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Blood test uses DNA strands of dying cells

Posted in Cancer - General, CANCER BIOLOGY & Innovations in Cancer Therapy, Chemical Biology and its relations to Metabolic Disease, Clinical Diagnostics, Curation, Diagnostics and Lab Tests, Genomic Expression, Neurodegenerative Diseases, Neurological Diseases, Population Health Management, Genetics & Pharmaceutical, tagged Amino acids, Anaplerosis, Biosynthesis, Cancer - General, circulating DNA, Diagnosis, lipogenesis, Methylation, mitochondria, nucleotides on April 9, 2016| Leave a Comment »

Blood test uses DNA strands of dying cells

Curators: Larry H. Bernstein, MD, FCAP and Aviva Lev-Ari, PhD, RN

LPBI

Hadassah-Developed Blood Test Detects Multiple Sclerosis, Cancer & Brain Damage

http://www.hadassah.org/news-stories/blood-test-detects-neurodegenerative-disease.html

A new blood test that uses the DNA strands of dying cells to detect diabetes, cancer, traumatic brain injury, and neurodegenerative disease has been developed by researchers at Hadassah Medical Organization (HMO) and The Hebrew University.

In a study involving 320 patients, the researchers were able to infer cell death in specific tissues by looking at the unique chemical modifications (called methylation patterns) of circulating DNA that these dying cells release. Previously, it had not been possible to measure cell death in specific human tissues non-invasively.

The findings are reported in the March 14, 2016 online edition of Proceedings of National Academy of Sciences USA, in an article entitled “Identification of tissue specific cell death using methylation patterns of circulating DNA.” Prof. Benjamin Glaser, head of Endocrinology at Hadassah, and Dr. Ruth Shemer and Prof. Yuval Dor from The Hebrew University of Jerusalem led an international team in performing the groundbreaking research.

Cell death is a central feature in health and disease. It can signify the early stages of pathology (e.g. a developing tumor or the beginning of an autoimmune or neurodegenerative disease); it can illuminate whether a disease has progressed and whether a particular treatment, such as chemotherapy, is working; and it can alert physicians to unintended toxic effects of treatment or the early rejection of a transplant.

As the researchers relate: “The approach can be adapted to identify cfDNA (cell-free circulating DNA) derived from any cell type in the body, offering a minimally invasive window for diagnosing and monitoring a broad spectrum of human pathologies as well as providing a better understanding of normal tissue dynamics.”

“In the long run,” notes Prof. Glaser, “we envision a new type of blood test aimed at the sensitive detection of tissue damage, even without a-priori suspicion of disease in a specific organ. We believe that such a tool will have broad utility in diagnostic medicine and in the study of human biology.”

The research was performed by Hebrew University students Roni Lehmann-Werman, Daniel Neiman, Hai Zemmour, Joshua Moss and Judith Magenheim, aided by clinicians and scientists from Hadassah Medical Center, Sheba Medical Center, and from institutions in Germany, Sweden, the USA and Canada, who provided precious blood samples from patients.

Scientists have known for decades that dying cells release fragmented DNA into the blood; however, since the DNA sequence of all cells in the body is identical, it had not been possible to determine the tissue of origin of the circulating DNA. Knowing that the DNA of each cell type carries a unique methylation and that methylation patterns of DNA account for the identity of cells, the researchers were able to use patterns of methylated DNA sequences as biomarkers to detect the origin of the DNA and to identify a specific pathology. For example, they were able to detect evidence of pancreatic beta-cell death in the blood of patients with new-onset type 1 diabetes, oligodendrocyte cell death in patients with relapsing multiple sclerosis, brain cell death in patients after traumatic or ischemic brain damage, and exocrine pancreatic tissue cell death in patients with pancreatic cancer or pancreatitis.

Support for the research came from the Juvenile Diabetes Research Foundation, the Human Islet Research Network of the National Institutes of Health, the Sir Zalman Cowen Universities Fund, the DFG (a Trilateral German-Israel-Palestine program), and the Soyka pancreatic cancer fund.

Identification of tissue-specific cell death using methylation patterns of circulating DNA.

Lehmann-Werman R¹, Neiman D¹, Zemmour H¹, Moss J¹, ….., Razin A¹, Cedar H¹, Shapiro AM¹⁵, Glaser B¹⁶, Shemer R¹⁷, Dor Y¹⁷.

Proc Natl Acad Sci U S A. 2016 Mar 29;113(13):E1826-34. http://dx.doi.org:/10.1073/pnas.1519286113

Minimally invasive detection of cell death could prove an invaluable resource in many physiologic and pathologic situations. Cell-free circulating DNA (cfDNA) released from dying cells is emerging as a diagnostic tool for monitoring cancer dynamics and graft failure. However, existing methods rely on differences in DNA sequences in source tissues, so that cell death cannot be identified in tissues with a normal genome. We developed a method of detecting tissue-specific cell death in humans based on tissue-specific methylation patterns in cfDNA. We interrogated tissue-specific methylome databases to identify cell type-specific DNA methylation signatures and developed a method to detect these signatures in mixed DNA samples. We isolated cfDNA from plasma or serum of donors, treated the cfDNA with bisulfite, PCR-amplified the cfDNA, and sequenced it to quantify cfDNA carrying the methylation markers of the cell type of interest. Pancreatic β-cell DNA was identified in the circulation of patients with recently diagnosed type-1 diabetes and islet-graft recipients; oligodendrocyte DNA was identified in patients with relapsing multiple sclerosis; neuronal/glial DNA was identified in patients after traumatic brain injury or cardiac arrest; and exocrine pancreas DNA was identified in patients with pancreatic cancer or pancreatitis. This proof-of-concept study demonstrates that the tissue origins of cfDNA and thus the rate of death of specific cell types can be determined in humans. The approach can be adapted to identify cfDNA derived from any cell type in the body, offering a minimally invasive window for diagnosing and monitoring a broad spectrum of human pathologies as well as providing a better understanding of normal tissue dynamics.

While impressively organ specific, they did not specifically prove that the DNA was from an actual dying cell. For example, you would need to see if Troponin levels were elevated when assuming the DNA is from injured myocardium. Also, for brain, though impractical , you’d want to see a brain biopsy or imaging for the brain related cases. The experiment of spiking with DNA was clever though. Also, what is the turnaround time for this test in practical use?

Larry HB

Very good comment. I was reluctant to put this up, but it was of interest and published in PNAS. Perhaps I can find more information. Troponin levels would be good for 48 hours, longer than CK and comparable to LD. What about Nat peptides?

Glutamine and cancer: cell biology, physiology, and clinical opportunities

Christopher T. Hensley,¹ Ajla T. Wasti,^1,2

J Clin Invest 2013 https://www.jci.org/articles/view/69600

Glutamine is an abundant and versatile nutrient that participates in energy formation, redox homeostasis, macromolecular synthesis, and signaling in cancer cells. These characteristics make glutamine metabolism an appealing target for new clinical strategies to detect, monitor, and treat cancer. Here we review the metabolic functions of glutamine as a super nutrient and the surprising roles of glutamine in supporting the biological hallmarks of malignancy. We also review recent efforts in imaging and therapeutics to exploit tumor cell glutamine dependence, discuss some of the challenges in this arena, and suggest a disease-focused paradigm to deploy these emerging approaches.

It has been nearly a century since the discovery that tumors display metabolic activities that distinguish them from differentiated, non-proliferating tissues and presumably contribute to their supraphysiological survival and growth (1). Interest in cancer metabolism was boosted by discoveries that oncogenes and tumor suppressors could regulate nutrient metabolism, and that mutations in some metabolic enzymes participate in the development of malignancy (2, 3). The persistent appeal of cancer metabolism as a line of investigation lies both in its ability to uncover fundamental aspects of malignancy and in the translational potential of exploiting cancer metabolism to improve the way we diagnose, monitor, and treat cancer. Furthermore, an improved understanding of how altered metabolism contributes to cancer has a high potential for synergy with translational efforts. For example, the demonstration that asparagine is a conditionally essential nutrient in rapidly growing cancer cells paved the way for L-asparaginase therapy in leukemia. Additionally, the avidity of some tumors for glucose uptake led to the development of ¹⁸fluoro-2-deoxyglucose imaging by PET; this in turn stimulated hundreds of studies on the biological underpinnings of tumor glucose metabolism.

There continue to be large gaps in understanding which metabolic pathways are altered in cancer, whether these alterations benefit the tumor in a substantive way, and how this information could be used in clinical oncology. In this Review, we consider glutamine, a highly versatile nutrient whose metabolism has implications for tumor cell biology, metabolic imaging, and perhaps novel therapeutics.

Glutamine in intermediary metabolism

Glutamine metabolism has been reviewed extensively and is briefly outlined here (4, 5). The importance of glutamine as a nutrient in cancer derives from its abilities to donate its nitrogen and carbon into an array of growth-promoting pathways (Figure 1). At concentrations of 0.6–0.9 mmol/l, glutamine is the most abundant amino acid in plasma (6). Although most tissues can synthesize glutamine, during periods of rapid growth or other stresses, demand outpaces supply, and glutamine becomes conditionally essential (7). This requirement for glutamine is particularly true in cancer cells, many of which display oncogene-dependent addictions to glutamine in culture (8). Glutamine catabolism begins with its conversion to glutamate in reactions that either donate the amide nitrogen to biosynthetic pathways or release it as ammonia. The latter reactions are catalyzed by the glutaminases (GLSs), of which several isozymes are encoded by human genes GLS and GLS2 (9). Classical studies revealed that GLS isozymes, particularly those encoded by GLS, are expressed in experimental tumors in rats and mice, where their enzyme activity correlates with growth rate and malignancy. Silencing GLS expression or inhibiting GLS activity is sufficient to delay tumor growth in a number of models (10–13). The role of GLS2 in cancer appears to be context specific and regulated by factors that are still incompletely characterized. In some tissues, GLS2 is a p53 target gene and seems to function in tumor suppression (14). On the other hand, GLS2 expression is enhanced in some neuroblastomas, where it contributes to cell survival (15). These observations, coupled with the demonstration that c-Myc stimulates GLS expression (12, 16), position at least some of the GLS isozymes as pro-oncogenic.

Figure 1Glutamine metabolism as a target for diagnostic imaging and therapy in cancer. Glutamine is imported via SLC1A5 and other transporters, then enters a complex metabolic network by which its carbon and nitrogen are supplied to pathways that promote cell survival and growth. Enzymes discussed in the text are shown in green, and inhibitors that target various aspects of glutamine metabolism are shown in red. Green arrows denote reductive carboxylation. ¹⁸F-labeled analogs of glutamine are also under development as PET probes for localization of tumor tissue. AcCoA, acetyl-CoA; DON, 6-diazo-5-oxo-L-norleucine; GSH, glutathione; NEAA, nonessential amino acids; ME, malic enzyme; OAA, oxaloacetate; TA, transaminase; 968, compound 968; α-KG, α-ketoglutarate.

Glutamate, the product of the GLS reaction, is a precursor of glutathione, the major cellular antioxidant. It is also the source of amino groups for nonessential amino acids like alanine, aspartate, serine, and glycine, all of which are required for macromolecular synthesis. In glutamine-consuming cells, glutamate is also the major source of α-ketoglutarate, a TCA cycle intermediate and substrate for dioxygenases that modify proteins and DNA. These dioxygenases include prolyl hydroxylases, histone demethylases, and 5-methylcytosine hydroxylases. Their requirement for α-ketoglutarate, although likely accounting for only a small fraction of total α-ketoglutarate utilization, makes this metabolite an essential component of cell signaling and epigenetic networks.

Conversion of glutamate to α-ketoglutarate occurs either through oxidative deamination by glutamate dehydrogenase (GDH) in the mitochondrion or by transamination to produce nonessential amino acids in either the cytosol or the mitochondrion. During avid glucose metabolism, the transamination pathway predominates (17). When glucose is scarce, GDH becomes a major pathway to supply glutamine carbon to the TCA cycle, and is required for cell survival (17, 18). Metabolism of glutamine-derived α-ketoglutarate in the TCA cycle serves several purposes: it generates reducing equivalents for the electron transport chain (ETC) and oxidative phosphorylation, becoming a major source of energy (19); and it is an important anaplerotic nutrient, feeding net production of oxaloacetate to offset export of intermediates from the cycle to supply anabolism (20). Glutamine oxidation also supports redox homeostasis by supplying carbon to malic enzyme, some isoforms of which produce NADPH (Figure 1). In KRAS-driven pancreatic adenocarcinoma cells, a pathway involving glutamine-dependent NADPH production is essential for redox balance and growth (21). In these cells, glutamine is used to produce aspartate in the mitochondria. This aspartate is then trafficked to the cytosol, where it is deaminated to produce oxaloacetate and then malate, the substrate for malic enzyme.

Recent work has uncovered an unexpected role for glutamine in cells with reduced mitochondrial function. Despite glutamine’s conventional role as a respiratory substrate, several studies demonstrated a persistence of glutamine dependence in cells with permanent mitochondrial dysfunction from mutations in the ETC or TCA cycle, or transient impairment secondary to hypoxia (22–25). Under these conditions, glutamine-derived α-ketoglutarate is reductively carboxylated by NADPH-dependent isoforms of isocitrate dehydrogenase to produce isocitrate, citrate, and other TCA cycle intermediates (Figure 1). These conditions broaden glutamine’s utility as a carbon source because it becomes not only a major source of oxaloacetate, but also generates acetyl-CoA in what amounts to a striking rewiring of TCA cycle metabolism.

Glutamine promotes hallmarks of malignancy

Deregulated energetics. One hallmark of cancer cells is aberrant bioenergetics (26). Glutamine’s involvement in the pathways outlined above contributes to a phenotype conducive to energy formation, survival, and growth. In addition to its role in mitochondrial metabolism, glutamine also suppresses expression of thioredoxin-interacting protein, a negative regulator of glucose uptake (27). Thus, glutamine contributes to both of the energy-forming pathways in cancer cells: oxidative phosphorylation and glycolysis. Glutamine also modulates hallmarks not traditionally thought to be metabolic, as outlined below. These interactions highlight the complex interplay between glutamine metabolism and many aspects of cell biology.

Sustaining proliferative signaling. Pathological cancer cell growth relies on maintenance of proliferative signaling pathways with increased autonomy relative to non-malignant cells. Several lines of evidence argue that glutamine reinforces activity of these pathways. In some cancer cells, excess glutamine is exported in exchange for leucine and other essential amino acids. This exchange facilitates activation of the serine/threonine kinase mTOR, a major positive regulator of cell growth (28). In addition, glutamine-derived nitrogen is a component of amino sugars, known as hexosamines, that are used to glycosylate growth factor receptors and promote their localization to the cell surface. Disruption of hexosamine synthesis reduces the ability to initiate signaling pathways downstream of growth factors (29).

Enabling replicative immortality. Some aspects of glutamine metabolism oppose senescence and promote replicative immortality in cultured cells. In IMR90 lung fibroblasts, silencing either of two NADPH-generating isoforms of malic enzyme (ME1, ME2) rapidly induced senescence, while malic enzyme overexpression suppressed senescence (30). Both malic enzyme isoforms are repressed at the transcriptional level by p53 and contribute to enhanced levels of glutamine consumption and NADPH production in p53-deficient cells. The ability of p53-replete cells to resist senescence required the expression of ME1 and ME2, and silencing either enzyme reduced the growth of TP53^+/+ and, to a lesser degree, TP53^–/– tumors (30). These observations position malic enzymes as potential therapeutic targets.

Resisting cell death. Although many cancer cells require glutamine for survival, cells with enhanced expression of Myc oncoproteins are particularly sensitive to glutamine deprivation (8, 12, 16). In these cells, glutamine deprivation induces depletion of TCA cycle intermediates, depression of ATP levels, delayed growth, diminished glutathione pools, and apoptosis. Myc drives glutamine uptake and catabolism by activating the expression of genes involved in glutamine metabolism, including GLSand SLC1A5, which encodes the Na⁺-dependent amino acid transporter ASCT2 (12, 16). SilencingGLS mimicked some of the effects of glutamine deprivation, including growth suppression in Myc-expressing cells and tumors (10, 12). MYCN amplification occurs in 20%–25% of neuroblastomas and is correlated with poor outcome (31). In cells with high N-Myc levels, glutamine deprivation triggered an ATF4-dependent induction of apoptosis that could be prevented by restoring downstream metabolites oxaloacetate and α-ketoglutarate (15). In this model, pharmacological activation of ATF4, inhibition of glutamine metabolic enzymes, or combinations of these treatments mimicked the effects of glutamine deprivation in cells and suppressed growth of MYCN-amplified subcutaneous and transgenic tumors in mice.

