INTRODUCTION
INTRODUCTION
Metabolism is the foundation of all living systems and is involved in all the chemical and physical changes that occur in a cell. It ensures normal cellular functioning and enables cells to respond to external and internal stimuli. The breakdown of complex organic constituents with the concomitant liberation of energy and the synthesis of biomolecules constitute an important function of cellular metabolism (Figure 1).
Figure 1: An Overview of metabolism
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While the metabolism of carbohydrates is pivotal in the generation of energy, the conversion of glutamine in nucleic acids plays a crucial role in cellular growth (Figure 2). Hence, in order to understand the metabolic status of a cell or a living system, it is essential to evaluate the cellular metabolic profile. The metabolic profile provides a detailed insight into the molecular working of a cell at a given moment. The quantitative and the qualitative status of the metabolites is the fingerprint of any cellular function. Metabolomics, the study of all the metabolites present in a cell at a specific moment provides a snapshot of the molecular machinery operative under a given condition and allows a better understanding of the macromolecules mediating the production of these metabolites (Stitt, M., 2003).
Figure 2: The central role of the TCA cycle in cellular metabolism (Adapted from Voet, 1990)
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Metabolomics
During the last decade, the genome of many organisms has been solved. However, the functions of many genes still remain obscure. Although the genome provides the genetic information a cell possesses, how this message is translated into biological action is still elusive (Griffin, J., 2003). Furthermore, a phenotype is rarely ever an exact replica of a genotype as the transmission of the genetic information may undergo a variety of regulatory controls that may not be dependent on the initial genetic message (Stitt, M., 2003). For instance, silent phenotypes arise due to the ability of organisms to adapt and utilize various metabolic routes without the direct dependence on genetic manipulation (Griffin, J., 2003). The basis of metabolomics that is the global detection and comparison of metabolites found in an organ, a tissue, a cell or a cell compartment enables the elucidation of a given phenotype i.e. the expressed information in a system (Allen, J., 2003). Thus, the metabolome, which is the set of metabolites synthesized, depends on the genotype of the cell and the environment of the cell (Brindle, K 2003). It allows us to decipher allelic changes, to observe the consequences of transcriptional, post-transcriptional, translational and post-translational manifestations and to evaluate metabolic fluxes (Stitt, M. 2003). These factors contribute to a cellular phenotype i.e. the cell we see. Metabolomics, the multilevel analysis of the concerted actions of a system can also simultaneously pinpoint the main regulatory processes involved. The identification of these key regulatory forks informs us on the metabolic fluxes/alterations. Metabolomics can also solve situations where one genetic variation affects more then one biochemical pathway (Griffin, J., 2003). The measurement of metabolites in a biological system reveals the response of an organism to changes in its environment, and defines the important changes a cell may be experiencing. Hence, the identification of changes in key metabolites can be correlated to changes in the phenotype (Figure 3). Thus, the specific metabolites present in the blood plasma of patients with coronary artery disease become markers for this disease and is the fingerprint of a certain phenotype.
Pathology, genetic intervention and drug toxicity elicit a multitude of responses leading to an altered metabolic network and subsequently, to an altered phenotype. A metabolic network can be subdivided into monofunctional units. Some units are more important then others in linking the metabolic network (Ravasz, E., 2002). For example, glycolysis and oxidative phosphorylation constitute two separate units integrated in a metabolic network. A small perturbation in one unit can be perpetuated in the entire metabolic networks with or without affecting the phenotype (Stitt, M., 2003). This is why metabolic fingerprinting is crucial if the genomic significance is to be of any value. Thus, the metabolic differences or changes may help us predict the function of an affected (deleted or upregulated) gene. In some systems, it is possible to observe the carbon and nitrogen metabolism arising from the metabolism of glucose, fructose or sucrose. Metabolomics also allows us to determine reaction pathways by examining metabolite concentrations at a specific time interval. Over time, the closer metabolites are to each other, the more likely they are connected by a single reaction (Brindle, K. 2003). Hence, metabolomics is a powerful tool that will enable the assignment of genetic functions.
Metabolite fingerprint can help elucidate the induction of key metabolites that are necessary for the survival and/or adaptation of a particular organism submitted to an extreme condition (Sauer, U., 2004). It will be also possible to examine metabolite levels before and after the application of pesticides on plants in order to determine the biochemical repercussions of these pesticides. In therapy, metabolomic studies may help in the manipulation of particular metabolic fluxes that may be upregulated, non functional or affected in disease states. The mechanisms involved in pathogen-host interactions can be readily evaluated by metabolomic studies (Fiehn, O. 2002). The possibility of modifying or altering the metabolic fluxes of an organism in order to obtain metabolites with pharmacological significance and the discovery of novel biochemical pathways and cellular networks are other areas where the role of metabolomics will be critical.
Figure 3: Strategy to study biochemical adaptation / toxicology
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Although metabolomics offers great promise, the identification of thousands of metabolites is not an easy task. Analyzing all the metabolites in a biological system at a given time is indeed a daunting proposition (Stitt, M., 2003). Furthermore, the reactivity and the half-life of these metabolites render this exercise challenging. The evolution of technologies is therefore critical if the identities of all the metabolites are to be elucidated. Nuclear magnetic resonance (NMR) is an inexpensive and non-invasive technique that gives a snapshot of the metabolic processes occurring at a defined state or level. This metabolic fingerprinting minimizes false positives and detects biochemical variations caused by a disorder, stress or disease. 1H NMR is utilized to link the response of metabolism or the phenotype to different mRNA and protein expression. Atoms such as 31P and 13C that can also be visualized by NMR extend the amount of metabolites that can be detected (Griffin, J., 2003). Other techniques involved in metabolic profiling include mass spectroscopy (MS) and high performance liquid chromatography (HPLC). MS has the advantage of being more sensitive to the metabolites present in low concentrations. Therefore, MS can identify a greater amount of metabolites. The study of metabolomics may also rely on the identification and separation of metabolites accomplished by the combined action of gas chromatography (GC) and liquid chromatography (LC) with MS. The limiting factor in metabolomics is the need to increase the number of different metabolites that are quantifiable in a given system (Griffin, J., 2003). These metabolites are difficult to capture due to their dynamic and chemical behavior. However, the use of superconducting probes (cryoprobes) will help increase the sensitivity towards unlabelled carbon atom. These superconducting probes will reduce thermal noise. Many diseases arise due to metabolic arrest or alteration. Often, the initial event that triggers the onset of the disease is difficult to pinpoint. In neurodegenerative diseases, oxidative stresses as well as metabolic abnormalities lead to the progression of neurodegeneration. However, the order by which these events occur is still unknown and metabolomic studies may likely solve this mystery. Hence, metabolomics is an essential tool if one is to delineate the molecular details of cellular functions and how an organism operates under stress. A stress is an abnormal situation a cell is compelled to face and may be chemical, physical or biological in origin (Appanna, V., 1999).
Oxidative stress and diseases
Oxidative stress is a major stress that all aerobic organisms have to deal with. In fact, most neurodegenerative disorders are a consequence of a defective oxidative energy metabolism. It is known that oxidative stress constitutes one of the earliest events in neurodegeneration (Brown, A. 2000). Are reactive oxygen species (ROS) and a defective metabolism caused by one another or are they consequences of a more proximate event? In the brain, metabolic disease and ROS lead to the same aberrant phenotype. However, oxidative conditions are not restricted to the brain area and are encountered throughout biological systems (Figure 4).
