have been proposed to explain how loss of function of FH enzyme could lead to tumor development. These include the activation of a hypoxia like pathway under normoxic conditions, an inefficient apoptotic response, and an increase in the production of oxygen reactive species. So far the majority of data support a role for pseudohypoxic drive in FH loss of function tumorigenesis.


1 Hanahan, D. and R.A. Weinberg, The hallmarks of cancer. Cell, 2000. 100(1): p. 57-70.
2. Garber, K., Energy deregulation: licensing tumors to grow. Science, 2006. 312(5777): p. 1158-9.
3. Baysal, B.E., et al., Mutations in SDHD, a mitochondrial complex II gene, in hereditary paraganglioma. Science, 2000. 287(5454): p. 848-51.
4. Gimm, O., et al., Somatic and occult germ-line mutations in SDHD, a mitochondrial complex II gene, in nonfamilial pheochromocytoma. Cancer Res, 2000. 60(24): p. 6822-5.
5. Astuti, D., et al., Gene mutations in the succinate dehydrogenase subunit SDHB cause susceptibility to familial pheochromocytoma and to familial paraganglioma. Am J Hum Genet, 2001. 69(1): p. 49-54.
6. Baysal, B.E., et al., Prevalence of SDHB, SDHC, and SDHD germline mutations in clinic patients with head and neck paragangliomas. J Med Genet, 2002. 39(3): p. 178-83.
7. Launonen, V., et al., Inherited susceptibility to uterine leiomyomas and renal cell cancer. Proc Natl Acad Sci U S A, 2001. 98(6): p. 3387-92.
8. Kiuru, M., et al., Familial cutaneous leiomyomatosis is a two-hit condition associated with renal cell cancer of characteristic histopathology. Am J Pathol, 2001. 159(3): p. 825-9.
9. Tomlinson, I.P., et al., Germline mutations in FH predispose to dominantly inherited uterine fibroids, skin leiomyomata and papillary renal cell cancer. Nat Genet, 2002. 30(4): p. 406-10.
10. Toro, J.R., et al., Mutations in the fumarate hydratase gene cause hereditary leiomyomatosis and renal cell cancer in families in North America. Am J Hum Genet, 2003. 73(1): p. 95-106.
11. Alam, N.A., et al., Genetic and functional analyses of FH mutations in multiple cutaneous and uterine leiomyomatosis, hereditary leiomyomatosis and renal cancer, and fumarate hydratase deficiency. Hum Mol Genet, 2003. 12(11): p. 1241-52.
12. Gellera, C., et al., Fumarase deficiency is an autosomal recessive encephalopathy affecting both the mitochondrial and the cytosolic enzymes. Neurology, 1990. 40(3 Pt 1): p. 495-9.
13. Bourgeron, T., et al., Mutation of the fumarase gene in two siblings with progressive encephalopathy and fumarase deficiency. J Clin Invest, 1994. 93(6): p. 2514-8.
14. Coughlin, E.M., et al., Molecular analysis and prenatal diagnosis of human fumarase deficiency. Mol Genet Metab, 1998. 63(4): p. 254-62.
15. Kiuru, M., et al., Few FH mutations in sporadic counterparts of tumor types observed in hereditary leiomyomatosis and renal cell cancer families. Cancer Res, 2002. 62(16): p. 4554-7.
16. Lorenzato, A., et al., A cancer-predisposing "hot spot" mutation of the fumarase gene creates a dominant negative protein. Int J Cancer, 2007.
17. Merino, M.J., et al., The Morphologic Spectrum of Kidney Tumors in Hereditary Leiomyomatosis and Renal Cell Carcinoma (HLRCC) Syndrome. Am J Surg Pathol, 2007. 31(10): p. 1578-1585.
18. Lehtonen, H.J., et al., Increased risk of cancer in patients with fumarate hydratase germline mutation. J Med Genet, 2006. 43(6): p. 523-6.
19. Semenza, G.L., HIF-1 and tumor progression: pathophysiology and therapeutics. Trends Mol Med, 2002. 8(4 Suppl): p. S62-7.
20. Covello, K.L. and M.C. Simon, HIFs, hypoxia, and vascular development. Curr Top Dev Biol, 2004. 62: p. 37-54.
21. Birner, P., et al., Overexpression of hypoxia-inducible factor 1alpha is a marker for an unfavorable prognosis in early-stage invasive cervical cancer. Cancer Res, 2000. 60(17): p. 4693-6.
22. Zhong, H., et al., Overexpression of hypoxia-inducible factor 1alpha in common human cancers and their metastases. Cancer Res, 1999. 59(22): p. 5830-5.