The PKC isoform PKC-ζ also regulates glutamine metabolism. Loss of PKC-ζ enhances glutamine utilization and enables cells to survive glucose deprivation (32). This effect requires flux of carbon and nitrogen from glutamine into serine. PKC-ζ reduces the expression of phosphoglycerate dehydrogenase, an enzyme required for glutamine-dependent serine biosynthesis, and also phosphorylates and inactivates this enzyme. Thus, PKC-ζ loss, which promotes intestinal tumorigenesis in mice, enables cells to alter glutamine metabolism in response to nutrient stress.

Invasion and metastasis. Loss of the epithelial cell-cell adhesion molecule E-cadherin is a component of the epithelial-mesenchymal transition, and is sufficient to induce migration, invasion, and tumor progression (33, 34). Addiction to glutamine may oppose this process because glutamine favors stabilization of tight junctions in some cells (35). Furthermore, the selection of breast cancer cells with the ability to grow without glutamine yielded highly adaptable subpopulations with enhanced mesenchymal marker expression and improved capacity for anchorage-independent growth, therapeutic resistance, and metastasis in vivo (36). It is unknown whether this result reflects a primary role for glutamine in suppressing these markers of aggressiveness in breast cancer, or whether prolonged glutamine deprivation selects for cells with enhanced fitness across a number of phenotypes.

Organ-specific glutamine metabolism in health and disease

As a major player in carbon and nitrogen transport, glutamine metabolism displays complex inter-organ dynamics, with some organs functioning as net producers and others as consumers (Figure 2). Organ-specific glutamine metabolism has frequently been studied in humans and animal models by measuring the arteriovenous difference in plasma glutamine abundance. In healthy subjects, the plasma glutamine pool is largely the result of release from skeletal muscle (37–39). In rats, the lungs are comparable to muscle in terms of glutamine production (40, 41), and human lungs also have the capacity for marked glutamine release, although such release is most prominent in times of stress (42, 43). Stress-induced release from the lung is regulated by an induction of glutamine synthase expression as a consequence of glucocorticoid signaling and other mechanisms (44, 45). Although this results in a small arteriovenous difference, the overall release of glutamine is significant because of the large pulmonary perfusion. In rats and humans, adipose tissue is a minor but potentially important source of glutamine (46, 47). The liver has the capacity to synthesize or catabolize glutamine, with these activities subject both to regional heterogeneity among hepatocytes and regulatory effects of systemic acidosis and hyperammonemia. However, the liver does not appear to be a major contributor to the plasma glutamine pool in healthy rats and humans (39, 48, 49).

Figure 2Model for inter-organ glutamine metabolism in health and cancer. Organs that release glutamine into the bloodstream are shown in green, and those that consume glutamine are in red; the shade denotes magnitude of consumption/release. For some organs (liver, kidneys), evidence from model systems and/or human studies suggests that there is a change in net glutamine flux during tumorigenesis.

Glutamine consumption occurs largely in the gut and kidney. The organs of the gastrointestinal tract drained by the portal vein, particularly the small intestine, are major consumers of plasma glutamine in both rats and humans (37, 38, 49, 50). Enterocytes oxidize more than half of glutamine carbon to CO₂, accounting for a third of the respiration of these cells in fasting animals (51). The kidney consumes net quantities of glutamine to maintain acid-base balance (37, 38, 52, 53). During acidosis, the kidneys substantially increase their uptake of glutamine, cleaving it by GLS to produce ammonia, which is excreted along with organic acids to maintain physiologic pH (52, 54). Glutamine is also a major metabolic substrate in lymphocytes and macrophages, at least during mitogenic stimulation of primary cells in culture (55–57).

Importantly, cancer seems to cause major changes in inter-organ glutamine trafficking (Figure 2). Currently, much work in this area is derived from studies in methylcholanthrene-induced fibrosarcoma in the rat, a model of an aggressively growing, glutamine-consuming tumor. In this model, fibrosarcoma induces skeletal muscle expression of glutamine synthetase and greatly increases the release of glutamine into the circulation. As the tumor increases in size, intramuscular glutamine pools are depleted in association with loss of lean muscle mass, mimicking the cachectic phenotype of humans in advanced stages of cancer (52). Simultaneously, both the liver and the kidneys become net glutamine exporters, although the hepatic effect may be diminished as the tumor size becomes very large (48, 49, 52). Glutamine utilization by organs supplied by the portal vein is diminished in cancer (48). In addition to its function as a nutrient for the tumor itself, and possibly for cancer-associated immune cells, glutamine provides additional, indirect metabolic benefits to both the tumor and the host. For example, glutamine was used as a gluconeogenic substrate in cachectic mice with large orthotopic gliomas, providing a significant source of carbon in the plasma glucose pool (58). This glucose was taken up and metabolized by the tumor to produce lactate and to supply the TCA cycle.

It will be valuable to extend work in human inter-organ glutamine trafficking, both in healthy subjects and in cancer patients. Such studies will likely produce a better understanding of the pathophysiology of cancer cachexia, a major source of morbidity and mortality. Research in this area should also aid in the anticipation of organ-specific toxicities of drugs designed to interfere with glutamine metabolism. Alterations of glutamine handling in cancer may induce a different spectrum of toxicities compared with healthy subjects.

Tumors differ according to their need for glutamine

One important consideration is that not all cancer cells need an exogenous supply of glutamine. A panel of lung cancer cell lines displayed significant variability in their response to glutamine deprivation, with some cells possessing almost complete independence (59). Breast cancer cells also demonstrate systematic differences in glutamine dependence, with basal-type cells tending to be glutamine dependent and luminal-type cells tending to be glutamine independent (60). Resistance to glutamine deprivation is associated with the ability to synthesize glutamine de novo and/or to engage alternative pathways of anaplerosis (10, 60).

Tumors also display variable levels of glutamine metabolism in vivo. A study of orthotopic gliomas revealed that genetically diverse, human-derived tumors took up glutamine in the mouse brain but did not catabolize it (58). Rather, the tumors synthesized glutamine de novo and used pyruvate carboxylation for anaplerosis. Cells derived from these tumors did not require glutamine to survive or proliferate when cultured ex vivo. Glutamine synthesis from glucose was also a prominent feature of primary gliomas in human subjects infused with ¹³C-glucose at the time of surgical resection (61). Furthermore, an analysis of glutamine metabolism in lung and liver tumors revealed that both the tissue of origin and the oncogene influence whether the tumor produces or consumes glutamine (62). MET-induced hepatic tumors produced glutamine, whereas Myc-induced liver tumors catabolized it. In the lung, however, Myc expression was associated with glutamine accumulation.

This variability makes it imperative to develop ways to predict which tumors have the highest likelihood of responding to inhibitors of glutamine metabolism. Methods to image or otherwise quantify glutamine metabolism in vivo would be useful in this regard (63). Infusions of pre-surgical subjects with isotopically labeled glutamine, followed by extraction of metabolites from the tumor and analysis of ¹³C enrichment, can be used to detect both glutamine uptake and catabolism (58, 62). However, this approach requires a specimen of the tumor to be obtained. Approaches for glutamine-based imaging, which avoid this problem, include a number of glutamine analogs compatible with PET. Although glutamine could in principle be imaged using the radioisotopes ¹¹C, ¹³N, or ¹⁸F, the relatively long half-life of the latter increases its appeal. In mice, ¹⁸F-(2S, 4R)4-fluoroglutamine is avidly taken up by tumors derived from highly glutaminolytic cells, and by glutamine-consuming organs including the intestine, kidney, liver, and pancreas (64). Labeled analogs of glutamate are also taken up by some tumors (65, 66). One of these, (4S)-4-(3-[¹⁸F] fluoropropyl)-L-glutamate (¹⁸F-FSPG, also called BAY 94-9392), was evaluated in small clinical trials involving patients with several types of cancer (65, 67). This analog enters the cell through the cystine/glutamate exchange transporter (xC^–transport system), which is linked to glutathione biosynthesis (68). The analog was well tolerated, with high tumor detection rates and good tumor-to-background ratios in hepatocellular carcinoma and lung cancer.

PET approaches detect analog uptake and retention but cannot provide information about downstream metabolism. Analysis of hyperpolarized nuclei can provide a real-time view of enzyme-catalyzed reactions. This technique involves redistribution of the populations of energy levels of a nucleus (e.g., ¹³C, ¹⁵N), resulting in a gain in magnetic resonance signal that can temporarily exceed 10,000-fold (69). This gain in signal enables rapid detection of both the labeled molecule and its downstream metabolites. Glutamine has been hyperpolarized on ¹⁵N and ¹³C (70, 71). In the latter case, the conversion of hyperpolarized glutamine to glutamate could be detected in intact hepatoma cells (70). If these analogs are translated to clinical studies, they might provide a dynamic view of the proximal reactions of glutaminolysis in vivo.

Pharmacological strategies to inhibit glutamine metabolism in cancer

Efforts to inhibit glutamine metabolism using amino acid analogs have an extensive history, including evaluation in clinical trials. Acivicin, 6-diazo-5-oxo-L-norleucine, and azaserine, three of the most widely studied analogs (Figure 1), all demonstrated variable degrees of gastrointestinal toxicity, myelosuppression, and neurotoxicity (72). Because these agents non-selectively target glutamine-consuming processes, recent interest has focused on developing methods directed at specific nodes of glutamine metabolism. First, ASCT2, the Na⁺-dependent neutral amino acid transporter encoded by SLC1A5, is broadly expressed in lung cancer cell lines and accounts for a majority of glutamine transport in those cells (Figure 1). It has been shown that γ-L-glutamyl-p-nitroanilide (GPNA) inhibits this transporter and limits lung cancer cell growth (73). Additional interest in GPNA lies in its ability to enhance the uptake of drugs imported via the monocarboxylate transporter MCT1. Suppressing glutamine uptake with GPNA enhances MCT1 stability and stimulates uptake of the glycolytic inhibitor 3-bromopyruvate (3-BrPyr) (74, 75). Because enforced MCT1 overexpression is sufficient to sensitize tumor xenografts to 3-BrPyr (76), GPNA may have a place in 3-BrPyr–based therapeutic regimens.

Two inhibitors of GLS isoforms have been characterized in recent years (Figure 1). Compound 968, an inhibitor of the GLS-encoded splice isoform GAC, inhibits the transformation of fibroblasts by oncogenic RhoGTPases and delays the growth of GLS-expressing lymphoma xenografts (13). Bis-2-(5-phenylacetamido-1,2,4-thiadiazol-2-yl)ethyl sulfide (BPTES) also potently inhibits GLS isoforms encoded by GLS (77). BPTES impairs ATP levels and growth rates of P493 lymphoma cells under both normoxic and hypoxic conditions and suppresses the growth of P493-derived xenografts (78).

Evidence also supports a role for targeting the flux from glutamate to α-ketoglutarate, although no potent, specific inhibitors yet exist to inhibit these enzymes in intact cells. Aminooxyacetate (AOA) inhibits aminotransferases non-specifically, but milliomolar doses are typically used to achieve this effect in cultured cells (Figure 1). Nevertheless, AOA has demonstrated efficacy in both breast adenocarcinoma xenografts and autochthonous neuroblastomas in mice (15, 79). Epigallocatechin gallate (EGCG), a green tea polyphenol, has numerous pharmacological effects, one of which is to inhibit GDH (80). The effects of EGCG on GDH have been used to kill glutamine-addicted cancer cells during glucose deprivation or glycolytic inhibition (17, 18) and to suppress growth of neuroblastoma xenografts (15).

A paradigm to exploit glutamine metabolism in cancer

Recent advances in glutamine-based imaging, coupled with the successful application of glutamine metabolic inhibitors in mouse models of cancer, make it possible to conceive of treatment plans that feature consideration of tumor glutamine utilization. A key challenge will be predicting which tumors are most likely to respond to inhibitors of glutamine metabolism. Neuroblastoma is used here as an example of a tumor in which evidence supports the utility of strategies that would involve both glutamine-based imaging and therapy (Figure 3). Neuroblastoma is the second most common extracranial solid malignancy of childhood. High-risk neuroblastoma is defined by age, stage, and biological features of the tumor, including MYCN amplification, which occurs in some 20%–25% of cases (31). Because MYCN-amplified tumor cells require glutamine catabolism for survival and growth (15), glutamine-based PET at the time of standard diagnostic imaging could help predict which tumors would be likely to respond to inhibitors of glutamine metabolism. Infusion of ¹³C-glutamine coordinated with the diagnostic biopsy could then enable inspection of ¹³C enrichment in glutamine-derived metabolites from the tumor, confirming the activity of glutamine catabolic pathways. Following on evidence from mouse models of neuroblastoma, treatment could then include agents directed against glutamine catabolism (15). Of note, some tumors were sensitive to the ATF4 agonist fenretinide (FRT), alone or in combination with EGCG. Importantly, FRT has already been the focus of a Phase I clinical trial in children with solid tumors, including neuroblastoma, and was fairly well tolerated (81).

Figure 3A strategy to integrate glutamine metabolism into the diagnosis, classification, treatment, and monitoring of neuroblastoma. Neuroblastoma commonly presents in children as an abdominal mass. A standard evaluation of a child with suspected neuroblastoma includes measurement of urine catecholamines, a bone scan, and full-body imaging with meta-iodobenzylguanidine (MIBG), all of which contribute to diagnosis and disease staging. In animal models, a subset of these tumors requires glutamine metabolism. This finding implies that approaches to image, quantify, or block glutamine metabolism (highlighted in red) in human neuroblastoma could be incorporated into the diagnosis and management of this disease. In particular, glutamine metabolic studies may help predict which tumors would respond to therapies targeting glutamine metabolism. HVA, homovanillic acid; VMA, vanillylmandelic acid.

Conclusions

Glutamine is a versatile nutrient required for the survival and growth of a potentially large subset of tumors. Work over the next several years should produce a more accurate picture of the molecular determinants of glutamine addiction and the identification of death pathways that execute cells when glutamine catabolism is impaired. Advancement of glutamine-based imaging into clinical practice should soon make it possible to differentiate tumors that take up glutamine from those that do not. Finally, the development of safe, high-potency inhibitors of key metabolic nodes should facilitate therapeutic regimens featuring inhibition of glutamine metabolism.

Therapeutic strategies impacting cancer cell glutamine metabolism

Michael J Lukey, Kristin F Wilson, and Richard A Cerione^*

Future Med Chem. 2013 Sep; 5(14): 1685–1700. doi: 10.4155/fmc.13.130

The metabolic adaptations that support oncogenic growth can also render cancer cells dependent on certain nutrients. Along with the Warburg effect, increased utilization of glutamine is one of the metabolic hallmarks of the transformed state. Glutamine catabolism is positively regulated by multiple oncogenic signals, including those transmitted by the Rho family of GTPases and by c-Myc. The recent identification of mechanistically distinct inhibitors of glutaminase, which can selectively block cellular transformation, has revived interest in the possibility of targeting glutamine metabolism in cancer therapy. Here, we outline the regulation and roles of glutamine metabolism within cancer cells and discuss possible strategies for, and the consequences of, impacting these processes therapeutically.

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Cancer cell metabolism & glutamine addiction

Interest in the metabolic changes characteristic of malignant transformation has undergone a renaissance of sorts in the cancer biology and pharmaceutical communities. However, the recognition that an important connection exists between cellular metabolism and cancer began nearly a century ago with the work of Otto Warburg [1–3]. Warburg found that rapidly proliferating tumor cells exhibit elevated glucose uptake and glycolytic flux, and furthermore that much of the pyruvate generated by glycolysis is reduced to lactate rather than undergoing mitochondrial oxidation via the tricarboxylic acid (TCA) cycle (Figure 1). This phenomenon persists even under aerobic conditions (‘aerobic glycolysis’), and is known as the Warburg effect [4]. Warburg proposed that aerobic glycolysis was caused by defective mitochondria in cancer cells, but it is now known that mitochondrial dysfunction is relatively rare and that most tumors have an unimpaired capacity for oxidative phosphorylation [5]. In fact, the most important selective advantages provided by the Warburg effect are still debated. Although aerobic glycolysis is an inefficient way to produce ATP (2 ATP/glucose vs ~36 ATP/glucose by complete oxidation), a high glycolytic flux can generate ATP rapidly and furthermore can provide a biosynthetic advantage by supplying precursors and reducing equivalents for the synthesis of macromolecules [4]. The mechanisms underlying the Warburg effect are also not yet fully resolved, although it is increasingly clear that a number of oncogenes and tumor suppressors contribute to the phenomenon. The PI3K/Akt/mTORC1 signaling axis, for example, is a key regulator of aerobic glycolysis and biosynthesis, driving the surface expression of nutrient transporters and the upregulation of glycolytic enzymes [6]. The HIF transcription factor also upregulates expression of glucose transporters and glycolytic enzymes in response to hypoxia and growth factors (or loss of the von Hippel–Landau [VHL] tumor suppressor), and the oncogenic transcription factor c-Myc similarly induces expression of proteins important for glycolysis [6].