Figure 4: Physiological conditions that trigger the generation of ROS
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In aerobic respiration, the electron transport chain is susceptible to electron leakages. These free electrons reduce nearly 2 % of the oxygen consumed and give rise to reactive oxygen intermediates such as O2-∙, H2O2, HOCl, OH· at the level of complex I and III (Figure 5) (Nulton-Persson, A., 2001). These are referred to as ROS. The properties of these oxidative intermediates have been exploited in the manufacturing of disinfectants and antibacterial agents (Elkins, J. 1999) and also as a cytotoxic tool during phagocytosis (Gonzalez-Flecha, B., 1995). Most ROS arise from the mitochondrial electron transport system. Consequently, mitochondria are also a major target of the deleterious events created by ROS (Tretter, L., 2000). Oxidative stress is thought to disrupt the mitochondrial membranes by modifying crucial thiol groups allowing the opening of the membrane permeability transition (MPT) pores and the subsequent apoptotic cascade. Another manifestation of oxidative stress is the peroxidation of membranes which involves the formation of highly reactive aldehydic products (Humphries, K. 1998). The latter deleteriously attacks proteins and propagates the oxidative condition. H2O2 has a relatively long half-life compared to other ROS like O2-∙, OH·.
Figure 5: ROS generated by mitochondrial processes
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H2O2 may also arise naturally due to the presence of numerous oxidases such as xanthine, monoamine, NADPH and urate oxidase, cyclooxygenase, and nitric oxide synthase (Zhang, Z. 2000). In biological systems, free redox active metal ions such as Fe+3 and Cu+2 are markers of oxidative stress. These metal ions mediate the formation of the highly toxic and ravaging hydroxyl radical (∙OH). It is impossible to completely eliminate ROS from the metabolism since they are a product of respiration. Furthermore, deliberate production is an important part of cellular activity as ROS have been shown to act as secondary messengers, gene regulators and mediate cell activation (Polla, A. 2003).
Nitric oxide (NO), is another potent oxidizing molecule and it functions both as a scavenger of superoxide and as a precursor of peroxynitrite (PN). ROS and reactive nitrogen species (RNS) are the major oxidants formed in the cell. NO-linked redox modifications of proteins include protein nitrosation and tyrosine nitration that may cause severe damage to the cell. Nevertheless, recently, the endogenous nitrosylation of proteins has been shown to be critical in the regulation of cellular functions (Shopfer, F. 2003).
Peroxide in biological systems
Peroxide plays a significant role in oxidative damage (Lord-Fontaine, S. 2002). Metabolically active organs such as the liver and the kidneys produce huge amounts of peroxide (Nath, K. 1995). On the other hand, the central nervous system is the most susceptible to oxidative stress as it is characterized with low catalase activity and relatively high amounts of polyunsaturated lipids which are easily oxidized. The oxidation of dopamine in the brain also perpetuates the damage to this organ (Carri, M. 2003).
In situation of H2O2 or O2-· stress, aconitase (ACN) (EC 4.2.1.3), α –ketoglutarate dehydrogenase (α-KGDH) (EC 1.2.4.2) and pyruvate dehydrogenase (PDH) (EC 1.2.4.1) are main enzymatic targets. ACN has been shown to interact directly with peroxide in vitro. The reactivation of the enzyme is achieved by the addition of free iron. This scenario may likely occur in vivo (Nulton-Persson, A., 2001). Lipoamide, an essential cofactor for the E2 subunit constituting α-KGDH and PDH, is a key target. This cofactor consists of the covalent assemblage of lipoic acid to a lysine residue of E2. The reduced form, dihydrolipoamide, is modified following H2O2 or O2-· exposure in both yeast and E-Coli (Figure 6).
Figure 6: Different subunits involved in α-keto acid dehydrogenases (Taken from Voet, 1990)
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The oxidation of these enzymes creates carbonyl groups that can be easily probed using anti 2, 4-Dinitrophenyl Hydrazone antibodies (Cabiscol, E. 2000). Therefore it is not surprising that H2O2 also causes the depolarization of membranes due to the inhibition of α-KGDH and the reduction of NADH synthesis by this enzyme. H2O2 is also known to cause the impairment of calcium utilization and the reduction of ATP synthesis (Chinopoulos, C. 1999). Another scenario implicates the direct inhibition of α-KGDH by H2O2 due to its interactions with SH groups in the E2 subunit of this enzyme. Other enzymes such as glyceraldehyde-3-phosphate dehydrogenase (GAPDH) and Ca+2 ATPases may be similarly impeded.
Due to its ability to modify and affect the activity of proteins, H2O2 is believed to be a signaling molecule for proteins like phosphatases. Peroxide can alter cellular functions by reversible oxidation of key sulfhydryl residues. Peroxide satisfies the requirement of a signaling molecule for its production and neutralization respond to physiological stimuli and is normally tightly regulated by multiple enzymatic and non enzymatic scavengers. Peroxide may therefore be an important regulator of mitochondrial functions. When rat mitochondria were treated with H2O2, SDH, α-KGDH and ACN declined in activity (Nulton-Persson, A. 2001). It was determined that the redox status of the mitochondria impacted the activity of SDH and α-KGDH and that the inhibition of these enzymes was not due to direct interaction with peroxide. In response to oxidative species such as H2O2, the organelle limits the generation of reducing factors (FADH2 and NADH) via SDH and α-KGDH and thereby limits the generation of H2O2 or O2-·. Thus, the inhibition of these enzymes may serve as an antioxidative mechanism preventing the generation of additional ROS. However, it is not clear how these enzymes are deactivated/reactivated. Most likely though, H2O2 does not directly interact with these enzymes. One possible mechanism is thought to involve the reversible glutathionylation of these enzymes since glutathione is also a redox responsive molecule (Nulton-Persson, A. 2001). In this manner, H2O2 is a signaling molecule and the TCA cycle may serve as an antioxidative system geared towards the prevention of ROS generation. The detoxification of H2O2 is mainly assumed by catalase and glutathione peroxidase, two molecules that require NADPH, a crucial dinucleotide in the metabolism of ROS (Lord-Fontaine, S., 2002)
Antioxidative defense mechanisms
ROS are critical in some cellular processes. However, when the prooxidant and antioxidant balance is disturbed, serious injuries are manifested. Therefore, organisms are stocked with many antioxidant defense systems. Some are non-enzymatic in nature and include bilirubin, albumin, flavanoids, GSH, ascorbic acid, α-tocophenol, β-carotene and uric acids (Prior, R.L. 1999). Evidence suggests that the treatment of Alzheimer’s disease patients with antioxidants such as vitamin E delay the deterioration of cognitive functions. Superoxide dismutase (SOD), catalase and GSH dependent peroxidase are enzymes that play a pivotal role in ROS defense. Apart from these scavengers, other mechanisms exist in order to reduce the levels of intracellular ROS (Elkins, J.G. 1999). Other protective mechanisms include metal transport systems and the induction of transcription factors (Figure 7). The protection against oxidants is elaborate. First, it is important for cells to prevent oxidative damage at the source by preventing electron leakage. Second the dangerous ROS can be intercepted before they can be deleterious by antioxidant molecules. A final checkpoint involves the repair of the damage caused by these ROS (Carrie, M. 2003). Toxicants can affect the free radical scavengers as well as the DNA repair mechanisms or toxicants can affect the production of free radicals. When this equilibrium is broken several biochemical consequences arise such as damage to the DNA, the depletion of GSH pools, the peroxidation of lipids, the alteration of metabolism and apoptosis or necrosis (Panaretakis, T. 2001). Other enzymes are also implicated albeit indirectly in maintaining the cellular redox balance such as glucose 6-phosphate dehydrogenase (G6PDH), malic enzyme (ME) and isocitrate dehydrogenase (ICDH).