23. Talks, K.L., et al., The expression and distribution of the hypoxia-inducible factors HIF-1alpha and HIF-2alpha in normal human tissues, cancers, and tumor-associated macrophages. Am J Pathol, 2000. 157(2): p. 411-21.
24. Iwai, K., et al., Identification of the von Hippel-lindau tumor-suppressor protein as part of an active E3 ubiquitin ligase complex. Proc Natl Acad Sci U S A, 1999. 96(22): p. 12436-41.
25. Ohh, M., et al., Ubiquitination of hypoxia-inducible factor requires direct binding to the beta-domain of the von Hippel-Lindau protein. Nat Cell Biol, 2000. 2(7): p. 423-7.
26. Bruick, R.K. and S.L. McKnight, A conserved family of prolyl-4-hydroxylases that modify HIF. Science, 2001. 294(5545): p. 1337-40.
27. Epstein, A.C., et al., C. elegans EGL-9 and mammalian homologs define a family of dioxygenases that regulate HIF by prolyl hydroxylation. Cell, 2001. 107(1): p. 43-54.
28. Jaakkola, P., et al., Targeting of HIF-alpha to the von Hippel-Lindau ubiquitylation complex by O2-regulated prolyl hydroxylation. Science, 2001. 292(5516): p. 468-72.
29. Semenza, G.L., Targeting HIF-1 for cancer therapy. Nat Rev Cancer, 2003. 3(10): p. 721-32.
30. Pollard, P.J., et al., Accumulation of Krebs cycle intermediates and over-expression of HIF1alpha in tumors which result from germline FH and SDH mutations. Hum Mol Genet, 2005. 14(15): p. 2231-9.
31. Isaacs, J.S., et al., HIF overexpression correlates with biallelic loss of fumarate hydratase in renal cancer: Novel role of fumarate in regulation of HIF stability. Cancer Cell, 2005. 8(2): p. 143-53.
32. Pollard, P., et al., Evidence of increased microvessel density and activation of the hypoxia pathway in tumors from the hereditary leiomyomatosis and renal cell cancer syndrome. J Pathol, 2005. 205(1): p. 41-9.
33. Nor, J.E., et al., Thrombospondin-1 induces endothelial cell apoptosis and inhibits angiogenesis by activating the caspase death pathway. J Vasc Res, 2000. 37(3): p. 209-18.
34. Selak, M.A., et al., Succinate links TCA cycle dysfunction to oncogenesis by inhibiting HIF-alpha prolyl hydroxylase. Cancer Cell, 2005. 7(1): p. 77-85.
35. Dalgard, C.L., et al., Endogenous 2-oxoacids differentially regulate expression of oxygen sensors. Biochem J, 2004. 380(Pt 2): p. 419-24.
36. Lu, H., et al., Reversible inactivation of HIF-1 prolyl hydroxylases allows cell metabolism to control basal HIF-1. J Biol Chem, 2005. 280(51): p. 41928-39.
37. Koivunen, P., et al., Inhibition of hypoxia-inducible factor (HIF) hydroxylases by citric acid cycle intermediates: possible links between cell metabolism and stabilization of HIF. J Biol Chem, 2007. 282(7): p. 4524-32.
38. Pollard, P.J., et al., Targeted inactivation of fh1 causes proliferative renal cyst development and activation of the hypoxia pathway. Cancer Cell, 2007. 11(4): p. 311-9.
39. Harris, A.L., Hypoxia--a key regulatory factor in tumor growth. Nat Rev Cancer, 2002. 2(1): p. 38-47.
40. Elstrom, R.L., et al., Akt stimulates aerobic glycolysis in cancer cells. Cancer Res, 2004. 64(11): p. 3892-9.
41. Sowter, H.M., et al., HIF-1-dependent regulation of hypoxic induction of the cell death factors BNIP3 and NIX in human tumors. Cancer Res, 2001. 61(18): p. 6669-73.
42. Erler, J.T., et al., Hypoxia-mediated down-regulation of Bid and Bax in tumors occurs via hypoxia-inducible factor 1-dependent and -independent mechanisms and contributes to drug resistance. Mol Cell Biol, 2004. 24(7): p. 2875-89.
43. Chandel, N.S., et al., Reactive oxygen species generated at mitochondrial complex III stabilize hypoxia-inducible factor-1alpha during hypoxia: a mechanism of O2 sensing. J Biol Chem, 2000. 275(33): p. 25130-8.
44. Johnstone, R.W., A.A. Ruefli, and S.W. Lowe, Apoptosis: a link between cancer genetics and chemotherapy. Cell, 2002. 108(2): p. 153-64.