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http://www.ncbi.nlm.nih.gov/pmc/articles/PMC4154374/bin/nihms610340f1.jpg

Cell proliferation requires metabolic reprogramming

A second major change in the metabolic program of many cancer cells, and the primary focus of this review, is the alteration of glutamine metabolism. Glutamine is the major carrier of nitrogen between organs, and the most abundant amino acid in plasma [7]. It is also a key nutrient for numerous intracellular processes including oxidative metabolism and ATP generation, biosynthesis of proteins, lipids and nucleic acids, and also redox homeostasis and the regulation of signal transduction pathways [8–10]. Although most mammalian cells are capable of synthesizing glutamine, the demand for this amino acid can become so great during rapid proliferation that an additional extracellular supply is required; hence glutamine is considered conditionally essential [11]. Indeed, many cancer cells are ‘glutamine addicted’, and cannot survive in the absence of an exogenous glutamine supply [12,13].

An important step in the elevation of glutamine catabolism is the activation of the mitochondrial enzyme glutaminase, which catalyzes the hydrolysis of glutamine to generate glutamate and ammonium. The subsequent deamination of glutamate releases a second ammonium to yield the TCA cycle intermediate α-ketoglutarate (α-KG), a reaction catalyzed by glutamate dehydrogenase (GLUD1). This series of reactions is particularly important in rapidly proliferating cells, in which a considerable proportion of the TCA cycle metabolite citrate is exported from mitochondria in order to generate cytosolic acetyl-CoA for lipid biosynthesis [14]. Replenishment of TCA cycle intermediates (anaplerosis) is therefore required, and glutamine often serves as the key anaplerotic substrate through its conversion via glutamate to α-KG (Figure 1).

Mammals express two genes for glutaminase enzymes [15–17]. The GLS gene encodes a protein initially characterized in kidney and thus called kidney-type glutaminase (KGA), although this enzyme and its shorter splice variant glutaminase C (GAC), collectively referred to as GLS, are now known to be widely distributed [18–20]. The KGA and GAC isoforms share identical N-terminal and catalytic domains, encoded by exons 1–14 of the GLS gene, but have distinct C-termini derived from exon 15 in the case of GAC and exons 16–19 in the case of KGA [21]. Upregulation of GLS, in particular the GAC iso-form, is common in cancer cells and the degree of GLS overexpression correlates with both the degree of malignancy and the tumor grade in human breast cancer samples [22,23]. The GLS2 gene encodes a protein originally discovered and characterized in liver, which has thus been referred to as liver-type glutaminase and, more recently, as glutaminase 2 (GLS2) [15].

Both KGA and GAC can be activated by inorganic phosphate (Pi), and this activation correlates closely with a dimer-to-tetramer transition for each enzyme [7, 22]. As the concentration of Pi is raised the apparent catalytic constant, k_cat^app, increases and simultaneously the apparent Michaelis constant, K_m^app, decreases; consequently the catalytic efficiency rises dramatically, especially in the case of GAC [22]. x-ray crystal structures of GAC and KGA in different states indicate that the positioning of a key loop within each monomer (Glu312 to Pro329), located between the active site and the dimer–dimer interface, is critical for mediating tetramerization-induced activation [22,24]. Given the ability of Pi to promote tetramerization and activation of GAC and KGA, it has been proposed that the elevated mitochondrial Pi levels found under hypoxic conditions, which are commonly encountered in the tumor microenvironment, could be one trigger for GLS activation [22].

Oncogenic alterations affecting glutamine metabolism

At least two classes of cellular signals regulate glutamine metabolism, influencing both the expression level and the enzymatic activity of GLS. The transcription factor c-Myc can suppress the expression of microRNAs miR-23a and miR-23b and, in doing so, upregulates GLS (specifically GAC) expression [13,25]. Independent of changes in GAC expression, oncogenic diffuse B-cell lymphoma protein (Dbl), a GEF for Rho GTPases and oncogenic variants of downstream Rho GTPases are able to signal to activate GAC in a manner that is dependent on NF-κB [23]. Mitochondria isolated from Dbl- or Rho GTPase-transformed NIH-3T3 fibroblasts demonstrate significantly higher basal glutaminase activity than mitochondria isolated from non-transformed cells [23]. Furthermore, the enzymatic activity of GAC immunoprecipitated from Dbl-transformed cells is elevated relative to GAC from non-transformed cells, indicating the presence of activating post-translational modification(s) [23]. Indeed, when GAC isolated from Dbl-transformed cells is treated with alkaline phosphatase, basal enzymatic activity is dramatically reduced [23]. Collectively, these findings point to phosphorylation events underlying the activation of GAC in transformed cells. Similarly, phosphorylation-dependent regulation of KGA activity downstream of the Raf-Mek-Erk signaling axis occurs in response to EGF stimulation [24].

It is becoming clear that, in addition to c-Myc and Dbl, many other oncogenic signals and environmental conditions can impact cellular glutamine metabolism. Loss of the retinoblastoma tumor suppressor, for example, leads to a marked increase in glutamine uptake and catabolism, and renders mouse embryonic fibroblasts dependent on exogenous glutamine [26]. Cells transformed by KRAS also illustrate increased expression of genes associated with glutamine metabolism and a corresponding increased utilization of glutamine for anabolic synthesis [27]. In fact, KRAS signaling appears to induce glutamine dependence, since the deleterious effects of glutamine withdrawal in KRAS-driven cells can be rescued by expression of a dominant-negative GEF for Ras [28]. Downstream of Ras, the Raf-MEK-ERK signaling pathway has been implicated in the upregulation of glutamine uptake and metabolism [24,29]. A recent study using human pancreatic ductal adenocarcinoma cells identified a novel KRAS-regulated metabolic pathway, through which glutamine supports cell growth [30]. Proliferation of KRAS-mutant pancreatic ductal adenocarcinoma cells depends on GLS-catalyzed production of glutamate, but not on downstream deamination of glutamate to α-KG; instead, transaminase-mediated glutamate metabolism is essential for growth. Glutamine-derived aspartate is subsequently transported into the cytoplasm where it is converted by aspartate transaminase into oxaloacetate, which can be used to generate malate and pyruvate. The series of reactions maintains NADPH levels and thus the cellular redox state [30].

Other recent studies have revealed that another pathway for glutamine metabolism can be essential under hypoxic conditions, and also in cancer cells with mitochondrial defects or loss of the VHL tumor suppressor [31–35]. In these situations, glutamine-derived α-KG undergoes reductive carboxylation by IDH1 or IDH2 to generate citrate, which can be exported from mitochondria to support lipogenesis (Figure 1). Activation of HIF is both necessary and sufficient for driving the reductive carboxylation phenotype in renal cell carcinoma, and suppression of HIF activity can induce a switch from glutamine-mediated lipogenesis back to glucose-mediated lipogenesis [32,35]. Furthermore, loss of VHL and consequent downstream activation of HIF renders renal cell carcinoma cells sensitive to inhibitors of GLS [35]. Evidently, the metabolic routes through which glutamine supports cancer cell proliferation vary with genetic background and with microenvironmental conditions. Nevertheless, it is increasingly clear that diverse oncogenic signals promote glutamine utilization and furthermore that hypoxia, a common condition within poorly vascularized tumors, increases glutamine dependence.

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Consistent with the critical role of TCA cycle anaplerosis in cancer cell proliferation, a range of glutamine-dependent cancer cell lines are sensitive to silencing or inhibition of GLS [23,93]. Although loss of GLS suppresses proliferation, in some cases the induction of a compensatory anaplerotic mechanism mediated by pyruvate carboxylase (PC) allows the use of glucose- rather than glutamine-derived carbon for anaplerosis [93]. Low glutamine conditions render glioblastoma cells completely dependent on PC for proliferation; reciprocally, glucose deprivation causes them to become dependent on GLUD1, presumably as a mediator of glutamine-dependent anaplerosis [94]. These studies provide insight into the possibility of inhibiting glutamine-dependent TCA cycle anaplerosis (e.g., with 968 or BPTES) and indicate that high expression of PC could represent a means of resistance to GLS inhibitors.

In c-Myc-induced human Burkitt lymphoma P493 cells, entry of glucose-derived carbon into the TCA cycle is attenuated under hypoxia, whereas glutamine oxidation via the TCA cycle persists [95]. Upon complete withdrawal of glucose, the TCA cycle continues to function and is driven by glutamine. The proportions of viable and proliferating cell populations are almost identical in glucose-replete and -deplete conditions so long as glutamine is present. Inhibition of GLS by BPTES causes a decrease in ATP and glutathione levels, with a simultaneous increase in reactive oxygen species production. Strikingly, whereas BPTES treatment under aerobic conditions suppresses proliferation, under hypoxic conditions it results in cell death, an effect ascribed to glutamine’s critical roles in alleviating oxidative stress in addition to supporting bioenergetics.

In addition to deamidation, glutamine-derived carbon can also reach the TCA cycle through transamination [96], and recent studies indicate that inhibition of this process could be a promising strategy for cancer treatment [30,97,98]. The transaminase inhibitor amino-oxyacetate selectively suppresses proliferation of the aggressive breast cancer cell line MDA-MB-231 relative to normal human mammary epithelial cells, and similar effects were observed with siRNA knockdown of aspartate transaminase [97]. Treatment with amino-oxyacetate killed glutamine-dependent glioblastoma cells, in a manner that could be rescued by α-KG and was dependent on c-Myc expression [13]. Transaminase inhibitors have also been found to suppress both anchorage-dependent and anchorage-independent growth of lung carcinoma cells [98].

Reductive carboxylation

The central metabolic precursor for fatty acid biosynthesis is acetyl-CoA, which can be generated from pyruvate in the mitochondria by pyruvate dehydrogenase. Since acetyl-CoA cannot cross the inner mitochondrial membrane, it is exported to the cytosol via the citrate shuttle following its condensation with oxaloacetate in the TCA cycle (Figure 3). In the cytosol, citrate is converted back to acetyl-CoA and oxaloacetate in a reaction catalyzed by ATP citrate lyase. In addition to its synthesis from glycolytic pyruvate, citrate can also be generated by reductive carboxylation of α-KG [99]. Across a range of cancer cell lines, 10–25% of lipogenic acetyl-CoA is generated from glutamine via this reductive pathway; indeed, reductive metabolism is the primary route for incorporation of glutamine, glutamate and α-KG carbon into lipids [32]. Some of the reductive carboxylation of α-KG is catalyzed by cytosolic IDH1, as well as by mitochondrial IDH2 and/or IDH3.

In A549 lung carcinoma cells, glutamine dependence and reductive carboxylation flux increases under hypoxic conditions [32,34], such that glutamine-derived α-KG accounts for approximately 80% of the carbon used for de novo lipogenesis. Similarly, in melanoma cells, the major source of carbon for acetyl-CoA, citrate and fatty acids switches from glucose under normoxia to glutamine (via reductive carboxylation) under hypoxia [31]. The hypoxic switch to reductive glutamine metabolism is dependent on HIF, and constitutive activation of HIF is sufficient to induce the preferential reductive metabolism of α-KG even under normoxic conditions [32]. Tumor cells with mitochondrial defects, such as electron-transport chain mutations/inhibition, also use glutamine-dependent reductive carboxylation as the major pathway for citrate generation, and loss of electron-transport chain activity is sufficient to induce a switch from glucose to glutamine as the primary source of lipogenic carbon [33].

Together these studies indicate that mitochondrial defects/inhibition, and/or hypoxia, might sensitize cancer cells to inhibition of GLS. The fact that P493 cells are more sensitive to BPTES under hypoxic conditions could in part be explained by an increased reliance on glutamine-dependent reductive carboxylation for lipogenesis [95]. Intriguingly, cancer cells harboring neoenzymatic mutations in IDH1, which results in production of the oncometabolite 2-hydroxyglutarate, are also sensitized to GLS inhibition [100]. 2-hydroxyglutarate is generated primarily from glutamine-derived α-KG [100,101], and therefore tumors expressing mutant IDH might be especially susceptible to alterations in α-KG levels.

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As with all therapies, the potential side effects of strategies impacting glutamine metabolism must be seriously considered. The widespread use of l-asparaginase to lower plasma asparagine and glutamine concentrations in ALL patients demonstrates the potential for glutamine metabolism to be safely targeted, and also sheds light on potential toxicological consequences. For example, glutamine is known to be essential for the proliferation of lymphocytes, macrophages and neutrophils, and immunosuppression is a known side effect of l-asparaginase treatment, requiring close monitoring [11,105]. Evidence from early trials using glutamine-mimetic anti-metabolites, such as l-DON, indicates that these unselective molecules can cause excessive gastrointestinal toxicity and neurotoxicity. Within the brain, GLS converts glutamine into the neurotransmitter glutamate in neurons; astrocytes then take up synaptically released glutamate and convert it back to glutamine, which is subsequently transported back to neurons [106,107].

……

It has become clear during the past decade that altered metabolism plays a critical, in some cases even causal, role in the development and maintenance of cancers. It is now accepted that virtually all oncogenes and tumor suppressors impact metabolic pathways [5]. Furthermore, mutations in certain metabolic enzymes (e.g., isocitrate dehydrogenase, succinate dehydrogenase and fumarate hydratase) are associated with both familial and sporadic human cancers [113]. With this realization has come a renewed interest in the possibility of selectively targeting the metabolism of cancer cells as a therapeutic strategy. The use of l-asparaginase to treat ALL by depleting plasma asparagine and glutamine levels and the promising outcome of the first use of dichloroacetate (which acts, at least in part, through its inhibition of the metabolic enzyme pyruvate dehydrogenase kinase) in glioblastoma patients [114,115], support the notion that cancer metabolism can be safely and effectively targeted in the clinic. The metabolic adaptations of cancer cells must balance the requirements for modestly increased ATP synthesis, dramatically upregulated macromolecular biosynthesis and maintenance of redox balance. By serving as a carbon source for energy generation, a carbon and nitrogen source for biosynthesis and a precursor of the cellular antioxidant glutathione, glutamine is able to contribute to each of these requirements.

The countless combinations of genetic alterations that are found in human neo-plasias mean that there is not a single rigid metabolic program that is characteristic of all transformed cells. This perhaps explains why some current anti-metabolite chemotherapies (e.g., those targeting nucleotide synthesis) are effective only for certain malignancies. A deeper understanding of the metabolic alterations within specific genetic contexts will allow for better-targeted therapeutic interventions. Furthermore, it seems highly likely that combination therapies based on drug synergisms will be especially important for exploiting therapeutic windows within which cancer cells, but not normal cells, are impacted [37]. Glucose and glutamine metabolic pathways, for example, might be able to compensate for one another under some circumstances. When glucose metabolism is impaired in glioblastoma cells, glutamine catabolism becomes essential for survival [94]; reciprocally, suppression of GLS expression causes cells to become fully dependent on glucose-driven TCA cycle anaplerosis via PC [93]. The implication is that PC inhibition could synergize with GLS inhibition.

A topic warranting further investigation is the role that GLS2 plays in cellular metabolism. GLS, in particular the GAC isoform, is upregulated downstream of oncogenes and downregulated by tumor suppressors, and is essential for growth of many cancer cells. In contrast, GLS2 is activated by the ‘universal’ tumor suppressor p53, and furthermore is significantly downregulated in liver tumors and can block transformed characteristics of some cancer cells when overexpressed [116–118]. Emphasizing the importance of genetic context, it was recently reported that GLS2 is significantly upregulated in neuroblastomas overexpressing N-Myc [119]. There are various possible explanations for the apparently different roles of two enzymes that catalyze the same reaction. Because the regulation of GLS and GLS2 is distinct, they will be called up under different conditions. The two enzymes have different kinetic characteristics, and therefore might influence energy metabolism and antioxidant defense in different manners [20]. There is also evidence that GLS2 may act, directly or indirectly, as a transcription factor [118]. Finally, it is possible that the different interactions of GLS and GLS2 with other proteins are responsible for their apparently different roles.

Mitochondria as biosynthetic factories for cancer proliferation

Christopher S Ahn and Christian M Metallo

Cancer & Metabolism (2015) 3:1 http://dx.doi.org:/10.1186/s40170-015-0128-2

Unchecked growth and proliferation is a hallmark of cancer, and numerous oncogenic mutations reprogram cellular metabolism to fuel these processes. As a central metabolic organelle, mitochondria execute critical biochemical functions for the synthesis of fundamental cellular components, including fatty acids, amino acids, and nucleotides. Despite the extensive interest in the glycolytic phenotype of many cancer cells, tumors contain fully functional mitochondria that support proliferation and survival. Furthermore, tumor cells commonly increase flux through one or more mitochondrial pathways, and pharmacological inhibition of mitochondrial metabolism is emerging as a potential therapeutic strategy in some cancers. Here, we review the biosynthetic roles of mitochondrial metabolism in tumors and highlight specific cancers where these processes are activated.