Figure 7: Oxidative and antioxidative systems (Taken from Carri, M., 2003)
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Glucose-6-Phosphate Dehydrogenase (G6PDH)
The enzyme G6PDH is utilized in the catabolism of glucose. In diabetes mellitus, the level of extracellular glucose is elevated. The resulting hyperglycemic condition is accompanied by an elevation in oxidative damage. This elevation is thought to be due to the decreased activity of G6PDH leading to decreased level of NADPH. This enzyme is critical in the production of NADPH and is the only NADPH producing enzyme in erythrocytes (Salvemini, F. 1999). Other enzymes such as ICDH-NADP+ and ME also contribute to the NADPH pool. The importance of G6PDH is also demonstrated by mutants that cannot tolerate the reactive oxygen intermediate superoxide which is converted to peroxide in biological systems. This is explained by the lack of activity of glutathione reductase caused by the absence of NADPH (Figure 8).
Figure 8: Dependence of NADPH for the antioxidative system (Taken from Ben-Yoseph, O., 1996)
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The enzyme G6PDH is also critical for cell growth. It is believed that when peroxide levels become too high to allow cell growth, G6PDH produces the required amounts of NADPH to rectify the imbalance of oxidants and the cells can grow (Tian W-N., 1998). This enzyme is also reported to be the most sensitive to oxidative stress in many biological organisms. Low levels of NADPH also impede the tetrameric and active formation of catalase (Salvemini, F. 1999). The importance of G6PDH in the cellular antioxidant system is undeniable.
Isocitrate dehydrogenase (ICDH-NADP+)
Another enzyme, the soluble ICDH-NADP+ is becoming an intriguing enzyme because of its location throughout the cell. In fact, this enzyme is found in the cytosol, mitochondria and peroxisomes (Wrenger, C., 2003). Although it was thought to be involved in the production of NADPH utilized for both lipid biosynthesis and the neutralization of hydroxyperoxides, its role in ROS defense is now being fully understood.
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The parasite Plamodium falciparum causes malaria and depends on an elaborate antioxidative system to inhabit the host. This organism possesses TCA cycle enzymes
and the generation of NADPH from G6PDH is poor. Therefore in parasites subjected to oxidative stress, both the mRNA transcripts and protein levels of ICDH-NADP+ are up-regulated allowing the parasite to survive the defense system of the host (Wrenger, C. 2003). Although some organisms may defend themselves against ROS insult, ROS severely impede such critical enzymes as ACN, α-KGDH and pyruvate dehydrogenase (PDH) as they do possess the oxygen sensitive Fe-S cluster and SH group respectively.
α-Ketoglutarate Dehydrogenase (KGDH)
Krebs cycle enzymes are susceptible and more sensitive to oxidative stress because they are localized in the mitochondria and most bacterial membrane, the birthplace of oxidative species. α-KGDH, a housekeeping enzyme is crucial to all cells. It is essential in maintaining the cells redox state via the production of the reducing equivalent NADH and the production of ATP. It is needed for the catabolism and utilization of glutamate in glutaminolysis. It is an enzyme with multiple copies of three different subunits, E1, E2 and E3. The subunit E1 is unique to α-KGDH whereas E2 and E3 are common to both α-KGDH and PDH. It is highly regulated by substrates, cofactors and effectors such as calcium, ADP, Pi, ATP or GTP (Gibson, G. 2000). This enzyme is a flux regulatory enzyme and reflects the flux of intermediates through the TCA cycle. It is an important enzyme in respiration and maximal exercise (Anderson, U. 1998), since it generates NADH, a powerful reducing agent for complex I. This enzyme does also play a pivotal role in the malate-aspartate shuttle and acts as a collector of amino groups in tissues.
Oxidative stress and the inhibition of α-KGDH are common to Parkinson’s disease (PD) and many other neurodegenerative abnormalities. Often, this enzyme is as abundant as usual but has unusually low activity manifested in both affected and non affected areas of the brain. In some cases however, the protein levels are also affected and lead to the decrease in enzyme activity (Brown, A. 2000). It is often difficult to recreate neurodegenerative disease in model organisms. However, MPTP (1-methyl-4-phenyl-1,2,5,6- tetrahydropyridine) induces PD in some organisms. When oxidized, MPP+ inhibits complex I. This results in an increase in free radical generation. It is suggested that α-KGDH is a target of these free radicals due to the modification of thiol residues on the E2 subunit (Joffe, G. 1998). The enzyme α-KGDH may be inhibited by free radicals without the modification of the dihydrolipoic acid moieties (Gibson, E. 2002). Furthermore, compared to other mitochondrial enzymes, α-KGDH is the most sensitive to peroxinitrite and superoxide anions. However, the exact mechanisms how these inhibitory influences exert themselves are not clear (Anderson, U. 1998).
One sign that the inhibition of α-KGDH proceeds via oxidative stress is that thiamine deficiency causes the inactivation of α-KGDH before the appearance of phenotypic lesion (Gibson, G. 2000). Thiamine is a cofactor involved in the functioning of α-KGDH and PDH. Thiamine reduces oxidative stress and reduces lipid peroxidation and increases the reduced form of glutathione (Gibson, E. 2002). Thiamine deficiency is also involved in numerous neurodegenerative diseases where ROS are elevated and where α-KGDH is deleteriously affected.
One of the products of lipid peroxidation 4-hydroxy-2-nonenal (HNE) inhibits α-KGDH in vitro at much lower concentrations than those found in oxidative environments. The enzyme α-KGDH is a complex arrangement of 3 subunits and the cofactor lipoic acid. The latter is a target of HNE (Lucas, D., 1999). Hydrophobic interactions occur between the hydrocarbon chain of lipoic acid and HNE (Figure 9).
Figure 9: Reaction between HNE and the lipoic residue of α-KGDH (Taken from Humphries, K. 1998)
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Not surprisingly, PDH is also inactivated as a result of lipid peroxidation. On the other hand, HNE does not appear to directly inhibit other NADH dehydrogenase or mitochondrial electron transport chain components (Humphries, K. 1998). Other lipid peroxidation products which are lipophilic electrophiles could also interact with the dihydrolipoic moiety of α-KGDH and PDH.
The enzyme α-KGDH is also reduced in the brain of Alzheimer’s disease (AD) patients most likely due to its sensitivity to oxidative stress. In fibroblast, α-KGDH is inhibited by H2O2. It was therefore proposed that α-KGDH may be a peroxide sensor (Gibson, E. 2001). The association of oxidative stress and the defect in α-KGDH functioning is correlated with a decreased level of ferritin. The consequence is the destabilization of the cellular iron buffering capacity allowing the iron to become redox active and unsafe (Gibson, E. 2003). These observations depict an intricate relation between oxidative stress, metabolism and metal homeostasis. PDH, an enzyme that lies at the intersection of carbohydrate, lipid and amino acid metabolism and links these pathways together is also affected by ROS. It is severely impeded in such neurodegenerative disease like Friedreich’s Ataxia and the oxidation of the lipoic acid in the E2 subunit appears to be the target for oxidative damage (Blass, J.P. 1976).