45. Wang, X., The expanding role of mitochondria in apoptosis. Genes Dev, 2001. 15(22): p. 2922-33.
46. Cory, S. and J.M. Adams, The Bcl2 family: regulators of the cellular life-or-death switch. Nat Rev Cancer, 2002. 2(9): p. 647-56.
47. Rathmell, J.C. and C.B. Thompson, The central effectors of cell death in the immune system. Annu Rev Immunol, 1999. 17: p. 781-828.
48. Savill, J. and V. Fadok, Corpse clearance defines the meaning of cell death. Nature, 2000. 407(6805): p. 784-8.
49. Downward, J., Cell biology: metabolism meets death. Nature, 2003. 424(6951): p. 896-7.
50. Ricci, J.E., et al., Disruption of mitochondrial function during apoptosis is mediated by caspase cleavage of the p75 subunit of complex I of the electron transport chain. Cell, 2004. 117(6): p. 773-86.
51. Eng, C., et al., A role for mitochondrial enzymes in inherited neoplasia and beyond. Nat Rev Cancer, 2003. 3(3): p. 193-202.
52. Gatenby, R.A. and R.J. Gillies, Why do cancers have high aerobic glycolysis? Nat Rev Cancer, 2004. 4(11): p. 891-9.
53. Kim, J.W. and C.V. Dang, Multifaceted roles of glycolytic enzymes. Trends Biochem Sci, 2005. 30(3): p. 142-50.
54. Majewski, N., et al., Hexokinase-mitochondria interaction mediated by Akt is required to inhibit apoptosis in the presence or absence of Bax and Bak. Mol Cell, 2004. 16(5): p. 819-30.
55. Pastorino, J.G., N. Shulga, and J.B. Hoek, Mitochondrial binding of hexokinase II inhibits Bax-induced cytochrome c release and apoptosis. J Biol Chem, 2002. 277(9): p. 7610-8.
56. Danial, N.N., et al., BAD and glucokinase reside in a mitochondrial complex that integrates glycolysis and apoptosis. Nature, 2003. 424(6951): p. 952-6.
57. Azoulay-Zohar, H., et al., In self-defence: hexokinase promotes voltage-dependent anion channel closure and prevents mitochondria-mediated apoptotic cell death. Biochem J, 2004. 377(Pt 2): p. 347-55.
58. Majewski, N., et al., Akt inhibits apoptosis downstream of BID cleavage via a glucose-dependent mechanism involving mitochondrial hexokinases. Mol Cell Biol, 2004. 24(2): p. 730-40.
59. Ravagnan, L., T. Roumier, and G. Kroemer, Mitochondria, the killer organelles and their weapons. J Cell Physiol, 2002. 192(2): p. 131-7.
60. Miramar, M.D., et al., NADH oxidase activity of mitochondrial apoptosis-inducing factor. J Biol Chem, 2001. 276(19): p. 16391-8.
61. Mates, J.M., et al., Glutamine and its relationship with intracellular redox status, oxidative stress and cell proliferation/death. Int J Biochem Cell Biol, 2002. 34(5): p. 439-58.
62. Chang, W.K., et al., Glutamine protects activated human T cells from apoptosis by up-regulating glutathione and Bcl-2 levels. Clin Immunol, 2002. 104(2): p. 151-60.
63. Lee, S., et al., Neuronal apoptosis linked to EglN3 prolyl hydroxylase and familial pheochromocytoma genes: Developmental culling and cancer. Cancer Cell, 2005. 8(2): p. 155-167.
64. Wortham, N.C., et al., Aberrant expression of apoptosis proteins and ultrastructural aberrations in uterine leiomyomas from patients with hereditary leiomyomatosis and renal cell carcinoma. Fertil Steril, 2006. 86(4): p. 961-71.
65. Raha, S. and B.H. Robinson, Mitochondria, oxygen free radicals, disease and ageing. Trends Biochem Sci, 2000. 25(10): p. 502-8.
66. Gerald, D., et al., JunD reduces tumor angiogenesis by protecting cells from oxidative stress. Cell, 2004. 118(6): p. 781-94.
67. Selak, M.A., R.V. Duran, and E. Gottlieb, Redox stress is not essential for the pseudo-hypoxic phenotype of succinate dehydrogenase deficient cells. Biochim Biophys Acta, 2006. 1757(5-6): p. 567-72.
68. Lehtonen, H.J., et al., Increased HIF1 alpha in SDH and FH deficient tumors does not cause microsatellite instability. Int J Cancer, 2007. 121(6): p. 1386-9.

2010-01-25T09:37:08 - Annalisa Ghiglia

Relazione delle Dott.sse Annalisa Ghiglia e Nadia Dani

AddThis Social Bookmark Button