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Recent characterizations of metabolic enzymes as tumor suppressors and oncogene-driven metabolic reprogramming have reinvigorated interest in cancer metabolism. Although therapies targeting metabolic processes have long been a staple in cancer treatment (e.g. inhibition of folate metabolism via methotrexate), the focused therapeutic potential surrounding these findings have generated a renewed appreciation for Otto Warburg’s work almost a century ago. Warburg observed that tumor cells ferment much of the glucose taken up during growth to lactate, thus using glycolysis as a major means of adenosine triphosphate (ATP) regeneration [1]. However, the observation of decreased respiration in cancer cells and idea that “the respiration of all cancer cells is damaged” belies the critical role of mitochondria in biosynthesis and cell survival [1]. On the contrary, functional mitochondria are present in all proliferative cells within our body (including all tumors), as they are responsible for converting the diverse nutrients available to cells into the fundamental building blocks required for cell growth. These organelles execute numerous functions in cancer cells to promote tumor growth and survival in response to stress. Here, we outline the critical biosynthetic functions served by mitochondria within tumors (Figure 1). Although many of these functions are similarly important in normal, proliferating cells, we have attempted to highlight potential points where mitochondrial metabolism may be therapeutically targeted to slow cancer growth. This review is organized by specific metabolic pathways or processes (i.e., glucose metabolism and lipogenesis, amino acid metabolism, and nucleotide biosynthesis). Tumors or cancer cell types where enzymes in each pathway have been specifically observed to by dysregulated are described within the text and summarized in Table 1.

Figure 1

Biosynthetic nodes within mitochondria. Metabolic pathways within mitochondria that contribute to biosynthesis in cancer and other proliferating cells. TCA metabolism and FOCM enable cells to convert carbohydrates and amino acids to lipids, non-essential amino acids, nucleotides (including purines used for cofactor synthesis), glutathione, heme, and other cellular components. Critical biosynthetic routes are indicated by yellow arrows. Enzymatic reactions that are dependent on redox-sensitive cofactors are depicted in red. https://static-content.springer.com/image/art%3A10.1186%2Fs40170-015-0128-2/MediaObjects/40170_2015_128_Fig1_HTML.gif

Table 1

Overview of mitochondrial biosynthetic enzymes important in cancer

TCA cycle, anaplerosis, and AcCoA metabolism

Cancers in which three or more mitochondrial enzymes have been studied and found to be differentially regulated (or mutated, as indicated) in cancers vs. control groups are included. Dysregulation of each enzyme was demonstrated in clinical tumors samples, animal models, or cell lines at the levels of genes, mRNA, protein, metabolites, and/or flux.

Figure 2

Coordination of carbon and nitrogen metabolism across amino acids. Glutamate and aKG are key substrates in numerous transamination reactions and can also serve as precursors for glutamine, proline, and the TCA cycle. Mitochondrial enzymes catalyzing these reactions are highlighted in blue, and TCA cycle intermediates are highlighted in orange (pyruvate enters the TCA cycle as acetyl-CoA or oxaloacetate).
https://static-content.springer.com/image/art%3A10.1186%2Fs40170-015-0128-2/MediaObjects/40170_2015_128_Fig2_HTML.gif

Figure 3

Biosynthetic sources for purine and pyrimidine synthesis. Sources and fates of nitrogen, carbon, and oxygen atoms are colored as indicated. Italicized metabolites can be sourced from the mitochondria or cytosol. The double bond formed by the action of DHODH/ubiquinone is also indicated. https://static-content.springer.com/image/art%3A10.1186%2Fs40170-015-0128-2/MediaObjects/40170_2015_128_Fig3_HTML.gif

Mitochondria operate as both engine and factory in eukaryotes, coordinating cellular energy production and the availability of fundamental building blocks that are required for cell proliferation. Cancer cells must therefore balance their relative bioenergetic and biosynthetic needs to grow, proliferate, and survive within the physical constraints of energy and mass conservation. In contrast to quiescent cells, which predominantly use oxidative mitochondrial metabolism to produce ATP and uptake glucose at much lower rates than proliferating cells, tumor cells exhibit increased glycolytic rates to provide an elevated flux of substrate for biosynthetic pathways, including those executed within mitochondria. Given these higher rates of nutrient utilization, metabolic flux through mitochondrial pathways and the associated ROS production can often be higher in cancer cells. Not surprisingly, activation of cellular antioxidant response pathways is commonly observed in cancer or subpopulations of cells within tumors [46,78]. Cellular compartmentalization affords a degree of protection from such damaging side products of metabolism, and methods which are able to deconvolute the relative contributions of each cellular compartment (e.g. mitochondria, cytosol, peroxisome, etc.) to cancer metabolism will be crucial to more completely understand the metabolism of cancer cells in the future [74,79]. Ultimately, while mitochondrial dysregulation is widely considered to be a hallmark of cancer, numerous mitochondrial functions remain critical for tumor growth and are emerging as clinical targets.

Following this point, it comes as no surprise that mitochondrial metabolism is highly active in virtually all tumors (i.e., cancer cells, stroma, or both), and investigators have begun targeting these pathways to explore potential efficacy. Indeed, some evidence suggests that biguanides such as metformin or phenformin may limit tumor incidence and burden in humans and animals [80,81]. These effects are presumably due, at least in part, to complex I inhibition of the ETC, which significantly perturbs mitochondrial function [82,83]. However, more insights are needed into the mechanisms of these compounds in patients to determine the therapeutic potential of targeting this and other components of mitochondria. In developing new therapies that target cancer metabolism, researchers will face challenges similar to those that are relevant for many established chemotherapies since deleterious effects on normal proliferating cells that also depend on mitochondrial metabolism (and aerobic glycolysis) are likely to arise.

As we acquire a more detailed picture of how specific genetic modifications in a patient’s tumor correlate with its metabolic profile, opportunities for designing targeted or combinatorial therapies will become increasingly apparent. Cancer therapies that address tumor-specific mitochondrial dysregulation and dysfunction may be particularly effective. For example, some cancer cells harbor mutations in TCA enzymes (e.g., FH, SDH, IDH2) or regulatory proteins that control mitophagy (i.e., LKB1) [84]. Such tumors may be compromised with respect to some aspects of mitochondrial biosynthesis and dependent on alternate pathways for growth and/or survival such that synthetically lethal targets emerge. Ultimately, such strategies will require clinicians and researchers to coordinate metabolic, biochemical, and genetic information in the design of therapeutic strategies.

David Terrano, M.D., Ph.D. commented on your update

“Not well versed in Nat peptides so I could not say. I also hesitate with any PNAS paper because those in their academy tend to have a fast track to publication. It has been that way since at least early 2000’s wh n I began research. I don’t doubt their goal and approach (this same group leads the way in methylation-based diagnosis of CNS neoplasms, which is apparently highly accurate). But when I see “dying cells” I know what that means biochemically and look for those hallmarks. Organ specific oligonucleosomes would be a nice cell death surrogate. “

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single nucleotide repair and tunable DNA-directed assembly of nanomaterials

Posted in Biological Networks, Gene Regulation and Evolution, Cell Biology, Chemical Biology and its relations to Metabolic Disease, Computational Biology/Systems and Bioinformatics, Curation, Disease Biology, Small Molecules in Development of Therapeutic Drugs, Genetics & Innovations in Treatment, Genetics & Pharmaceutical, Genome Biology, Genomics Pharmacy, Innovations, tagged DNA repair, DNAzyme, enzyme cascades, nanomaterials, nucleotides, tunable on November 1, 2015| Leave a Comment »

single nucleotide repair and tunable DNA-directed assembly of nanomaterials

Larry H. Bernstein, MD, FCAP, Curator

LPBI

Expanding DNAzyme functionality through enzyme cascades with applications in single nucleotide repair and tunable DNA-directed assembly of nanomaterials

Yu Xiang,^a Zidong Wang,^b Hang Xing^a and Yi Lu*^a^b Show Affiliations

Chem. Sci., 2013, 4, 398-404 DOI: <a href="http://dx.doi.org:/10.1039/C2SC20763J”>http://dx.doi.org:/10.1039/C2SC20763J

Many biological functions require two or more enzymes working together in cascades. While many examples of protein and RNA enzyme cascades are known, few enzyme cascades containing solely DNAzymes have been reported. Herein we demonstrate the combination of an 8–17 DNAzyme with RNA cleavage activity and an E47 DNAzyme with DNA ligation activity to achieve a new function of single ribonucleotide repair in DNA while maintaining the integrity of the original DNA sequence, which is difficult for a single DNAzyme to achieve. In addition, this method is applied to modify the sequences of DNA strands immobilized on the surface ofnanoparticles to control the DNA-directed assembly selectively and sequentially. Such an approach can be applied to other DNAzymes with different activities to expand the functions of DNAzymes and the scope of their applications.

Graphical abstract: Expanding DNAzyme functionality through enzyme cascades with applications in single nucleotide repair and tunable DNA-directed assembly of nanomaterials

The discovery of deoxyribozymes (DNAzymes) with enzymatic activity in the 1990s1,2 has demonstrated that DNA molecules are not simply inert biopolymers for genetic storage; they can be active catalysts as well.3–8 Since then, many DNAzymes have been obtained with catalytic functions such as cleavage,2,9–13 ligation,14–16 phosphorylation,17 adenylation18 or depurination19 of nucleic acids, as well as other reactions including porphyrin metallation, C–C bond formation, nucleopeptide linkage formation, oxygen transfer and thymine dimer repair.20–26 Because DNAzymes are facile to synthesize and more stable than protein and RNA enzymes, they have been widely used in applications such as nanomaterial assembly,27,28 biosensing,29–31 logical computing,32 nanomachine engineering,33 antiviral or gene therapy,34 and in vitro RNA manipulation.35 Despite these successes, the application of DNAzymes is limited by the narrower range of catalytic functionality compared to protein enzymes. One possible approach to addressing this issue would be to combine enzymes with different reactivities to form a cascade of successive enzymatic reactions, which together create new functionality. Indeed, many such examples exist in biology, since nearly all important biological functions, such as the pathways involved in DNA repair and protein synthesis, require a cascade of multiple protein enzymes to carry out their full function. In contrast, little has been reported about the use of DNAzyme cascades to realize enhanced functionality. Such a strategy could expand the functionality of DNAzymes to a level more on par with protein and RNA enzymes, which should greatly increase the range of possible applications.

One such application is single nucleotide repair, i.e., excision of a misincorporated ribonucleotide in single-stranded DNA and subsequent insertion of the corresponding deoxyribonucleotide at the excision site. The misincorporation of ribonucleotides into DNA strands can occur from exposure to external oxidizing agents or ionizing radiation,36 or spontaneously during DNA replication.37 Misincorporation of ribonucleotide can distort the structure of DNA,38 reduce its stability,39 and interfere with the normal interaction between DNA and DNA polymerases.40 In fact, the overexpression of DNA polymerases that are prone to ribonucleotide misincorporation has been linked to many cancers, including ovarian, prostate, breast and colon cancers.41 In nature, protein enzymes such as RNase H and FEN-1 can efficiently excise misincorporated ribonucleotides in DNA by cleaving the DNA at the ribonucleotide site and then restoring the correct deoxyribonucleotides by DNA polymerases,42,43 which is an example of an enzyme cascade. It would be interesting to nd out if a similar function could be achieved through DNAzyme cascades.

Another potential application is in tuning the properties of DNA-functionalized nanomaterials. For example, DNA-functionalized gold nanoparticles27 have emerged as an attractive platform for biosensing,32,44–50 nanomedicine,45 and as building blocks for controlled nanoassemblies.51–55 Although much research has been focused on the surface modification of gold nanoparticles with DNA for various applications, there are still limited methods to modify the sequences of DNA already immobilized on gold nanoparticles in order to make the properties of the DNA-modified nanomaterials tunable aer fabrication. The use of DNAzymes is a promising approach for DNA modification on nanomaterials56 due to the excellent stability of DNAzymes and their smaller size compared to protein enzymes, thereby minimizing steric effects between the enzyme and the DNA in order to avoid reduction in reaction efficiency. However, it is still very challenging to modify a specific DNA sequence on multiple-DNA-functionalized nanomaterials to tune their functions in a selective and sequential fashion.

Herein, we demonstrate a cascade of two DNAzymes with RNA cleavage and DNA ligation activities, respectively, in order to carry out single nucleotide repair or selective sequence modification of DNA. In a one-pot reaction, a single misincorporated ribonucleotide in a DNA strand was converted to the corresponding deoxyribonucleotide while maintaining sequence integrity. Furthermore, the sequences of DNA strands immobilized on multiple functional nanoparticles were successfully modified in order to control and alter the DNAdirected assembly of nanoparticles in a stepwise and selective fashion.

Results and discussion To demonstrate that single nucleotide repair in DNA can be achieved by the cascade of two DNAzymes, we used a 26-nt DNA strand (O1) containing a misincorporated cytidine (rC) ribonucleotide as an example. The goal was to convert the rC in O1 into a deoxycytidine (C), as seen in O4 (Fig. 1a), while maintaining the integrity of the DNA sequence. The DNAzymes 17Em1 (Fig. 1a, blue) with RNA cleavage activity2,57–59 and E47 (Fig. 1a, red) with DNA ligation activity14,60 were chosen as the cascade pair in this study. The 17Em1 DNAzyme catalyzes the hydrolysis of the 30 phosphodiester linkage of the internal rC in the DNA strand when metal ion cofactors such as Pb2+ and Zn2+ are present (Fig. S1a in ESI†). On the other hand, the E47 DNAzyme can induce the catalytic ligation of the 50 –OH of the DNA substrate with another 30 -phosphorylated DNA strand (activated by imidazole)14 in the presence of Cu2+ or Zn2+ as the metal cofactor (Fig. S1b in ESI†). Therefore, by sequential cleavage and ligation reactions catalyzed by these two DNAzymes on O1 containing rC, O1 could first be cleaved at the 30 phosphodiester of the rC by 17Em1 and then undergo ligation at the cleavage site with another 30 -phosphorylated DNA strand of an identical sequence (except with deoxyribonucleotide C in place of ribonucleotide rC) by E47. The product O4 has a sequence identical to the starting strand O1, with the rC replaced with C.

Fig. 1 (a) Conversion of a single ribonucleotide (rC) in a DNA strand O1 to a deoxyribonucleotide (C) by the cascade of DNAzymes 17Em1 and E47: O1 is cleaved by 17Em1 to afford products of O2 and Oc; O2 is then ligated with O3 (activated by imidazole) to form O4 by E47. (b) Sequence modification of a DNA strand O5 to O4 through a similar protocol by the cascade of DNAzymes 17Em2 and E47.

Initially, 30 -fluorescein-labeled O1 was treated with 17Em1 to form DNA duplex O1-17Em1 via 18 matched base pairs (9 on each binding arm). In the presence of Pb2+, O1 was efficiently cleaved by 17Em1 into fragments O2 and Oc, resulting in the dehybridization of the duplex because the melting temperature of the duplex between 17Em1 and O2 or between 17Em1 and Oc is below room temperature (Fig. 1a). The fluorescence image after polyacrylamide gel electrophoresis (PAGE) suggested the complete cleavage of O1 and formation of O2 (Fig. 2a, lane 1 and 2 for O1 and O2, respectively), while Oc was not visible on the gel due to the lack of a fluorescein label. The cleavage reaction product O2 was also confirmed by the result from matrixassisted laser desorption ionization-time of flight (MALDI-TOF) mass spectrum (Table 1 and Fig. S2 in ESI†). Control experiments using a DNAzyme of a different sequence (17Em2) or without Pb2+ showed negligible cleavage of the substrate O1 (Fig. S3 in ESI†) due to the specificity of the DNAzyme and the essential role of the metal ion cofactor.2,57–59 Subsequently, without any purification of O2 from the mixture solution after the previous cleavage step, E47 and 30 -phosphorylated O3 (imidazole-activated) were added into the solution to generate another DNA complex O2–O3-E47, which gave O4 as the product after the E47-catalyzed ligation reaction in the presence of Cu2+ (Fig. 1a).14,60 The formation of O4 was confirmed by both fluorescent PAGE (Fig. 2a, the upper band of lane 3) and MALDITOF MS (Table 1 and Fig. S2 in ESI†), while some unreacted O2 was also observed on the gel (Fig. 2a, the lower band of lane 3). Here, O3 was invisible due to the lack of a fluorescein label. Considerably lower levels of ligation between O2 and O3 were observed if either E47 or Cu2+ was absent (Fig. S3 in ESI†). Together these results indicate that the reactions catalyzed by the DNAzyme cascade were achieved through a one-pot reaction without isolation and purification of the intermediate O2.