Aconitase (ACN)
The enzyme ACN is a 4Fe-4S cluster-containing enzyme that binds citrate and catalyzes its isomerization to isocitrate (Figure 10).
Figure 10: Iron sulfur clusters are susceptible to oxidation in ACN (Taken from Voet, 1990)
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This protein is also crucial in the controlling of iron homeostasis. In fact, ACN is an iron sensor and helps in the attainment of iron homeostasis. This enzyme is known as an iron regulatory protein (IRP). These proteins are capable of binding iron regulatory elements (IRE) situated on the mRNA directing the synthesis of specific proteins involved in iron homeostasis (Eisenstein, R., 1998). When iron levels are high and sufficient for cellular processes, the Fe-S cluster of ACN is cubane and the enzyme is active. When iron is scarce the Fe-S cluster loses its Fe, and then is able to act as IRP. The IRP bind to IRE on the ferritin mRNA and prevents its translation and at the same time, stabilizes the mRNA responsible for the transferrin receptor. The result is the increase in iron internalization and in available iron (Figure 11).
Figure 11: State of iron levels and the modulation of ACN activity (Taken from Cairo, G., 2002)
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ACN is the most sensitive tricarboxylic acid (TCA) cycle enzyme to oxidative stress. The oxidation of iron-sulfur clusters by superoxide anions (O2·-) and H2O2 causes the inhibition of ACN. This necessarily results in the reduction of the flux of citrate through the TCA cycle. Patients with exercise intolerance are known to exhibit a reduction in ACN activity. It is believed that ACN acts as a circuit breaker preventing the continuous production of ROS through oxidative metabolism (Andersson, U. 1998). However, in some cases, this inactivation does not necessarily affect the energy metabolism since many organisms can convert glutamate to α-ketoglutarate and assure the continuity of the TCA cycle.
REDOX ACTIVE METALS and OXIDATIVE STRESS
Iron
Although ROS production is an integral component of oxidative cellular metabolism, abnormal metal homeostasis may be an important contributor of this toxic situation. The homeostasis of iron is tightly controlled at the uptake, utilization and storage levels (Figure 12). Any interference of these processes liberates iron which is now able to catalyze the one-electron reduction of oxygen reactive species that ultimately lead to the generation of free radicals (Kruszewski, M. 2003).
Figure 12: Pathways of iron homeostasis in hepatocytes (Taken from Goswami, T., 2002)
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Iron is crucial to metabolism. When iron is bound to transferrin, it is said to be in a safe state. Also, this association enables iron uptake by cells (Goswami, T. 2002). Excessive free iron has been thought to perpetuate oxidative stress in biological systems. In an organism, the highly damaging OH∙ is produced through the Haber-Weiss and Fenton reactions whenever iron is available.
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Increase in free iron and depletion of GSH pools are also associated with Parkinson’s disease (Gu, M. 1998). What is more compelling is that the amyloid β-peptide, a marker in AD can also generate H2O2 in the presence of redox metal such as Fe+3. When iron is found bound to the β-amyloid peptide, it becomes a catalytic site for the production of H2O2 from O2 (Cuajungca, M. 2003). Hence, free redox active iron ions are excellent markers of oxidative stress. It has been shown that in the reperfused cat retina, mobilized Fe+3 increases the severity of oxidative damage (Banin, E., 2000). Therefore, iron potentiates ROS-related diseases (Figure13).
Figure 13: Iron-mediated damage in biological systems (Taken from Polla, A., 2003)
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It has been postulated that a decrease in iron stores may retard the onset of age-related disease. Through the menstrual cycle, women eliminate large quantities of iron and this may account for their longer lifespan (Casanueva, E. 2003). Furthermore, the importance of iron for rapidly dividing cells renders the understanding of its metabolism an interesting target for therapeutic approaches to conquer cancer and other diseases. Hence, various iron chelating compound such as desferral are utilized as a therapy in order to abolish ROS-mediated diseases.
Copper
Another important metal implicated in the mediation of oxidative stress is copper. Since this metal is mainly stored in the liver, diseases targeting this organ will eventually affect copper homeostasis (Puig, S. 2002). Ceruloplasmin is the main copper binding protein and is responsible for copper transport to tissues. Redox active transition metals such as Cu+2 are useful as enzymatic cofactors and play a key role in oxidative metabolism. Proteins such as cytochrome C oxidase, tyrosinase, p-hydroxyphenyl pyruvate hydrolase and Cu-Zn SOD require copper for their functioning (Figure 14).
Figure 14: The cytochrome C oxidase reaction (Taken from Voet, 1990)
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However, this redox reactivity allows for the generation of oxidative species in situations where the metal is unbound. Patients suffering from Wilson’s disease are characterized with a faulty copper excretion mechanism through the bile and this may explain the noticeable increase in mitochondrial membrane peroxidation and reduced levels of vitamin E (Gaetke, L. 2003). This evidence suggests a role for copper in the generation of oxidative stress (Figure 15). Indeed, one explanation of copper toxicity lies in the redox capacity of the metal. Both copper atoms can generate oxidative intermediates through the following reactions:
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Figure 15: Cytotoxic mechanisms of copper Taken from Pourahmad, J., 2001)
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Like iron, in normal tissue, free Cu+2 does not exist (Figure 16). From the extracellular environment, Cu+2 atoms enter the cells and are delivered to chaperones such as cyclooxygenase (COX17) and autotaxin (Atx1)). Then, Cu+2 ions are brought to copper containing proteins such as SOD and cytochrome C oxidase. When the chaperoning of Cu+2 to enzymatic substrates is altered, the metal mediates oxidative stress. In the neurodegenerative disease amyotrophic lateral sclerosis (ALS), Cu+2 ions were unable to bind SOD. This enzyme represents 1% of the total proteins in many cells and is instrumental in scavenging superoxides. The inability of SOD to neutralize superoxide is the causative agent of this disease.
Figure 16: Homeostasis of copper (Taken from Puig, S., 2002)
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Another possible mechanism of copper toxicity is through the alteration of iron metabolism. When transport of copper to its Cu-proteins is affected, ceruloplasmin (Cp), a copper containing protein is rapidly disintegrated. This is detrimental since Cp also has efficient ferroxidase activity, allowing for the reduction of iron and its subsequent incorporation in transferrin where it is said to be safe (Carri, M. 2003). Therefore, deficiencies in copper homeostasis trigger improper iron handling and an inevitable rise in ROS.
Finally, when the incorporation of copper in SOD is faulty, the enzyme is inactivated. The inactivation of SOD creates albeit by an unknown mechanism an increase in the cellular demand for iron. Possibly, the ineffectiveness of SOD leads to an increase in ROS which in turn inactivate Fe-S cluster containing enzymes by oxidizing the iron and releasing it from the enzyme. This event triggers the need for iron. Many mitochondrial enzymes are therefore susceptible to ROS damage (Figure 17).
Figure 17: Alteration of iron metabolism by copper (Taken from Carri, M., 2003).
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NON-REDOX Metals, oxidative stress and metabolism
Zinc
Although the role of redox metals in the homeostasis of ROS is well documented, the involvement of non-redox metals is only beginning to become evident. Zinc is one such metal and its possible significance in neurodegeneration is now emerging. Recent reports suggest that small amounts of zinc bind to and cause the deposition of the β-amyloid peptide in the brain, a hallmark of Alzheimer’s disease (AD) (Cuajungco, M. 2003). Although the mechanism of action is different, non-redox metals can also lead to oxidative stress. Zinc does not participate in redox or Fenton chemistry reactions. Zinc is necessary for transcriptional and translational processes (Dineley, K.2003). However, in excess as in the case of injury, free intracellular zinc is toxic, particularly in the brain and is involved in some forms of AD. The homeostasis of zinc is crucial to neuronal survival. Although metallothionein may buffer elevated levels of zinc, intracellular zinc can be high in abnormal situations. A metal such as zinc can have various targets. It is believed that ROS generated by an increase in intracellular Zn+2 causes neurodegeneration (Sensi, S. 1999). The mitochondria appear to be the main target for Zn+2. Zinc also affects glycolysis through the inhibition of glyceraldehyde 3-phosphate dehydrogenase (GAPDH) thereby diminishing ATP synthesis. Furthermore, zinc impairs complex I respiration as a consequences of α-KGDH inhibition (Dineley, K. 2003). Although the precise mechanism of action is not known, it appears that zinc binds to the SH group in α-KGDH. The inhibition is first thought to be reversible but after increased exposure, zinc binds more tightly to the enzymatic complex and the inhibition becomes irreversible. Zinc may also lead to the generation of huge amount of peroxide in vitro by favoring the lipoamide dehydrogenase (α-KGDH component) reduction of molecular oxygen (Gazaryan, I. 2002).
NADH + O2 NAD+ + H2O2 Oxidase
It has been shown that other mitochondrial dehydrogenase enzymes such as succinate dehydrogenase (SDH), glutamate dehydrogenase (GDH) and malate dehydrogenase (MDH) are usually not affected by zinc. However, like α-KGDH, PDH is also inhibited by Zn+2. Zinc may also bind to and inhibit complex III of the electron transport chain (ETC), preventing the translocation of proton to the intramembraneous region, thereby affecting the mitochondrial membrane potential (Figure 18).
Figure 18: Biochemical targets of zinc (Taken from Dineley, K., 2003)
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Aluminium-mediated oxidative stress
Aluminum has no known biological functions but is toxic to all organisms. The availability of aluminum has greatly increased and the daily encounters with this metal can no longer be ignored. It is said that nearly 8 µg of aluminum accumulates in the human brain every year (Zatta, P. 2001). It has been suggested that aluminum may accumulate in bones whereby it is released at an older age because of bone demineralization. It would then accumulate in other organs such as the brain (Nayak, P., 2002). Intracellular aluminum is localized in organelles such as the mitochondria and the nucleus and is associated with a variety of neurodegenerative complications.
The relationship between aluminum neurotoxicity and Alzheimer’s disease is not fully understood. This trivalent metal ion interferes with the metabolism of carbohydrates and energy. It leads to the reduction in the process of acetylcholine biosynthesis and may also interact with ATPases thus affecting neurotransmitters release/uptake. It may also inevitably affect metal transporters. More importantly, aluminum may induce oxidative stress as a consequence of brain phospholipids peroxidation. Aluminum leads to in vivo lipid peroxidation in mice overexpressing the human amyloid precursor protein, a marker of AD. In this AD model, when mice were fed aluminum and vitamin E simultaneously, the lipid peroxidation was reduced. Lipid peroxidation was evident by the detection of isoprostan, a stable end product of free radical oxidation of polyunsaturated fatty acids. These isoprostans are present in postmortem AD individuals (Pratico, D. 2002). This study further establishes the involvement of oxidative stress in the cascade of events leading to neurodegeneration (Figure 19).
Figure 19: Relationship between aluminum exposure, oxidative stress and disease
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Aluminum may also have the ability to influence the pro-oxidant effect of iron in biological systems. Not only can aluminum displace iron in biomolecules it may also form complexes with superoxide with considerable oxidative power that are necessarily involved in biological oxidation.
O2-· + Al+3 AlO2·2-
This aluminum-superoxide complex has the potential to reduce Fe+3 back to Fe+2, enabling iron to undergo oxidation by H2O2 in a Fenton type reaction (Exley, C. 2004).
Fe+2 + O2 Fe+3 + O2-·
2O2-· + 2H- H2O2 + O2
Fe+2 + H2O2 OH- + HO· + Fe+3
If aluminum does indeed interfere with iron metabolism and does in fact form superoxide complexes, it may not be surprising that gallium (Ga+3), another trivalent metal ion may act in a similar fashion as a pro-oxidant non-redox metal (Exley, C., 2004)
Gallium
As our era of technological development continues, gallium arsenide (GaAs) semiconductors are being widely utilized as photon emitters in lasers for optical data storage and for high-speed optical communication. Satellite communication systems and ultra fast supercomputers depend on the semiconducting properties of gallium. Inevitably, gallium is becoming more prominent in our daily life. Therefore, the amount of gallium available in the environment has increased and will continue to increase in the foreseeable future (Chang, K., 2003). The toxicological repercussion cannot be ignored and studies are being designed to delineate the metabolic impact of gallium on living organisms. When GaAs was fed to rats, testicular toxicity was observed. The toxicity may not be attributable to arsenic itself and gallium has been shown to participate in mediating the aberrant conditions although the biological targets have not yet been fully identified (Omura, 1996).
The toxicity of gallium may inevitably be due to its capacity to interfere with iron metabolism as both have numerous common features including their positive charge of +3 and ionic radii of 0.67Ǻ. Nevertheless, their interactions with living organisms are different. Effectively, gallium a non-redox metal has no known biological functions (Al-Aoukaty, A., 1992), while iron is essential in all biological systems. Gallium can manifest its toxicity by displacing iron in essential molecules. The cytotoxicity of gallium is greatly enhanced when this trivalent metal is bound to transferrin. The affinity of gallium binding sites on transferrin is approximately ten fold lower than for iron. If present in high concentrations it binds to this ligand. In biological situations, gallium increases transferrin receptor expression due to the simulation of iron deprivation (Seligman, P. 1992). Ga+3-transferrin inhibits Fe+3-transferrin uptake. More importantly, Ga+3-transferrin alters the pH of the endosome, an intracellular iron store, thus decreasing the release of iron from internalized Fe+3-transferrin. Therefore, Ga+3-transferrin mimics iron depletion which causes the increase expression of transferrin receptors and the inhibition of cellular proliferation. Iron is essential in the synthesis of DNA, as it is a key component of ribonucleotide reductase, the only enzyme involved in the conversion of ribonucleic acid into deoxyribonucleic acid. This enzyme is inhibited by gallium as it competes for the iron binding site on the R2 subunit (Figure 20).
Figure 20: Ribonucleotide Reductase (Taken from Uhlin, 1994)
[pic]
The toxicity of gallium has been utilized to engineer this trivalent metal in the creation of antitumor drugs. Out of all antitumor group IIIa elements, gallium is the most potent and shows promise in the treatment of bladder cancer. Cells that proliferate at higher rates like cancer cells necessitate increase amounts of iron and therefore gallium is detrimental to these cells (Chitambar, C. 2003). This property allows the utilization of gallium in cancer therapy. Radiolabelled gallium (Ga67) is also used to visualize tumors. In vitro, gallium has been shown to destabilize the DNA helix. It can also compete with DNA-Mg+2 binding site and interferes with DNA processes. Gallium is also known to induce chromatin condensation, inhibit DNA polymerase, tyrosine phosphatase and prevents tubulin polymerization (Collery, P. 2002). Improving the bioavailability and targeting of gallium will allow the maximizing of antitumor activity of this metal.