Fig. 2 (a) Fluorescent PAGE (20% denaturing gel) images of the transformation from O1 to O4 by DNAzymes 17Em1 and E47. Lanes in (a): 1, O1; 2, 1 after cleavage by 17Em1 to yield O2 and Oc in the presence of Pb2+; 3, 2 after ligation to yield O4 by E47 in the presence of O3 and Cu2+; 4, 2 after ligation to yield O4 + 8A by E47 in the presence of O3 + 8A and Cu2+; 5, O4 in the presence of Pb2+ and 17Em1. (b) Fluorescent PAGE images of the transformation from O5 to O4 by 17Em2 and E47: Lanes in (b): 1, O5; 2, 1 after cleavage by 17Em2 to yield O2 in the presence of Pb2+; 3, 2 after ligation to yield O4 by E47 in the presence of O3 and Cu2+; 4, 2 after ligation to yield O4 + 8A by E47 in the presence of O3 + 8A and Cu2+; 5, O4 in the presence of Pb2+ and 17Em2

To provide further confirmation of the above successful conversion of rC in O1 to C in O4, while keeping other sequences identical, a longer O3 + 8A (O3 extended by A8 at 50 ) was used in place of O3 (Fig. 1a). Under the same conditions, a longer product O4 + 8A was obtained (Fig. 2a, the upper band of lane 4) with slower gel migration compared to O4 (Fig. 2a, the upper band of lane 3), suggesting that the ligation reaction occurred mostly between O2 and imidazole-activated O3, and not between O2 and un-activated Oc (Oc is the product from the previous cleavage reaction of O1 and 17Em1), in which case a band with the same migration as O4 would have been observed. The presence of C rather than rC in the product O4 was supported by the lower molecular weight of O4 in the MALDI-TOF mass spectrum as compared to that of O1 (Table 1), as well as the increased resistance to hydrolysis of O4 even in the presence of Pb2+ and 17Em1 (Fig. 2a, lane 5), which can catalyze the cleavage of a substrate containing an internal ribonucleotide linkage (O1),2,57 but not a substrate containing entirely deoxyribonucleotides (O4).

Table 1 Measured and calculated molecular weight (m/z) in MALDI-TOF mass spectra of 30 -fluorescein-labeled DNAs (O1, O2, O4 and O5). For full spectra, see Fig. S2 in ESI.†

DNA O1 O2 O4 O5 Measured 8831.6 4824.1 8812.3 8819.1 Calculated 8831.9 4826.3 8815.9 8821.9duct O4 + 8A was obtained (Fig. 2a, the upper band of lane 4) with slower gel migration compared to O4 (Fig. 2a, the upper band of lane 3), suggesting that the ligation reaction occurred mostly between O2 and imidazole-activated O3, and not between O2 and un-activated Oc (Oc is the product from the previous cleavage reaction of O1 and 17Em1), in which case a band with the same migration as O4 would have been observed. The presence of C rather than rC in the product O4 was supported by the lower molecular weight of O4 in the MALDI-TOF mass spectrum as compared to that of O1 (Table 1), as well as the increased resistance to hydrolysis of O4 even in the presence of Pb2+ and 17Em1 (Fig. 2a, lane 5), which can catalyze the cleavage of a substrate containing an internal ribonucleotide linkage (O1),2,57 but not a substrate containing entirely deoxyribonucleotides (O4).

In addition to the single nucleotide repair functionality, it is also possible to use this methodology to edit the sequence of a DNA strand, which was used to convert the DNA strand O5 into O4 using the same cascade and conditions as before (Fig. 1b and S4 in ESI†). The product O4 was confirmed by PAGE (Fig. 2b) and MALDI-TOF MS (Table 1 and Fig. S2 in ESI†) and found to be identical to that obtained from the method in Fig. 1a.

Encouraged by the above results, we applied this method to modify the sequence of DNA immobilized on gold nanoparticles27 (AuNPs) to control the DNA-directed assembly of the AuNPs in a selective manner. DNA-functionalized gold AuNPs have been used in a variety of applications due to both their unique properties and the sequence-dependent hybridization of ssDNA immobilized on the AuNPs for controlled assembly.51–55 As shown in Fig. 3, when AuNPs are functionalized by complementary DNAs, the AuNPs can assemble into an “aggregated” state via DNA hybridization, which shows red-shifted and broadened absorption spectra compared to AuNPs functionalized by non-complementary DNAs.

Fig. 3 Assembly of two types of DNA-functionalized gold nanoparticles. If the sequences of the two DNAs are not complementary, the gold nanoparticles are in a “dispersed” state and exhibit a red color with a sharp absorption band peaked around 532 nm (left). In contrast, the assembly of the gold nanoparticles with complementary DNAs causes the formation of an “aggregated” state with a broad absorption band around 600 nm (right).

By modifying the sequences of DNA on the AuNPs, the assembly of the particles can be effectively controlled. Although methods for fabrication of DNAfunctionalized AuNPs have been developed,27 there are still limited methods to modify the DNA sequences already immobilized on AuNPs in order to tune their functions. It is even more challenging to modify a specific DNA sequence on multiply-functionalized AuNPs with different DNA sequences on each nanoparticle. Selective modification can allow each different function of the AuNP to be controlled in a selective fashion for potential applications.

AuNPs of 13 nm diameter were functionalized with DNA molecules via 30 -end thiols and used for this study. The formation of the AuNP assembly was confirmed by TEM images (Fig. S5 in ESI†) and characterized by large changes in absorption spectra27 (A700/A532 changed from <0.15 to >0.50 as illustrated in Fig. 5 and Table S1 in ESI†). As shown in Fig. 4, O6- functionalized AuNPs (red) were found to be able to form aggregates with O9-functionalized AuNPs (blue) via DNAdirected assembly through 12 complementary base pairs (Fig. 5 and S5 and Table S1†), but not with O10-functionalized AuNPs (purple), because of the 4 mismatched base pairs in the middle of the binding arm (Fig. 5 and S5 and Table S1†). After being treated with 17Em2 and Pb2+, the O6 on the surface of AuNPs was cleaved and converted to O8, which could not hybridize with either O9 or O10 efficiently. Thus no DNA-directed assembly was observed between the resulting O8-functionalized AuNPs and either O9- or O10-functionalized AuNPs (Fig. 5 and S5 and Table S1†). However, after a subsequent ligation reaction catalyzed by E47 in the presence of imidazole-activated O3 and Cu2+, O8 on the surface of AuNPs could be extended to O7, making the AuNPs capable of assembling with O10-, but not O9- functionalized AuNPs (Fig. 5 and S5 and Table S1†).

Fig. 4 Controlling the assembly of DNA-functionalized gold nanoparticles via cascade-mediated modification of the DNA sequences. The solid and dashed lines indicate the successful and unsuccessful formation of aggregates, respectively. The inset shows the assembly of AuNPs modified with the complementary strands O6 and O9 (top) and the lack of assembly between AuNPs modified with noncomplementary strands O9 and O7 (obtained by treating O6-functionalized AuNPs with 17Em2 and E47) (bottom)

Fig. 5 Absorption spectra of O6-, O7- or O8-functionalized AuNPs in the presence of O9- and O10-functionalized AuNPs, respectively. The red-shift of the peak indicates the formation of AuNP aggregations due to the hybridization of complementary DNAs on the AuNPs.27 For the ratios of absorbance (A700/A532), see Table S1 in ESI.†

Interestingly, the product, O7-functionalized AuNPs, showed inverse characteristics in the formation of DNA-directed assembly with O10- and O9-functionalized AuNPs, compared to the original O6-functionalized AuNPs. TEM images of O6- functionalized AuNPs, either with or without treatment with 17Em2/E47, mixed with O10-functionalized AuNPs, are displayed in the inset of Fig. 4. These results clearly demonstrate that the ability to edit and replace DNA on AuNPs allows for exquisite programmable control over the assembly of nanoparticles.

Taking advantage of the specificity of DNAzyme to its nucleic acid substrates by complementary base pairing in the binding arms, selective modification of DNA sequences on surface of multiple functional AuNPs was also achieved in this work. As depicted in Fig. 6, O6 (red) and O11 (blue) bi-functional AuNPs could be modified by 17Em1, 17Em2 and E47 selectively and sequentially. As shown in Fig. 6, AuNPs capable of forming DNA-directed assembly with both (A and E), either (B, C and F), or neither (D) of the O9- and O10-functionalized AuNPs could be obtained by monitoring the significant increase of A700/A532 as indication of assembly formation (Fig. 7 and Table S2 in ESI†). Such a result, which is challenging to achieve by other techniques, can be used for the construction of tunable nanoassemblies for various applications.

Fig. 6 (a) Scheme showing stepwise modification of DNA sequences on multiply-functional gold nanoparticles by the collaboration of 17Em1 or 17Em2 and E47. (b) DNA sequences of O6–O8 and O11.

Fig. 7 Absorption spectra of DNA-functionalized gold nanoparticles (A–F) (Fig. 6a) in the presence of O9- and O10-functionalized AuNPs, respectively. The red-shift of the peak indicates the formation of AuNP aggregations due to the hybridization of complementary DNAs on the AuNPs.27 For the ratios of absorbance (A700/A532), see Table S2 in ESI.†

In summary, by putting together a cascade of two DNAzymes with cleaving and ligating activities, we have generated a new functionality for effective DNA modification. This function was applied in the conversion of a single misincorporated ribonucleotide into the corresponding dexoyribonucleotide in DNA and the modification of DNA sequences on the surface of gold nanoparticles to modify and control their self-assembly through DNA hybridization. The results suggest that combining DNAzymes with different catalytic activities may achieve more interesting functions and thus broaden the applications of DNAzymes.

Notes and references

1 D. L. Robertson and G. F. Joyce, Nature, 1990, 344, 467–468.

2 R. R. Breaker and G. F. Joyce, Chem. Biol., 1994, 1, 223–229.

3 D. Sen and C. R. Geyer, Curr. Opin. Chem. Biol., 1998, 2, 680– 687.

4 Y. F. Li and R. R. Breaker, Curr. Opin. Struct. Biol., 1999, 9, 315–323.

5 Y. Lu, Chem.–Eur. J., 2002, 8, 4588–4596.

6 R. R. Breaker, Nature, 2004, 432, 838–845.

7 K. Schlosser and Y. F. Li, Chem. Biol., 2009, 16, 311–322.

8 S. K. Silverman, Acc. Chem. Res., 2009, 42, 1521–1531.

9 R. R. Breaker and G. F. Joyce, Chem. Biol., 1995, 2, 655–660.

10 N. Carmi, S. R. Balkhi and R. R. Breaker, Proc. Natl. Acad. Sci. U. S. A., 1998, 95, 2233–2237.

11 A. R. Feldman and D. Sen, J. Mol. Biol., 2001, 313, 283–294.

12 J. W. Liu, A. K. Brown, X. L. Meng, D. M. Cropek, J. D. Istok, D. B. Watson and Y. Lu, Proc. Natl. Acad. Sci. U. S. A., 2007, 104, 2056–2061.

13 M. Chandra, A. Sachdeva and S. K. Silverman, Nat. Chem. Biol., 2009, 5, 718–720.

14 B. Cuenoud and J. W. Szostak, Nature, 1995, 375, 611–614.

15 A. Sreedhara, Y. F. Li and R. R. Breaker, J. Am. Chem. Soc., 2004, 126, 3454–3460.

16 W. E. Purtha, R. L. Coppins, M. K. Smalley and S. K. Silverman, J. Am. Chem. Soc., 2005, 127, 13124–13125.

17 Y. F. Li and R. R. Breaker, Proc. Natl. Acad. Sci. U. S. A., 1999, 96, 2746–2751.

18 Y. F. Li, Y. Liu and R. R. Breaker, Biochemistry, 2000, 39, 3106–3114.

19 T. L. Sheppard, P. Ordoukhanian and G. F. Joyce, Proc. Natl. Acad. Sci. U. S. A., 2000, 97, 7802–7807.

20 Y. F. Li and D. Sen, Nat. Struct. Biol., 1996, 3, 743–747

Citations

Portable and quantitative monitoring of heavy metal ions using DNAzyme-capped mesoporous silica nanoparticles with a glucometer readout

Libing Fu, Junyang Zhuang, Wenqiang Lai, Xiaohua Que, Minghua Lu and Dianping Tang

J. Mater. Chem. B, 2013, 1, 6123 DOI: <a href="http://dx.doi.org:/10.1039/c3tb21155j”>http://dx.doi.org:/10.1039/c3tb21155j

A Label-Free Electrochemical Aptasensor for the Analysis of The Potassium Ion

Xiaoxia Ji, Jialei Li and Chengjian Yang

Journal of Immunoassay and Immunochemistry, 2015, 36, 162 DOI: http://dx.doi.org:/10.1080/15321819.2014.915221

In vitro selection of DNA-cleaving deoxyribozyme with site-specific thymidine excision activity

M. Wang, H. Zhang, W. Zhang, Y. Zhao, A. Yasmeen, L. Zhou, X. Yu and Z. Tang

Nucleic Acids Research, 2014, 42, 9262 DOI: http://dx.doi.org:/10.1093/nar/gku592

Applications of Synchrotron-Based Spectroscopic Techniques in Studying Nucleic Acids and Nucleic Acid-Functionalized Nanomaterials

Peiwen Wu, Yang Yu, Claire E. McGhee, Li Huey Tan and Yi Lu

Adv. Mater., 2014, 26, 7849 DOI: http://dx.doi.org:/10.1002/adma.201304891

DNA as a Powerful Tool for Morphology Control, Spatial Positioning, and Dynamic Assembly of Nanoparticles

Li Huey Tan, Hang Xing and Yi Lu

Acc. Chem. Res., 2014, 47, 1881 DOI: http://dx.doi.org:/10.1021/ar500081k

Kun Xiong, Li Li, Li Zhang, Wei Ding, Lishan Peng, Yao Wang, Siguo Chen, Shiyu Tan and Zidong Wei

J. Mater. Chem. A, 2015,3, 1863-1867

Abstract

One-dimensional NiMo₂C nanowires with a weaker hydrogen binding energy were directly grown onto Ni foam for enhancing the HER

A one-step, cost-effective green method to in situ fabricate Ni(OH)₂ hexagonal platelets on Ni foam as binder-free supercapacitor electrode materials

Lingjie Li, Jing Xu, Jinglei Lei, Jie Zhang, Frank McLarnon, Zidong Wei, Nianbing Li and Fusheng Pan

J. Mater. Chem. A, 2015,3, 1953-1960

Abstract

The Ni(OH)₂ hexagonal platelets were in situ fabricated on Ni foam as a binder-free supercapacitor electrode material with high performance and excellent cycling stability by a one-step, cost-effective, green hydrothermal treatment of three-dimensional (3D) Ni foam in a 15 wt% H₂O₂ aqueous solution.

An extraordinarily stable catalyst: Pt NPs supported on two-dimensional Ti₃C₂X₂ (X = OH, F) nanosheets for oxygen reduction reaction

Xiaohong Xie, Siguo Chen, Wei Ding, Yao Nie and Zidong Wei

Chem. Commun., 2013,49, 10112-10114

Abstract

Pt nanoparticles supported on Ti₃C₂X₂ (X = OH, F) nanosheets show enhanced ORR stability and activity compared with the commercial Pt/C catalyst.

Pd-induced Pt(IV) reduction to form Pd@Pt/CNT core@shell catalyst for a more complete oxygen reduction

Meirong Xia, Ying Liu, Zidong Wei, Siguo Chen, Kun Xiong, Li Li, Wei Ding, Jinsong Hu, Li-Jun Wan, Rong Li and Shahnaz Fatima Alvia

J. Mater. Chem. A, 2013,1, 14443-14448

Abstract

A facile and controllable process for preparing Pd@Pt/CNT core@shell catalysts for the oxygen reduction reaction (ORR) via Pd-induced Pt(IV) reduction on Pd/CNT.

Acid- and base-functionalized core-confined bottlebrush copolymer catalysts for one-pot cascade reactions

Linfeng Xiong, Hui Zhang, Aiqing Zhong, Zidong He and Kun Huang

Chem. Commun., 2014,50, 14778-14781

Abstract

A novel method that enables the formation of core-confined bottlebrush copolymers (CCBCs) as catalyst supports for one-pot cascade reactions is reported for the first time.

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Isoenzymes in cell metabolic pathways

Posted in Best evidence, Biomarkers & Medical Diagnostics, Chemical Biology and its relations to Metabolic Disease, tagged ATP, Cori cycle, ECT, gluconeogemesis, Glycolysis, hydride transfer, isoenzymes, LDH, liver, MDH, NADH, nucleotides, ocular lens, TCA cycle on October 6, 2014| Leave a Comment »

Larry H. Bernstein, MD, FCAP, Author and Curator

Isozymes

An example of an isozyme is glucokinase, a variant of hexokinase which is not
inhibited by glucose 6-phosphate. Its different regulatory features and lower
affinity for glucose (compared to other hexokinases), allows it to serve different
functions in cells of specific organs, such as

control of insulinrelease by the beta cells of the pancreas, or
initiation ofglycogen synthesis by liver
Both of these processes must only occur when glucose is abundant,or
problems occur.

Isozymes or Isoenzymes are proteins with different structure which catalyze
the same reaction. Frequently they are oligomers made with different
polypeptide chains, so they usually differ in regulatory mechanisms and in
kinetic characteristics.

From the physiological point of view, isozymes allow the existence of similar
enzymes with different characteristics, “customized” to specific tissue
requirements or metabolic conditions.

One example of the advantages of having isoenzymes for adjusting the
metabolism to different conditions and/ or in different organs is the following:

Glucokinase and Hexokinase are typical examples of isoenzymes. In fact,
there are four Hexokinases: I, II, III and IV. Hexokinase I is present in all
mammalian tissues, and Hexokinase IV, aka Glucokinase, is found mainly
in liver, pancreas and brain.

Both enzymes catalyze the phosphorylation of Glucose:

Glucose + ATP —–à Glucose 6 (P) + ADP

Hexokinase I has a low Km and is inhibited by glucose 6 (P). Glucokinase
is not inhibited by Glucose 6 (P) and his Km is high. These two facts
indicate that the activity of glucokinase depends on the availability
of substrate and not on the demand of the product.

Since Glucokinase is not inhibited by glucose 6 phosphate, in
conditions of high concentrations of glucose this enzyme
continues phosphorylating glucose, which can be used for
glycogen synthesis in liver. Additionally, since Glucokinase
has a high Km, its activity does not compromise the supply
of glucose to other organs; in other words, if Glucokinase
had a low Km, and since it is not inhibited by its product, it
would continue converting glucose to glucose 6 phosphate
in the liver, making glucose unavailable for other organs
(remember that after meals, glucose arrives first to the liver
through the portal system).

The enzyme Lactate Dehydrogenase is made of two (H-
and M-) sub units, combined in different Permutations
and Combinations depending on the tissue in which it
is present as shown in table,

Type	Composition	Location
LDH1	HHHH	Heart and Erythrocyte
LDH2	HHHM	Heart and Erythrocyte
LDH3	HHMM	Brain and Kidney
LDH4	HMMM	Skeletal Muscle and Liver
LDH5	MMMM	Skeletal Muscle and Liver

While isozymes may be almost identical in function
(defined by Michaelis constant, KM)
they differ in amino acidsubstitutions that change the
electric charge of the enzyme (such as replacing
aspartic acid with glutamic acid)
The sum of zwitterion charges result in identifyjng
difference inmigratiion toward the anode by gel
electrophoresis, and this forms the basis for the use
of isozymes as molecular markers.
To identify isozymes, a crude protein extract is made by
grinding animal or plant tissue with an extraction buffer,
and the components of extract are separated according
to their charge by gel electrophoresis.
They were classically purified by ion-exchange column
chromatography after first precipitation with ammonium
sulfate, followed by dialysis.

The cytochrome P450 isozymes play important roles in
metabolism and steroidogenesis. The multiple forms of
phosphodiesterase also play major roles in various
biological processes.

These isoforms of the enzyme are unequally distributed
in the various cells of an organism.

Further the main isoenzymes may have closely grouped
“isoforms” having unclear significance.

There are many examples of isoenzymes in cell
metabolism that distinguish cells:

Adenylate kinase (AL in liver, and myokinase) – that
are distinguished by reactivity with sulfhydryl reagents
Pyruvate kinase
AMPK, and Calmodulin kinase
Malate, isocitrate, alcohol, and aldehyde dehydrogenase
Nitric oxide synthase (i, e, and n)…

References[edit]

Hunter, R. L. and C.L. Markert. (1957) Histochemical
demonstration of enzymes separated by zone electrophoresis
in starch gels. Science 125: 1294-1295

Uzunov, P. and Weiss, B.(1972) “Separation of multiple
molecular forms of cyclic adenosine 3′,5′-monophosphate
phosphodiesterase in rat cerebellum by polyacrylamide
gel electrophoresis.” Biochim. Biophys. Acta 284:220-226.

Uzunov, P., Shein, H.M. and Weiss, B.(1974) “Multiple
forms of cyclic 3′,5′-AMP phosphodiesterase
of rat cerebrum and cloned astrocytoma and
neuroblastoma cells.” Neuropharmacology 13:377-391.

Weiss, B., Fertel, R., Figlin, R. and Uzunov, P. (1974)
“Selective alteration of the activity of the multiple forms
of adenosine 3′,5′-monophosphate phosphodiesterase
of rat cerebrum.” Mol. Pharmacol.10:615-625.

Lactate dehydrogenase

In cells, the immediate energy sources involve glucose oxidation. In anaerobic metabolism, the donor of the phosphate group is adenosine triphosphate (ATP), and the reaction is catalyzed via the hexokinase or glucokinase: Glucose +ATP-Mg²+ = Glucose-6-phosphate (ΔGo = – 3.4 kcal/mol with hexokinase as the co-enzyme for the reaction.).
In the following step, the conversion of G-6-phosphate into F-1-6-bisphosphate is mediated by the enzyme phosphofructokinase with the co-factor ATP-Mg²+. This reaction has a large negative free energy difference and is irreversible under normal cellular conditions. In the second step of glycolysis, phosphoenolpyruvic acid in the presence of Mg²+ and K+ is transformed into pyruvic acid. In cancer cells or in the absence of oxygen, the transformation of pyruvic acid into lactic acid alters the process of glycolysis.
The energetic sum of anaerobic glycolysis is ΔGo = -34.64 kcal/mol. However a glucose molecule contains 686kcal/mol and, the energy difference (654.51 kcal) allows the potential for un-controlled reactions during carcinogenesis. The transfer of electrons from NADPH in each place of the conserved unit of energy transmits conformational exchanges in the mitochondrial ATPase. The reaction ADP³+ P²¯ + H²–à ATP + H2O is reversible. The terminal oxygen from ADP binds the P2¯ by forming an intermediate pentacovalent complex, resulting in the formation of ATP and H2O. This reaction requires Mg²+ and an ATP-synthetase, which is known as the H+-ATPase or the Fo-F1-ATPase complex. Intracellular calcium induces mitochondrial swelling and aging. [12].
The known marker of monitoring of treatment in cancer diseases, lactate dehydrogenase (LDH) is an enzyme that is localized to the cytosol of human cells and catalyzes the reversible reduction of pyruvate to lactate via using hydrogenated nicotinamide deaminase (NADH) as co-enzyme.
The causes of high LDH and high Mg levels in the serum include neoplastic states that promote the high production of intracellular LDH and the increased use of Mg²+ during molecular synthesis in processes pf carcinogenesis (Pyruvate acid>> LDH/NADH >>Lactate acid + NAD), [13].
LDH is released from tissues in patients with physiological or pathological conditions and is present in the serum as a tetramer that is composed of the two monomers LDH-A and LDH-B, which can be combined into 5 isoenzymes: LDH-1 (B4), LDH-2 (B3-A1), LDH-3 (B2-A2), LDH-4 (B1-A3) and LDH-5 (A4). The LDH-A gene is located on chromosome 11, whereas the LDH-B gene is located on chromosome 12. The monomers differ based on their sensitivity to allosteric modulators. They facilitate adaptive metabolism in various tissues. The LDH-4 isoform predominates in the myocardium, is inhibited by pyruvate and is guided by the anaerobic conversion to lactate.
Total LDH, which is derived from hemolytic processes, is used as a marker for monitoring the response to chemotherapy in patients with advanced neoplasm with or without metastasis. LDH levels in patients with malignant disease are increased as the result of high levels of the isoenzyme LDH-3 in patients with hematological malignant diseases and of the high level of the isoenzymes LDH-4 and LDH-5, which are increased in patients with other malignant diseases of tissues such as the liver, muscle, lungs, and conjunctive tissues. High concentrations of serum LDH damage the cell membrane [11, 31].

Relation between LDH and Mg as Factors of Interest in the Monitoring and Prognoses of Cancer

Aurelian Udristioiu, Emergency County Hospital Targu Jiu Romania, Clinical Laboratory Medical Analyses, E-mail: aurelianu2007@yahoo.com

Lactate Dehydrogenase (LDH) is ubiquitous in animals and
man, and it occurs in different organs of the body, each
region having a unique conformation of the subunits, but
the significance was once disputed. Perhaps the experiments
of Jakob and Monod on the lac 1 operon put to rest any
notions that isoenzymes and their conformational forms are
something of no real significance. This concept does not
necessarily apply in all cases of isoenzyme differences, by
which I mean that there may be a difference in reactivity at
the active site.

For that matter, Jakob and Monod discovered and elucidated
allosterism.

In biochemistry, allosteric regulation is the regulation of a
protein by binding an effector molecule at a site other than
the protein’s active site.

The site the effector binds to is termed the allosteric site.
Allosteric sites allow effectors to bind to the protein, often
resulting in a conformational change. Effectors that enhance
the protein’s activity are referred to as allosteric activators,
whereas those that decrease the protein’s activity are called
allosteric inhibitors.

Allosteric regulations are a natural example of control loops,
such as feedback from downstream products or feedforward
from upstream substrates. Long-range allostery is especially
important in cell signaling. Allosteric regulation
is also particularly important in the cell’s ability to adjust
enzyme activity.

The term allostery comes from the Greek allos (ἄλλος), “other,”
and stereos (στερεὀς), “solid (object).” This is in reference
to the fact that the regulatory site of an allosteric protein is
physically distinct from its active site.

Jacob and Monod model of transcriptional regulation of the lac operon by lac repressor

Jacob and Monod model of lac repressor

Most allosteric effects can be explained by the concerted
MWC model put forth by Monod, Wyman, and Changeux, [2]
or by the sequential model described by Koshland, Nemethy,
and Filmer.[3] Both postulate that enzyme subunits exist in
one of two conformations, tensed (T) or relaxed (R), and
that relaxed subunits bind substrate more readily than
those in the tense state. The two models differ most in
their assumptions about subunit interaction and the pre-
existence of both states.

Allosteric_Regulation Model

Monod, J. Wyman, J.P. Changeux. (1965). On the nature of
allosteric transitions:A plausible model. J. Mol. Biol.;12:88-118.
E. Jr Koshland, G. Némethy, D. Filmer (1966). Comparison of
experimental binding data and theoretical models in proteins
containing subunits. Biochemistry. Jan;5(1):365-8

The sequential model (2) of allosteric regulation holds that subunits
are not connected in such a way  that a  conformational change in
one induces a similar change in the others. Thus, all enzyme
subunits do not necessitate the  same conformation. Moreover,
the sequential model dictates that molecules of substrate
bind via an induced fit  protocol. In general, when a subunit
randomly collides with a molecule of substrate, the active site,
in essence, forms a  glove around its substrate.

While such an induced fit converts a subunit from the tensed
state to relaxed state, it does not propagate the conformational
change to adjacent subunits. Instead, substrate-binding at
one subunit only slightly alters the structure of other
subunits so that their binding sites are more receptive to
substrate.
To summarize:

subunits need not exist in the same conformation
molecules of substrate bind via induced-fit protocol
conformational changes are not propagated to all
subunits

The discovery of morpheeins has revealed a previously
unforeseen mechanism to target universally essential
enzymes for species-specific drug design and discovery.
A morpheein-based inhibitor would function by binding
to and stabilizing the inactive morpheein form of the
enzyme, thereby shifting the equilibrium to favor that form (3).

K. Jaffe, S.H. Lawrence (2008). “Expanding the
concepts in protein structure-function relationships
and enzyme kinetics: Teaching using morpheeins”.
Biochemistry and Molecular Biology Education36 (4)
: 274–283. http://dx.doi.org:/10.1002/bmb.20211.
PMC 2575429. PMID 19578473

Important related points are:

Non-regulatory allostery

A non-regulatory allosteric site refers to any non-regulatory
component of an enzyme (or any protein), that is not itself
an amino acid. For instance, many enzymes require sodium
binding to ensure proper function. However, the sodium
does not necessarily act as a regulatory subunit; the sodium
is always present and there are no known biological processes
to add/remove sodium to regulate enzyme activity. Non-
regulatory allostery could comprise any other ions besides
sodium (calcium, magnesium, zinc), as well as other chemicals
and possibly vitamins.

Lactate and malate dehydrogenases

LDH is a key enzyme in glycolysis. Anaerobic glycolysis is the
conversion of pyruvate into lactate acid in the absence
of oxygen. This pathway is important to glycolysis in two main
ways. The first is that

if pyruvate were to build up glycoysis
the generation of ATP would slow.

The second is anaerobic respiration

allows for the regeneration of NAD+ from NADH.

NAD+ is required when glyceraldehyde-3-phosphate
dehydrogenase oxidizes glyceraldehyde-3-phosphate in
glycolysis, which generates NADH. Lactate dehydrogenase
is responsible for the anaerobic conversion of NADH to
NAD+. Click here to see the residues which form
interactions with pyruvate in the Lactate Dehydrogenase
from Cryptosporidium parvum (2fm3). (Wikipedia)

Glycolysis ends with the synthesis of pyruvate.  But, to be
self-functioning, it must end with lactate.  Why?  Anaerobic
means “without oxygen”.  This is tantamount to saying
“without mitochondria”.

The mitochondria are especially adept at oxidizing
NADH to NAD+. NAD+ is needed to keep the glyceraldehyde-
3-PO4 dehydrogenase reaction functioning.
If glycolysis is to continue when no oxygen is present or in
short supply (as in a working muscle), an alternative means
of oxidizing NADH must occur.

Pyruvate has 2 metabolic fates:

it can either be converted into lactate or to acetyl-CoA .
Note that in animals and plants the electrons in  NADH
are transferred  to pyruvate which reduces the carbonyl
carbon in the pyruvate molecule to an alcohol. The
reaction is catalyzed by the enzyme lactate dehydrogenase.
Lactate (or L-lactate to be more precise)  is thus  a
“waste product”, since it has no metabolic fate other
than to be converted back into pyruvate in a reverse of
the  forward reaction.
More importantly, the NAD+ feeds back to the glyceraldehyde-
3-PO4 dehydrogenase reaction, which allows glycolysis
to continue. Were it not for lactate formation, glycolysis
as a self-functioning pathway could not exist.

In yeast a slightly different end of glycolysis becomes apparent.
Yeast do not synthesize lactate.  They do, however, oxidize
NADH back to NAD+ anaerobically.  How do they do this?  The
answer is they make ethanol.  In the reaction the pyruvate is
converted into acetaldehyde.  The reaction is catalyzed by a
lyase enzyme, pyruvate decarboxylase, which removes the
carboxyl group as a CO2.  Acetaldehyde is formed because
the electron pair that bonds the –COO group is not removed
by the decarboxylation.  A proton is plucked from the
environment giving the final product, acetaldehyde.
Acetaldehyde is now the substrate that will oxidize NADH to
NAD+ and in the process ethanol is formed.

There is another advantage to the pyruvate-lactate interchange.
The lactate formed by lactate dehydrogenase can be
reconverted. This allows a cell to synthesize glucose from lactate.
Converting lactate to glucose is a major feature of gluconeogenesis,
an anabolic pathway that synthesizes glucose from smaller
precursors such as lactate. This is important because acetyl-CoA
cannot be converted back to pyruvate and hence cannot be a
source of carbons for glucose biosynthesis.

ADP. ADP is required in the 3-phosphoglycerate kinase reaction
and in the pyruvate kinase reaction. It is formed from ATP in the
hexokinase reaction and the phosphofructokinase-I reaction.

NADH, ADP and PO4.   NADH oxidation is important in glycolysis.
NADH is converted into NAD+ in the mitochondria.  That
reaction is promoted by O2 ; NAD+ stays in the mitochondria.
Also in the mitochondria, ATP is formed by condensing ADP
with PO4.  Thus, O2 allows mitochondria to out-compete the
cytosol for ADP,  NADH and PO4, all limiting  substrates or
coenzymes.

In vertebrates, gluconeogenesis takes place mainly in the liver
and, to a lesser extent, in the cortex of kidneys. In many
animals, the process occurs during periods of fasting,
starvation, low-carbohydrate diets, or intense exercise.
The process is highly endergonic until it is coupled to the
hydrolysis of ATP or GTP, effectively making the process
exergonic. For example, the pathway leading from pyruvate
to glucose-6-phosphate requires 4 molecules of ATP and
2 molecules of GTP to proceed spontaneously. Gluco-
neogenesis is a target of therapy for type II diabetes,
such as metformin, which inhibits glucose formation
and stimulates glucose uptake by cells.

Lactate is formed at the endstage of glycolysis with insufficient
oxygen is transported to the liver where it is converted into
pyruvate by the Cori cycle using the enzyme lactate
dehydrogenase. In this reaction lactate loses two electrons
(becomes oxidized) and is converted to pyruvate. NAD+
gains two electrons (is reduced) and is converted to NADH.