Coincidentally, it was discovered that when gallium is used in cancer treatments, hypocalcemia is encountered. Consequently, even though gallium nitrate (Ga(NO3)3) can be toxic, it was ideal in the treatment of hypercalcemia related to cancers. Surprisingly, it was even effective in hypercalcemia cases where the level of parathyroid hormone related protein was present at high level. This metal will naturally accumulate at sites of bone remodeling through a transferrin or non-transferrin dependent pathway. Gallium appears to inhibit ATPases in the membrane and decrease the acidity of the extracellular environment, thereby reducing bone resorption (Bernstein, L., 1998). In fact, it is possible that gallium affects gene expression as well as protein synthesis since the metal has been shown to localize in the nuclei of osteoclast. Although the exact mechanism is unknown, gallium reduces the levels of serum alkaline phosphatase (increases bone formation) and encourages the excretion of hydroxyproline thereby inhibiting osteoclast activity and suppressing bone resorption (Hadjipavlou, A. 2001). Synergistically, gallium also acts on osteoblasts in an anabolic fashion and favors the production of type I collagen, a major component of the bone matrix. Gallium is also utilized for the treatment of Paget’s disease, a condition affecting the spinal structure and bone mass. Although gallium has some inherent therapeutic properties, its toxicity is undeniable due to its ability to mimic iron in various biological systems.
THESIS OBJECTIVES
When an organism is faced with an abnormal situation, there are three possible outcomes: 1. the organism may adapt. 2. the organism may die and 3. the organism may become dormant. Our laboratory has been elucidating the molecular mechanisms living systems utilize to adapt to environmental stress (Appanna, V., 1999). The literature is replete with information on adaptation scenarios usually involving proteins/enzymes necessary to combat stress situation; for instance, an ATP-dependent pump in cadmium (Cd+2) resistance or a mercury (Hg) reductase in the volatilization of Hg. The influence of these processes on the global cellular machinery has usually been ignored. We have utilized a holistic approach in an effort to delineate adaptation to stress; the involvement of the cellular metabolic network in stress adaptation strategy has been probed using the soil microorganism, Pseudomonas fluorescens, as a model organism. We have identified how metabolism is tailored in an effort to survive the stress imposed by a metal like aluminum (Al) (Hamel, R., 2003). Pseudomonas is an ideal choice as it proliferates rapidly in almost any carbon source. It is highly adaptable to various environments and offers enormous potential for commercial applications. Al is immobilized with the aid of oxalate and phosphatidylethanolamine (PE). The complete cellular metabolism is reconfigured in order to provide the precursors that drive the genesis of these moieties (Hamel, R., 2001). Isocitrate lyase (ICL) an enzyme that generates glyoxylate, a precursor for oxalate biosynthesis is markedly increased (Hamel, R., 2004). Enzymes such as isocitrate dehydrogenase (ICDH-NADP+) and glucose 6-phosphate dehydrogenase (G6PDH) that produce NADPH are also sharply enhanced.
Hence this study is undertaken with the aim of identifying the metabolic changes that enable P. fluorescens to survive gallium, a pro-oxidant and iron mimetic. Although the ability of this microbe to tolerate 1mM gallium has been shown (Al-Aoukaty., 1992), the significance of the metabolic network mediating the survival of P. fluorescens is not known. Thus, various enzymes involved in glycolysis, gluconeogenesis, the pentose phosphate pathway, the glyoxylate cycle, the tricarboxylic acid cycle and the oxidative phosphorylation have been evaluated. The influence of gallium especially as a generator of ROS and as an iron mimetic on these metabolic circuits has been assessed.
MATERIALS AND METHODS
List of Reagents and Equipments
2-Thiobarbituric acid; Sigma Chemical Company (St. Louis, Missouri)
2,4-Dinitrophenol; ICN Biochemicals (Cleveland,Ohio)
2,6-Dichloroindophenol; Sigma Chemical Company (St. Louis, Missouri)
5,5’- Dithio-bis-(2-nitrobenzoic acid); Sigma Chemical Company (St. Louis, Missouri)
6-Phosphogluconic acid (Barium salt); Sigma Chemical Company (St. Louis, Missouri)
Accumet pH Meter 910;Fisher Scientific (Unionville, Ontario)
Acrylamide; Bio-Rad Laboratories (Mississauga, Ontario)
Acetyl coenzyme A; Sigma Chemical Company (St. Louis, Missouri)
Adenosine 5' triphosphate (ATP); Sigma Chemical Company (St. Louis, Missouri)
α- Ketoglutaric acid;ICN Biochemicals (Cleveland,Ohio)
Ammonium chloride (NH4Cl); Sigma Chemical Company (St. Louis, Missouri)
Ammonium molybdate; Fisher Scientific (Unionville, Ontario)
Ammonium persulphate (APS);Bio-Rad Laboratories (Mississauga, Ontario)
Ammonium sulphate (NH4)2SO4; Sigma Chemical Company (St. Louis, Missouri)
Bio-Rad Mini-Protein II Dual Slab Cell; Bio-Rad Laboratories (Mississauga, Ontario)
Bis(2-hydroxyethyl)imino-tris(hydroxymethyl)methane ; Sigma Chemical Company (St. Louis, Missouri)
Bovine serum albumin; Sigma Chemical Company (St. Louis, Missouri)
Calcium chloride; ; BDH Laboratory Chemicals Division (Toronto, Ontario)
Centrifuge Model J2-MI; Beckman Instruments (Mississauga, Ontario)
Citric-2,4-13C acid; Isotech Inc (Miamisberg, Ohio)
Citric acid monohydrate; Sigma Chemical Company (St. Louis, Missouri)
Coenzyme A (sodium salt); Sigma Chemical Company (St. Louis, Missouri)
Coomassie G 250 Sigma Chemical Company (St. Louis, Missouri)
Coomassie R 250 Sigma Chemical Company (St. Louis, Missouri)
Deuterium oxide, 99.9 atom %D; Sigma Chemical Company (St. Louis, Missouri)
D- glucose; Sigma Chemical Company (St. Louis, Missouri)
D,L-aspartic acid; Fisher Scientific (Unionville, Ontario)DL-Dithiothreitol; Sigma Chemical Company (St. Louis, Missouri)
D,L-isocitric acid trisodium Salt; ICN Biochemicals (Cleveland,Ohio)
D,L-aspartic acid; Fisher Scientific (Unionville, Ontario)
ECL Plus™ reagents; Amersham Pharmacia Biotech (Piscataway, NJ, USA)
Ethylenediaminetetraacetic acid disodium salt; BDH Laboratory Chemicals Division (Toronto, Ontario)
ε-amino-n-caproic acid; Sigma Chemical Company (St. Louis, Missouri)
Ferric chloride (FeCl36H2O); Fisher Scientific (Unionville, Ontario):
Fumaric acid; Fisher Scientific (Unionville, Ontario)