Both lactate and NAD+ bind to the active site of the enzyme
lactate dehydrogenase and both lactate and NAD+ participate
in the catalysis reaction. In fact, catalysis could not occur
unless the coenzyme NAD+ bound to the active site.

lactat-pyr.LDH

http://academic.brooklyn.cuny.edu/biology/bio4fv/page/couple.gif

What is not shown:

The liver LDH is composed of predominantly M-type subunits.
The forward reaction is regulated in the H-type LDH, but not
the M-type enzyme by the formation of a ternary complex
of LDH-ox. NAD-lactate
The formation and breakup of the ternary complex is
dependent on the pyruvate in the forward reaction in a
concentration dependent manner.
The M-type LDH doesn’t have this tight binding of the LDH –
NAD+ – lactate (see catalysis below)
As lactate concentration builds in the circulation from heavy
muscle production (M-type), or from circulatory insufficiency,
the circulating lactic acid reaches the liver.
The lactic acid is taken up by the liver, and the high
concentration of lactic acid drives the backward reaction,
unrestricted.

Pyruvate, the first designated substrate of the gluconeogenic
pathway, can then be used to generate glucose. Transamination
or deamination of amino acids facilitates entering of their
carbon skeleton into the cycle directly  (as pyruvate or
oxaloacetate), or indirectly via the citric acid cycle.  It is
known that odd-chain fatty acids can be  oxidized to yield
propionyl-CoA, a precursor for succinyl-CoA, which can
be converted to  pyruvate and  enter  into gluconeogenesis.

gluconeogenesis

http://upload.wikimedia.org/wikipedia/commons/thumb/6/63/Amino_acid_catabolism.svg/300px-Amino_acid_catabolism.svg.png

Catalysis

Studies have shown that the reaction mechanism of LDH follows an ordered sequence.

mechanism of LDH reaction

In the forward reaction

NADH must bind to the enzyme Several residues are
involved in the binding of NADH. Once the NADH is
bound to the enzyme,
pyruvatebinds (substrate oxamate is shown; the CH3
group is replaced by NH2 to form oxamate). (see the
direction of the arrow)
binds to the enzyme between the nicotinamide ring
and several LDH residues.-
transfer of a hydride ion then happens quickly
in either direction giving a mixture of the two ternary
complexes,
enzyme-NAD+-lactate and enzyme-NADH-pyruvate .
finally L-lactate dissociates from the enzyme followed
by NAD+[2].

What is not shown is:

The dissocation of NAD+ and lactate from the H-type LDHs
is dependent on the pyruvate in the forward reaction in a
concentration dependent manner
This results in inhibition of the reaction as it proceeds as
a result of the abortive ternary complex that forms in about
500 msec carried out in the Aminco-Morrow stop flow analyzer.
The regulatory effect of the tighter binding of the LDH (H)-
NAD+-lactate is not seen with the M-type LDH.
The result of this is that the H-type LDH is regulated by the
formation of oxidized coenzyme bound with reduced substrate.

Genetics and Mutagenesis of Fish 1973, pp 243-276.
Developmental and Biochemical Genetics of Lactate
Dehydrogenase Isozymes in Fishes.
G. S. Whitt, E. T. Miller, J. B. Shaklee
http://link.springer.com/article/10.1007%2F978-3-642-
65700-9_23/lookinside/000.png

In the teleost there are only three of the isoenzymes. LDH-1,
3, and 5 (H4, H2M2, M4).

Lactic dehydrogenase isozymes in lens and cornea
Larry Bernstein, Michael Kerrigan, Harry Maisel
Experimental Eye Research Oct 1966; 5, (4): Pp 309–314, IN23–IN28
http://dx.doi.org:/10.1016/S0014-4835(66)80041-6

Lactic dehydrogenase isozymes of bovine and rabbit lens and
cornea were analyzed by starch gel electrophoresis.
Although there was a progressive loss of enzyme activity in
the lenses of both species with increasing age, the loss of
isozymes was more clearly evident in the bovine lens. In
the adult bovine lens,

lactic dehydrogenase isozyme Iwas predominant,
while in the adult rabbit lens, isozymes 3–5were mainly present.

The mobility of lens isozymes was identical to that of isozymes
in other tissues. Furthermore, the isozymes were not localized
to any major specific lens crystallin.

Lactate Dehydrogenase Isozyme Patterns of Human
Platelets and Bovine Lens Fibers
Elliot S. Vesell
Science 24 Dec 1965; 150(3704): pp.1735-1737
http://dx.doi.org:/10.1126/science

Since the platelets and lens fibers, like mature human erythrocytes,
lack a nucleus, the results strengthen the case for a

previously developed association between LDH-5 and the
cell nucleus.

These three cell types are mainly anaerobic, and therefore

their isozyme patterns are incompatible with the theory
that anaerobic ` tissues exhibit predominantly LDH-5
and aerobic tissues mainly LDH-1.

Lactate dehydrogenase isozymes and their relationship
to lens cell differentiation
James A. Stewart, John Papaconstantinou
Biochimica et Biophysica Acta (BBA) – General Subjects
26 May 1966; 121,(1): Pp 69–78
http://dx.doi.org:/10.1016/0304-4165(66)90349-7

Changes in the activity of lactate dehydrogenase (LDH) (l-lactate:
NAD+ oxidoreductase EC 1.1.1.27) isozymes are associated with
the growth and differentiation of bovine lens cells. Calf and adult
lens epithelial cells contain all 5 isozymes. The cathodal forms are
most active in the calf-epithelial cells; the anodal forms are most
active in the fiber cells. This transition from cathodal to anodal
forms of lactate dehydrogenase in the epithelial cells is associated
with cellular aging.

During the differentiation of an epithelial cell to a fiber cell, in calf
and adult lenses there is an enhancement of

the transition from cathodal forms to anodal forms.

The regulation of lactate dehydrogenase subunit synthesis may
be associated, therefore, with

the replicative activity of these cells.

In cells having the greatest replicative activity (calf epithelial
cells) the cathodal isozymes are most active; in cells having a
decreased mitotic activity (adult epithelial cells) the anodal
isozymes are most active. The non-replicative

fiber cell of calf and adult shows a transition toward the
anodal forms.

Although lens fiber cells have a low rate of oxidative metabolism
lactate dehydrogenase-I is the most active isozyme in these
cells. Kinetically,

lactate dehydrogenase-I factors other than, or in addition
to, the regulation of carbohydrate metabolism
are involved in regulating the synthesis of lactate dehydrogenase subunits.

Abbreviations LDH; lactate dehydrogenase

What is not examined to resolve the discrepancy (see the next item):

The Vessell paper was a challenge to the work in Nathan
Kaplan’s lab. However, there is sufficient complexity revealed
in these works that there is no conceptual foundation.

The analogy is to the loss of cell nuclei in crystallin lens
fiber formation with the LDH-H type subunits (aerobic?)
The findings are reproduced in several laboratories.
In the lens, glucose is catabolized primarily to lactic
acid, and is not appreciably combusted to CO2
(J Kinoshita. Glucose metabolism of Lens)
However, synthetic processes, including nuclear DNA and
cell replication requires TPNH. This is produced by means
of the Pentose Shunt.
The most favorable conditions for the lens are achieved
by incubating in a medium containing glucose in the
presence of oxygen. Under these conditions of
incubation (Kinoshita)

the lens remains completely transparent,
it maintains normal levels of high energy phosphate
bonds and cations, and
it shows a high rate of arginine incorporationinto protein.

incubation in the absence of glucose, but in the presence of oxygen

a haze is found in the lens,
a drop in high energy phosphate level is observed, and
Changes in cation levels are apparent.
A 50 percent decrease in the incorporation of arginine
into lens protein is also observed.

the most unfavorable condition for the lens is an anaerobic
incubation in a medium without glucose

Pirie2 observed that a-glycerophosphate is one of the end products
of lens metabolism. Its oxidation with DPN as the cofactor could
channel its electrons directly into the ETC to produce energy without
involving the Krebs cycle. a-Glycerophosphate is formed from intermediates of the glycolytic scheme by reduction of dihydroxy-
acetone phosphate, one of the triose phosphates produced in
glycolysis.

the dehydrogenase of the mitochondria catalyzes the transfer
of elections to form DPNH by the following reactions:

a-glycerophosphate + DPN+ ± dihydroxyacetone ……..

phosphate + DPNH.

The DPNH is channeled into the oxidative phosphorylation
mechanism to form ATP. The dihydroxyacetone phosphate
then diffuses out into the soluble cytoplasm, interacts with
the glycolytic intermediates by the reversal of the above reaction,

and the cyclic mechanism is begunover again.

That this type of electron transport system functions in the
lens was proposed by Pirie.
http://www.iovs.org/content/4/4/619.full.pdf

Lactate dehydrogenase activity and its isoenzymes in
concentric layers of adult bovine and calf lenses.
Sempol D, Osinaga E, Zigman S, Korc I, Korc B, Sans A, Radi R, et al.
Curr Eye Res. 1987 Apr;6(4):555-60.

The activity of lactate dehydrogenase (LDH) and its isoenzyme
pattern were studied in four concentric layers of adult
bovine and calf lenses. In both groups the specific activity of
the total LDH diminished progressively toward the internal
nuclear layer; the decrease was greater in the adult lenses.
The enzyme activities in the cortical layers of the calf lens
were lower than in the adult lens, but in the inner nuclear layers,
the opposite was found. All of the 5 LDH isoenzymes were found
in each layer. In both groups of animals the LDH1 isoenzyme
prevailed, followed by LDH2. No differences were found in the
percentage of each isoenzyme in the different lens layers.
The differences in the activitie(s) of LDH found may be due

to post-translational or post-synthetic modifications which
may occur during the aging process.

Structural basis for altered activity of M- and H-isozyme
forms of human lactate dehydrogenase.

Read JA1, Winter VJ, Eszes CM, Sessions RB, Brady RL.
Author information Proteins. 2001 May 1;43(2):175-85

Lactate dehydrogenase (LDH) interconverts pyruvate and
lactate with concomitant interconversion of NADH and NAD(+).
Although crystal structures of a variety of LDH have previously
been described, a notable absence has been any of the
three known human forms of this glycolytic enzyme. We have
now determined the crystal structures of two isoforms of
human LDH-the M form, predominantly found in muscle; and
the H form, found mainly in cardiac muscle. Both structures
have been crystallized as ternary complexes in the presence
of the NADH cofactor and oxamate, a substrate-like inhibitor.

Although each of these isoforms has different kinetic properties,
the domain structure, subunit association, and active-site regions
are indistinguishable between the two structures.

The pK(a) that governs the K(M) for pyruvate for the two isozymes
is found to differ by about 0.94 pH units, consistent with variation in
pK(a) of the active-site histidine.

The close similarity of these crystal structures suggests the distinctive
activity of these enzyme isoforms is likely to result

directly from variation of charged surface residues peripheral to the active site,
a hypothesis supported by electrostatic calculations based on each structure.

Proteins 2001;43:175-185.

Mechanistic aspects of biological redox reactions involving NADH.
Part 4. Possible mechanisms and corresponding intermediates for
the catalytic reaction in L-lactate dehydrogenase

J Molec Structure: THEOCHEM,25 Feb 1993; 279, Pp 99-125
Kathryn E. Norris, Jill E. Gready

The catalytic step in the conversion of pyruvate to L-lactate in the
enzyme L-lactate dehydrogenase involves the transfer of both a
proton and a hydride ion (A.R. Clarke, T. Atkinson and J.J. Holbrook,
TIBS, 14 (1989) 101.) However, it is not known whether the
reaction is concerted or, if a multistep process, the order in
which the transfers of the proton and the hydride ions take
place. Four possible non-concerted mechanisms can be
proposed, which differ in the order of the transfers of the
proton and hydride ion and the protonation state of the substrate
carboxylate group during the transfers. The energies and
optimized geometries of the corresponding intermediates,
protonated pyruvate, protonated pyruvic acid, deprotonated
L-lactate and deprotonated L-lactic acid, are computed using
the semiempirical AM 1 and ab initio SCF/3–21 G – methods.
These calculations are complementary to the study of
the substrates for the enzyme discussed in a previous paper
(K.E. Norris and J.E. Gready, J. Mol. Struct. (Theochem),
258 (1992) 109.) The structures and energetics of protonated
pyruvate and deprotonated L-lactate provide some
important insights into the requirements for enzymic reaction
and the characteristics of the transition state.

Pyruvate production by Enterococcus casseliflavus A-12
from gluconate in an alkaline medium

J Fermentation and Bioengineering, 1992; 73(4):287-291
H Yanase, N Mori, M Masuda, K Kita, M Shimao, N Kato

A newly isolated lactic acid bacterium, Enterococcus casseliflavus
A-12, produced pyruvic acid (16 g/l) during aerobic culture in
an alkaline medium containing sodium gluconate (50 g/l) as
the carbon source. The production was dependent on the pH
of the culture, the optimum initial pH being 10.0. With static
culture, the organism produced lactic acid (2.7 g/l) from both
gluconate and glucose. Pyruvate did not accumulate in growing
cultures on glucose, but resting cells obtained from a culture
on gluconate produced pyruvate from glucose as well as
gluconate. The enzyme profiles of the organism, which
grew on gluconate and glucose, suggested that gluconate
was metabolized via the Entner-Doudoroff and Embdem-
Meyerhof-Parnas pathways in aerobic culture, and that glucose
was oxidized mainly via the latter pathway under both aerobic
and anaerobic conditions. Gluconokinase, a key enzyme in
the aerobic metabolism of gluconate, was partially purified
from this strain and characterized.

A specific, highly active malate dehydrogenase by redesign
of a lactate dehydrogenase framework

HM Wilks, KW Hart, R Feeney, CR Dunn, H Muirhead, WN Chia, et al.

Department of Biochemistry, University of Bristol, United Kingdom.
Science 16 Dec1988: 242(4885), pp. 1541-1544
http://dx.doi.org:/10.1126/science.3201242

Three variations to the structure of the nicotinamide adenine
dinucleotide (NAD)-dependent L-lactate dehydrogenase
from Bacillus stearothermophilus were made to try to
change the substrate specificity from lactate to malate:
Asp197—-Asn, Thr246—-Gly, and Gln102—-Arg).

Each modification shifts the specificity from lactate to malate, although

only the last (Gln102—-Arg) provides an effective and
highly specific catalyst for the new substrate.

This synthetic enzyme has a ratio of catalytic rate (kcat) to
Michaelis constant (Km) for oxaloacetate of 4.2 x 10(6)M-1 s-1,

equal to that of native lactate dehydrogenase for its natural
substrate, pyruvate, and a maximum velocity (250 s-1),
which is double that reported for a natural malate from B.
stearothermophilus.

Malate dehydrogenase: distribution, function and properties.

Musrati RA1, Kollárová M, Mernik N, Mikulásová D.
Author information
Gen Physiol Biophys. 1998 Sep;17; (3):193-210.

Malate dehydrogenase (MDH) (EC 1.1.1.37) catalyzes the
conversion of oxaloacetate and malate. This reaction is
important in cellular metabolism, and it is coupled with
easily detectable cofactor oxidation/reduction. It is a
rather ubiquitous enzyme, for which several isoforms
have been identified, differing in their subcellular
localization and their specificity for the cofactor NAD
or NADP. The nucleotide binding characteristics can
be altered by a single amino acid change. Multiple
amino acid sequence alignments of MDH show there is a

low degree of primary structural similarity, apart from
several positions crucial for catalysis, cofactor binding
and the subunit interface.
Despite the low amino acids sequence identity their
3-dimensional structures are very similar.
MDH is a group of multimeric enzymes consisting of
identical subunits usually organized as either dimer
or tetramers with subunit molecular weights of 30-35 kDa.

Malate dehydrogenase, mitochondrial (MDH2)

UniProt Number:

P40926

Alternate Names:

Malate DH

Structure and Function:

Malate dehydrogenase (MDH2) is an enzyme in the citric
acid cycle that catalyzes the conversion of malate into
oxaloacetate (using NAD+) and vice versa (this is a
reversible reaction). Malate dehydrogenase is also
involved in gluconeogenesis, the synthesis of glucose
from smaller molecules.Pyruvate in the mitochondria is acted upon by pyruvate
carboxylase to form oxaloacetate, a citric acid cycle
intermediate.In order to get the oxaloacetate out of the mitochondria,
malate dehydrogenase reduces it to malate, and it then
traverses the inner mitochondrial membrane.Once in the cytosol, the malate is oxidized back to
oxaloacetate by cytosolic malate dehydrogenase.

Finally, phosphoenol-pyruvate carboxy kinase (PEPCK)
converts oxaloacetate to phosphoenol pyruvate.

Malate Dehydrogenase (MDH)(PDB entry 2x0i) is most known
for its role in the metabolic pathway of the tricarboxylic acid cycle,
critical to cellular respiration; The enzyme has other metabolic roles in –

glyoxylate bypass,
amino acid synthesis,
glucogenesis, and
oxidation/reduction balance .