Gallium (III) nitrate (hydrate); Sigma Chemical Company (St. Louis, Missouri)
Glacial acetic acid; CanLab (Toronto, Ontario)
Glucose-6-phosphate (disodium salt); Sigma Chemical Company (St. Louis, Missouri)
Glutamic acid (monosodium salt); Sigma Chemical Company (St. Louis, Missouri)
Glycerol; Sigma Chemical Company (St. Louis, Missouri)
Glycine; Sigma Chemical Company (St. Louis, Missouri)
Glyoxylic acid (monohydrate); Sigma Chemical Company (St. Louis, Missouri)
Guanidine hydrochloride; Sigma Chemical Company (St. Louis, Missouri)
Gyratory waterbath shaker model G 76; New Brunswick Scientific (Edison, New Jersey)
Hybond™- P: PVDF membrane; Amersham Pharmacia Biotech (Piscataway, NJ, USA)
Hydrochloric acid (HCl); CanLab (Toronto, Ontario)
Hydrogen peroxide (30% (w/w) solution); Sigma Chemical Company (St. Louis, Missouri)
Iodonitrotetrazolium chloride; Sigma Chemical Company (St. Louis, Missouri)
Isocitrate dehydrogenase EC 1.1.1.42 (porcine heart); Sigma Chemical Company (St. Louis, Missouri)
Lysozyme Grade 1; Sigma Chemical Company (St. Louis, Missouri)
Magnesium chloride hexahydrate (MgCl26H2O); BDH Laboratory Chemicals Division (Toronto, Ontario)
Malachite green (oxalate salt); Sigma Chemical Company (St. Louis, Missouri)
Malic acid; BDH Laboratory Chemicals Division (Toronto, Ontario)
Malic dehydrogenase (from porcine heart); Sigma Chemical Company (St. Louis, Missouri)
Malonic acid (disodium salt); Sigma Chemical Company (St. Louis, Missouri)
Menadione (sodium bisulfite); Sigma Chemical Company (St. Louis, Missouri)
n-Dodecyl β-D-maltoside; Sigma Chemical Company (St. Louis, Missouri)
Nitric acid (HNO3); CanLab (Toronto, Ontario)
Nicotinamide adenine dinucleotide (oxidized form); Sigma Chemical Company (St. Louis, Missouri)
Nicotinamide adenine dinucleotide (reduced form); Sigma Chemical Company (St. Louis, Missouri)
Nicotinamide adenine dinucleotide phosphate (oxidized form); Sigma Chemical Company (St. Louis, Missouri)
Nicotinamide adenine dinucleotide phosphate (reduced); Sigma Chemical Company (St. Louis, Missouri)
N,N-Methylene-bis-acrylamide; Bio-Rad Laboratories (Mississauga, Ontario)
N,N,N=,N=- Tetramethylenediamine (TEMED); Bio-Rad Laboratories (Mississauga, Ontario)
Oxaloacetic acid; Sigma Chemical Company (St. Louis, Missouri)
P-Anisidine; Sigma Chemical Company (St. Louis, Missouri)
Peroxidase (EC 1.11.1.7); Sigma Chemical Company (St. Louis, Missouri)
Phenazine methosulphate ; Sigma Chemical Company (St. Louis, Missouri)
Phenylmethylsulphonylfluoride (PMSF); Sigma Chemical Company (St. Louis, Missouri)
Potassium phosphate monobasic (KH2PO4); Sigma Chemical Company (St. Louis, Missouri)
Pseudomonas fluorescens ATCC 13525; American Type Culture Collection (Rockville, Maryland)
Pyruvate dehydrogenase Antibody: University of Glascow
Pyruvic acid (sodium salt crystalline); Sigma Chemical Company (St. Louis, Missouri)
Sodium carbonate anhydrous; Mallinckrodt Inc. (Kentucky)
Sodium phosphate dibasic (Na2HPO4); Sigma Chemical Company (St. Louis, Missouri)
Sodium hydroxide (NaOH); Fisher Scientific (Unionville, Ontario)
Sodium dodecyl sulphate (SDS); Sigma Chemical Company (St. Louis, Missouri)
Succinic acid; BDH Laboratory Chemicals Division (Toronto, Ontario)
Sucrose ; Sigma Chemical Company (St. Louis, Missouri)
Sulphuric acid (H2SO4);CanLab (Toronto, Ontario)
Tricarballylic acid; Sigma Chemical Company (St. Louis, Missouri)
Trichloroacetic acid; Fisher Scientific (Unionville, Ontario)
Tricine; Sigma Chemical Company (St. Louis, Missouri)
Tris(hydroxymethyl)aminomethane (Tris) HCl and Tris base; Sigma Chemical Company; (St. Louis, Missouri)
Tween-20; Bio-Rad Laboratories (Mississauga, Ontario)
Organism and culture conditions
The bacterial strain Pseudomonas fluorescens 13525 was obtained from the American Type Culture Collection (ATCC). The microbe was kept on a mineral medium containing citric acid in 2% agar. The sterile agar test tubes were maintained in a refrigerator at 4oC. The bacteria were subcultured every week.
Agar Media
In 250 ml of double distilled water were added Na2HPO4 (2.4 g); KH2PO4 (1.2 g); NH4Cl (0.4 g); MgSO4•7H2O (0.08 g); citric acid monohydrate (1.6 g) and 400 µl trace elements. (Trace element solution consisted of: FeCl3•6H2O (2µM); MgCl2•4H2O (1µM); Zn(NO3)2•6H2O (0.05 µM); CaCl2 (1µM); CoSO4•7H2O (0.25 µM) CuCl2•2H2O (0.1 µM); NaMoO4•2H2O (0.1 µM). The pH of the trace element solution was adjusted to 2.75 with diluted HCl to prevent precipitation of the metals and the solution was stored at 4oC). The pH was raised to 6.8 with dilute NaOH and the final volume was brought to 400 ml with double distilled water. The solution was gently heated and Bactoagar® (6.6 g) was added and stirred until completely dissolved. Approximately 7 to 10 ml were placed in test tubes and capped for slants. Following sterilization (autoclaved for 20 min at 17 lbs/in2, 121o C) the test tubes were laid on an angle and allowed to solidify at room temperature. Slants were stored in the refrigerator at 4 oC.
Preculture Media
The media used for the liquid preculture contained the following: Na2HPO4 (6.0 g); KH2PO4 (3.0 g); NH4Cl (0.8 g); MgSO4•7H2O (0.2 g); Citric acid monohydrate (4.0 g); Trace element solution (1.0 ml), per litre of deionized water. The pH was raised to 6.8 with dilute NaOH and the media was divided into 100 ml aliquots in 250 ml Erlenmeyer flasks. The flasks were capped with foam plugs and autoclaved for 20 min at 17 lbs/in2, 121 oC. These preculture media were inoculated with a loop of P. fluorescens stored on agar slants. Stationary phase was attained following 24 to 48 hrs of incubation.
Growth media
The following reagents were added in the following order: Na2HPO4 (6g); KH 2PO4 (3g); NH4Cl (0.8 g); MgSO4•7H2O (0.2 g); trace element solution (1 ml); citric acid monohydrate (4.0 g) or metal-citrate solution in the case of metal supplemented medium to 600 ml of double distilled water. The pH of the medium was raised to 6.8 with dilute NaOH and the volume was adjusted to 1 L with water. The media was separated in 200 ml amounts in 500 ml Erlenmeyer flasks and inoculated with 1 ml of the precultured bacteria. The cultures were incubated at 26oC in a gyratory water bath shaker model G76 (New Brunswick Scientific) at 140 rev. min- 1.
Control growth media
The media without the test metal(s) constituted the control media. The media were dispensed in 200 ml amounts in 500 ml Erlenmeyer flasks, stoppered with foam plugs and autoclaved for 20 min at 121oC.