An oxidoreductase, MDH has been extensively studied due to its
isozymes The enzyme exists in two places inside a cell:

the mitochondria and cytoplasm.
In the mitochondria, the enzyme catalyzes the reaction of
malate to oxaloacetate;
in the cytoplasm, the enzyme catalyzes oxaloacetate to
malate to allow transport.

The enzyme malate dehydrogenase is composed of either
a dimer or tetramer depending on the location of the enzyme
and the organism it is located in. During catalysis, the enzyme
subunits are

non-cooperative between active sites.

The mitochondrial MDH is complexly,

allosterically controlled by citrate, but no other known
metabolic regulation mechanisms have been discovered.
the exact mechanism of regulation has yet to be discovered.

Kinetically, the pH of optimization is 7.6 for oxaloacetate
conversion and 9.6 for malate conversion. The reported
K(m) value for malate conversion is 215 uM and the V(max)
value is 87.8 uM/min.

Comment:

The mMDH and the cMDH both form ternary complex
of MDH-NAD+-OAA formed during the forward reaction,
like the LDH H-type isozyme LDH-NAD+-PYR (mot the M-type).
However, the binding of the Enz-coenzyme-substrate is not
as strong as for the H-type LDH. .The regulatory role has
not been established.

References

↑Minarik P, Tomaskova N, Kollarova M, Antalik M. Malate
dehydrogenases–structure and function. Gen Physiol Biophys.
2002 Sep;21(3):257-65. PMID:12537350
↑Musrati RA, Kollarova M, Mernik N, Mikulasova D.
Malate dehydrogenase: distribution, function and properties.
Gen Physiol Biophys. 1998 Sep;17(3):193-210. PMID:9834842
↑Boernke WE, Millard CS, Stevens PW, Kakar SN, Stevens FJ,
Donnelly MI. Stringency of substrate specificity of
Escherichia coli malate dehydrogenase. Arch Biochem
Biophys. 1995 Sep 10;322(1):43-52. PMID:7574693
doi:http://dx.doi.org/10.1006/abbi.1995.1434
↑Goward CR, Nicholls DJ. Malate dehydrogenase: a model
for structure, evolution, and catalysis. Protein Sci. 1994
Oct;3(10):1883-8. PMID:7849603
doi:http://dx.doi.org/10.1002/pro.5560031027

Kinetic determination of malate dehydrogenase isozymes.

L H Bernstein, M B Grisham

Journal of Molecular and Cellular Cardiology (Impact Factor: 5.15).
11/1978; 10(10):931-44. http://dx.doi.org/10.1016/0022-2828(78)90339-5

Source: PubMed

ABSTRACT These studies determine the levels of malate
dehydrogenase isoenzymes in cardiac muscle by a steady
state kinetic method which depends on the differential inhibition
of these isoenzyme forms by high concentrations of oxaloacetate.
This inhibition is similar to that exhibited by lactate dehydrogenase
in the presence of high concentrations of pyruvate. The results
obtained by this method are comparable in resolution to those
obtained by CM-Sephadex fractionation and by differential
centrifugation for the analyses of mitochondrial malate
dehydrogenase and cytoplasmic malate dehydrogenase in
tissues. The use of standard curves of percent inhibition of
malate dehydrogenase activity plotted against the ratio of
mitochondrial MDH activity to the total of mMDH and cMDH
activities [ malate dehydrogenase ratio] (percent m-type) is
introduced for studies of comparative mitochondrial
function in heart muscle of different species or in different
tissues of the same species.

Calmodulin and Protein Kinase C Increase Ca21-stimulated
Secretion by Modulating Membrane-attached Exocytic Machinery

YA Chen, V Duvvuri, H Schulmani, and RH Scheller
Hughes Medical Institute, Department of Molecular and Cellular
Physiology, and the iDepartment of Neurobiology, Stanford
University School of Medicine, Stanford, California 94305-5135
JBC Sep 10, 1999; 274( 37): 26469–26476

Using a reconstituted [3H]norepinephrine
release assay in permeabilized PC12 cells, we
found that essential proteins that support the triggering
stage of Ca21-stimulated exocytosis are enriched in an
EGTA extract of brain membranes. Fractionation of this
extract allowed purification of two factors that stimulate
secretion in the absence of any other cytosolic proteins.
These are calmodulin and protein kinase Ca
(PKCa). Their effects on secretion were confirmed using
commercial and recombinant proteins. Calmodulin enhances
secretion in the absence of ATP, whereas PKC
requires ATP to increase secretion, suggesting that
phosphorylation is involved in PKC- but not calmodulin
mediated stimulation. Both proteins modulate release
events that occur in the triggering stage of exocytosis.

Endothelial nitric oxide synthase (eNOS) variants in
cardiovascular disease: pharmacogenomic implications

Indian J Med Res May 2011; 133: 464-466

Commentary

Manjula Bhanoori

Department of Biochemistry, University College of Science,
Osmania University, Hyderabad 500 007, India

The maintenance of regular vascular tone substantially
depends on the bioavailability of endothelium-derived
nitric oxide (NO) synthesized by eNOS. The essential
role of NO, as the elusive endothelium-derived relaxing
factor (EDRF), was the topic of research that won the
1998 Nobel Prize in Physiology or Medicine. The eNOS
gene, as a candidate gene in the investigations on
hypertension genetics, has attracted the attention of
several researchers because of the established role
of NO in vascular homeostasis. The eNOS variants
located in the 7q35-q36 region have been investigated
for their association with CVD, particularly hypertension.
Three variants, viz., (i) G894T substitution in exon 7
resulting in a Glu to Asp substitution at codon 298 (rs1799983),
(ii) an insertion-deletion in intron 4 (4a/b) consisting of two
alleles (the a*-deletion which has four tandem 27-bp repeats
and the b*-insertion having five repeats), and (iii) a T786C
substitution in the promoter region (rs2070744), have been
extensively studied20-22. Individual SNPs often cause only
a modest change in the resulting gene expression or function.
It is, therefore, the concurrent presence of a number of SNPs
or haplotypes within a defined region of the chromosome that
determines susceptibility to disease development and progression,
particularly in case of polygenic diseases.

Shankarishan et al24 analysed for the first time the prevalence
of eNOS exon 7 Glu298Asp polymorphism in tea garden community
of North Eastern India, who are a high risk group for CVD. This study
also included indigenous Assamese population and found no
significant difference between the distribution patterns of eNOS
exon 7 Glu298Asp variants between the communities. They have
rightly mentioned that for developing public health policies and
programmes it is necessary to know the prevalence and distribution
of the candidate genes in the population, as well as trends in
different population groups. They have also observed that the
eNOS exon 7 homozygous GG wild genotype (75.8%) was
predominant in the study population followed by heterozygous
GT genotype (21.5%) and homozygous TT genotype (2.7%).
The frequency distribution of the homozygous GG, heterozygous
GT and homozygous mutant TT genotypes were comparable to
that of the north Indian and south Indian population.

Polymorphisms in the endothelial nitric oxide synthase gene have
been associated inconsistently with cardiovascular diseases.
Varying distribution of eNOS variants among ethnic groups may
explain inter-ethnic differences in nitric oxide mediated vasodilation
and response to drugs28. Different population studies showed
association of eNOS polymorphisms with variations in NO
formation and response to drugs. Cardiovascular drugs including
statins increase eNOS expression and upregulate NO formation
and this effect may be responsible for protective, pleiotropic
effects produced by statins31. With respect to hypertension,
studies have reported interactions between diuretics and
polymorphisms in eNOS gene. Particularly, the Glu298Asp
polymorphism made a statistically significant contribution to
predicting blood pressure response to diuretics.

Neuronal Nitric Oxide Synthase and Its Interaction
With Soluble Guanylate Cyclase Is a Key Factor for
the Vascular Dysfunction of Experimental Sepsis

GM. Nardi, K Scheschowitsch, D Ammar, SK de
Oliveira, TB. Arruda; J Assreuy

Vascular dysfunction plays a central role in sepsis, and it is
characterized by hypotension and hyporesponsiveness to
vasoconstrictors. Nitric oxide is regarded as a central element
of sepsis vascular dysfunction. The high amounts of nitric
oxide produced during sepsis are mainly derived from the
inducible isoform of nitric oxide synthase 2.
We have previously shown that nitric oxide synthase 2 levels
decrease in later stages of sepsis, whereas levels and activity
of soluble guanylate cyclase increase. Therefore, we studied
the putative role of other relevant nitric oxide sources, namely,

the neuronal (nitric oxide synthase 1) isoform, in sepsis
and its relationship with soluble guanylate cyclase.

We also studied the consequences of

nitric oxide synthase 1 blockade in the hyporesponsiveness
to vasoconstrictors.

1) Both nitric oxide synthase 1 and soluble guanylate cyclase
are expressed in higher levels in vascular tissues during sepsis;

2) both proteins physically interact and nitric oxide synthase 1
blockade inhibits cyclic guanosine monophosphate production;

3) pharmacological blockade of nitric oxide synthase 1 using
7-nitroindazole or S-methyl-l-thiocitrulline reverts the hypo
responsiveness to phenylephrine and increases the vaso
constrictor effect of norepinephrine and phenylephrine.

Sepsis induces increased expression and physical association
of nitric oxide synthase 1/soluble guanylate cyclase and a higher
production of cyclic guanosine monophosphate that together
may help explain sepsis-induced vascular dysfunction.

In addition, selective inhibition of nitric oxide synthase 1
restores the responsiveness to vasoconstrictors.

Therefore, inhibition of nitric oxide synthase 1 (and possibly
soluble guanylate cyclase) may represent a valuable
alternative to restore the effectiveness of vasopressor
agents during late sepsis. (Crit Care Med 2014; XX:00–00)

Nitric Oxide Synthase Inhibitors That Interact with Both Heme
Propionate and Tetrahydrobiopterin Show High Isoform Selectivity

S Kang, W Tang, H Li, G Chreifi, P Martásek, LJ. Roman,
TL. Poulos, and RB. Silverman

†Department of Chemistry, Department of Molecular Biosciences,
Chemistry of Life Processes Institute, Center for Molecular Innovation
and Drug Discovery, Northwestern University, Evanston, Illinois
‡Departments of Molecular Biology and Biochemistry, Pharmaceutical
Sciences, and Chemistry, University of California, Irvine, California,
Department of Biochemistry, University of Texas Health Science Center,
San Antonio, Texas

Overproduction of NO by nNOS is implicated in the pathogenesis of
diverse neuronal disorders. Since NO signaling is involved in
diverse physiological functions, selective inhibition of nNOS
over other isoforms is essential to minimize side effects. A series of
α-amino functionalized aminopyridine derivatives (3−8) were
designed to probe the structure−activity relationship between ligand,
heme propionate, and H4B. Compound 8R was identified as the
most potent and selective molecule of this study, exhibiting a Ki of
24 nM for nNOS, with 273-fold and 2822-fold selectivity against
iNOS and eNOS, respectively.Although crystal structures of 8R
complexed with nNOS and eNOS revealed a similar binding mode,
the selectivity stems from the distinct electrostatic environments in
two isoforms that result in much lower inhibitor binding free energy
in nNOS than in eNOS. These findings provide a basis for further
development of simple, but even more selective and potent, nNOS
inhibitors

Aurelian
Aurelian Udristioiu

Lab Director at Emergency County Hospital Targu Jiu

In cells, the immediate energy sources involve glucose oxidation. In anaerobic metabolism, the donor of the phosphate group is adenosine triphosphate (ATP), and the reaction is catalyzed via the hexokinase or glucokinase: Glucose +ATP-Mg²+ = Glucose-6-phosphate (ΔGo = – 3.4 kcal/mol with hexokinase as the co-enzyme for the reaction.).
In the following step, the conversion of G-6-phosphate into F-1-6-bisphosphate is mediated by the enzyme phosphofructokinase with the co-factor ATP-Mg²+. This reaction has a large negative free energy difference and is irreversible under normal cellular conditions. In the second step of glycolysis, phosphoenolpyruvic acid in the presence of Mg²+ and K+ is transformed into pyruvic acid. In cancer cells or in the absence of oxygen, the transformation of pyruvic acid into lactic acid alters the process of glycolysis.
The energetic sum of anaerobic glycolysis is ΔGo = -34.64 kcal/mol. However a glucose molecule contains 686kcal/mol and, the energy difference (654.51 kcal) allows the potential for un-controlled reactions during carcinogenesis. The transfer of electrons from NADPH in each place of the conserved unit of energy transmits conformational exchanges in the mitochondrial ATPase. The reaction ADP³+ P²¯ + H²–à ATP + H2O is reversible. The terminal oxygen from ADP binds the P2¯ by forming an intermediate pentacovalent complex, resulting in the formation of ATP and H2O. This reaction requires Mg²+ and an ATP-synthetase, which is known as the H+-ATPase or the Fo-F1-ATPase complex. Intracellular calcium induces mitochondrial swelling and aging. [12].
The known marker of monitoring of treatment in cancer diseases, lactate dehydrogenase (LDH) is an enzyme that is localized to the cytosol of human cells and catalyzes the reversible reduction of pyruvate to lactate via using hydrogenated nicotinamide deaminase (NADH) as co-enzyme.
The causes of high LDH and high Mg levels in the serum include neoplastic states that promote the high production of intracellular LDH and the increased use of Mg²+ during molecular synthesis in processes pf carcinogenesis (Pyruvate acid>> LDH/NADH >>Lactate acid + NAD), [13].
LDH is released from tissues in patients with physiological or pathological conditions and is present in the serum as a tetramer that is composed of the two monomers LDH-A and LDH-B, which can be combined into 5 isoenzymes: LDH-1 (B4), LDH-2 (B3-A1), LDH-3 (B2-A2), LDH-4 (B1-A3) and LDH-5 (A4). The LDH-A gene is located on chromosome 11, whereas the LDH-B gene is located on chromosome 12. The monomers differ based on their sensitivity to allosteric modulators. They facilitate adaptive metabolism in various tissues. The LDH-4 isoform predominates in the myocardium, is inhibited by pyruvate and is guided by the anaerobic conversion to lactate.
Total LDH, which is derived from hemolytic processes, is used as a marker for monitoring the response to chemotherapy in patients with advanced neoplasm with or without metastasis. LDH levels in patients with malignant disease are increased as the result of high levels of the isoenzyme LDH-3 in patients with hematological malignant diseases and of the high level of the isoenzymes LDH-4 and LDH-5, which are increased in patients with other malignant diseases of tissues such as the liver, muscle, lungs, and conjunctive tissues. High concentrations of serum LDH damage the cell membrane [11, 31].

Relation between LDH and Mg as Factors of Interest in the Monitoring and Prognoses of Cancer

Aurelian Udristioiu, Emergency County Hospital Targu Jiu Romania, Clinical Laboratory Medical Analyses, E-mail: aurelianu2007@yahoo.com
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Larry Bernstein

CEO/CSO at Triplex Consulting

The inhibition be pyruvate is related by a ternary complex formed by NAD+ formed in the catalytic forward reaction Pyruvate + NADH –> Lactate + NAD(+). The reaction can be followed in an Aminco-Morrow stop-flow analyzer and occurs in ~ 500 msec. The reaction does not occur with the muscle type LDH, and it is regulatory in function. I did not know about the role of intracellular Mg(2+) in the catalysis, as my own work was in Nate Kaplan’s lab in 1970-73.

This difference in the behavior of the isoenzyme types was considered to be important then in elucidating functional roles, but it was challenged by Vessell earlier. The isoenzymes were first described by Clement Markert at Yale. I think, but don’t know, that the Mg++ would have a role in driving the forward reaction, but I can’t conceptualize how it might have any role in the difference between muscle and heart.

I didn’t quite know why oncologists used it specifically. Cancer cells exhibit the reliance on the anaerobic (muscle) type enzyme, which is also typical of liver, but with respect to the adenylate kinases – the liver AK and muscle AK (myokinase) are different. That difference was discovered by Masahiro Chiga, and differences in the reaction with sulfhydryl reagents were identified by Percy Russell.

Oddly enough, Vessell had a point. The RBC has the heart type predominance, not the M-type. He thought that it was related to the loss of nuclei from the reticulocyte. I did not buy that, and I had worked on the lens of the eye at the time.
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Aurelian
Aurelian Udristioiu

Lab Director at Emergency County Hospital Targu Jiu

Very interesting scientific comments. Thanks. !
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Aurelian

Aurelian Udristioiu

Lab Director at Emergency County Hospital Targu Jiu

The IDH1 and IDH2 genes are mutated in > 75% of different malignant diseases. Two distinct alterations are caused by tumor-derived mutations in IDH1 or IDH2,
IDH1 and IDH2 mutations have been observed in myeloid malignancies, including de novo and secondary AML (15%–30%), and in pre-leukemic clone malignancies, including myelodysplastic syndrome and myeloproliferative neoplasm (85% of the chronic phase and 20% of transformed cases in acute leukemia.
Aurelian Udristioiu, M.D
City Targu Jiu, Romania
AACC, NACB, Member, USA.