Metal growth media
Media supplemented with various metals were prepared. These media were prepared in the same manner as their respective control medium with the following modifications: 4 g citric acid monohydrate and the metal were first allowed to complex in approximately 50 ml deionized water for approximately 60 min prior to being added to the remainder of the media. Studies involving the use of a Ga-citrate media contained 1 mM Ga(NO3)3 complexed to citric acid. The media were prepared as above, however 0.2557 g of Ga(NO3)3 (M.W. 255.7) and 4.0 g of citric acid monohydrate (MW 210.1) was used in order to obtain a final concentration of Ga3+ and citric acid of 1 mM and 19 mM respectively. Studies utilizing other metal-citrate media were also utilized and consisted of 19 mM citrate complexed to 0.1 mM gallium,15 mM aluminum, 1 mM calcium or 1mM gallium/20 µM iron or 1 mM gallium/100 µM iron respectively.
Harvest of P. fluorescens
P. fluorescens were collected from the growth medium by centrifugation at 10,000 g for 15 min at 4oC. The supernatant was removed and 0.85% NaCl was utilized to suspend the bacterial pellet. The bacteria was centrifuged again for 15 min and the procedure was repeated (Figure 21).
Figure 21: Collection of bacterial cells
Preparation of Cell Free Extract (CFE) From Whole Cells
When the bacteria were harvested as indicated above, they were suspended in cell storage buffer consisting of 50 mM Tris-HCl, 5 mM MgCl2, 1 mM PMSF, 1 mM DTT at pH 7.3. The cells were disrupted by sonication using a Brunswick sonicator, power level 4 for 15 sec at 4 intervals. Samples were kept on ice and allowed to cool between intervals for at least 10 min (Figure 22). The supernatant fraction of CFE was collected and centrifuged at 180,000 × g for 60 min at 4oC to yield membrane and soluble components. The soluble fraction was removed and centrifuged again at 180,000 × g for 2 hrs to insure a membrane free system. Both the membrane and soluble fraction were kept on ice in the refrigerator for a maximum of three days.
Figure 22: Isolation of CFE from whole cells
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Isolation of spheroplasts of P. fluorescens
The inner membrane of P. fluorescens was isolated by a modified version of the method described by Mizuno MACROBUTTON HtmlResAnchor (Mizuno and Kageyama, 1978) (Figure 23). The cells were harvested (as seen in Figure 21) and then washed with 20% (w/v) sucrose. Cells (1.5 g wet weight) were suspended in 18 ml of ice-cold 20% (w/v) sucrose and ice-cold reagents were slowly added to the suspension in an ice bath in the following order; 9 ml of 2 M sucrose, 10 ml of 0.1 M Tris-HCl (pH 7.8 at 25oC), 0.8 ml of 1% Na-EDTA (pH 7.0), and 1.8 ml of 0.5% lysozyme. The mixture was then warmed to 30 oC within a period of 5 min and kept in the gyratory bath at that temperature for 60 min. The suspension was centrifuged to remove the spheroplasts at 10 000 × g for 30 min at 30 oC. The spheroplasts were incubated with the lysis buffer consisting of 40 ml of 50 mM Tris, 5 mM MgCl2 1mM PMSF and 1 mM DTT and the spheroplast membranes were recovered by centrifugation at 100 000 × g for 30 min and washed in the same buffer.
Figure 23: Isolation of spheroplasts of P. fluorescens
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13C NMR analyses of citrate metabolism in CFE
13C NMR analyses were performed using a Varian Gemini 2000 spectrometer operating at 50.38 MHz for 13C (carbon). Experiments were conducted with a 5mm dual probe (35o pulse, l-s relaxation delay, 8 kilobytes of data. Chemical shifts were referenced to shifts of standard compounds obtained under the same conditions. Membrane fraction equivalent to 2 mg/ml of proteins obtained after 25 hrs of growth in a citrate medium or 65 hrs for Ga-stressed medium were placed in a phosphate buffer (10% D2O). The reaction was initiated in 1.5 ml conical tubes by addition of labeled (5 mM) Ga-citrate (1:1) and [2,4- 13C2] citrate respectively and, if required for enzyme activity, 0.5 mM of the respective cofactor was utilized. Following 60 min incubation at 26 oC, the membrane fractions were frozen overnight and subjected to 13C NMR proton decoupled analyses.
1H NMR analysis of citrate metabolism in CFE from P. fluorescens
1H NMR analyses were performed using the Varian Gemini 2000 spectrometer operating at 200 MHz for 1H. Enzymes were assayed in 1H NMR buffer (10 mM phosphate, 5 mM MgCl2 and pH 7.4) with 500 µg of membrane protein, 2 mM substrate and, if required for enzyme activity, 1 mM of the respective cofactor was utilized. The experiments were performed in 1.5 ml conical tubes and the reactions were stopped by placing the tubes in a boiling water bath for 3 min. The formation of any precipitate and/or the presence of membranes were removed by centrifugation at 20 000 × g for 15 min. The supernatant was lyophilized and dissolved in 500 µl deuterium oxide (D2O), 99.9 atom % D. The water resonance was suppressed with the aid of the homodecoupler set to the signal attributable to water. The following parameters were used: Decoupler modulation mode (dmm=ccc), where c=continuous; decoupler modulation (dm=nyn), where n=no and y=yes; decoupler low power (dlp=2000); the first delay (d1=0); the second delay (d2=5); the first pulse (p1=2); and the acquisition time (at=1). The number of transients varied among samples. Occasionally, d2, p1, at, and dlp were varied to achieve maximal water suppression. Experiments were executed with a 5 mm dual probe at a 90o pulse angle, and 8 kilobytes of data.
Measurement of oxidized lipids
Thiobarbituric acid is known to react with oxidized lipids. The amount of thiobarbituric acid reactive species was measured in the inner membrane fraction of control and Ga-stressed cells at various times of growth as described by Buege (Buege, J.A., 1978). Briefly, 2 mg of protein equivalent of inner membrane was heated with 15% TCA, 0.375% TBA/0.25N HCl in a final volume of 1.0ml for 15 min. A pinkish color developed and the samples were centrifuged for 10 min at 10 000 × g. The supernatant was isolated and the absorbance was measured at 532 nm. Blanks did not contain any inner membranes. The extinction coefficient was ε= 1.56 E 105 M-1cm-1.
[pic]
Protein carbonyl measurement
The protein carbonyl content was determined according to the method described by Vendemiale (Vendemiale, G., 2001) in control and Ga-stressed cells at stationary phase of growth. Briefly, 1.0 mg of soluble protein equivalent was allowed to react with 2% DNPH in a final volume of 1.0 ml for 60 min. Subsequently, 200 µl of 50% TCA was added to each sample to precipitate the proteins. The proteins were then spun in a tabletop centrifuge at 10,000 × g for 10 min. The supernatant was discarded and washed with a solution of 10% TCA and recentrifuged. This was repeated two more times upon which the pelleted proteins were washed with a solution of ethylacetate:ethanol in a 1:1 ratio three times. The final precipitate was dissolved in 1.0ml of 6M guanidine and the absorbance was measured at 370 nm. The extinction coefficient for hydrazones was 21.5 nmol*L-1cm-1. Blanks did not contain any soluble proteins.
[pic]
Peroxide measurement in cellular fractions exposed to gallium
The amount of H2O2 was measured in the membrane fraction of control cells (3 mg), in 25 mM Tris-HCl/ 5 mM MgCl2 buffer (pH=7.3), subjected to either 5 mM citrate or 5mM Ga-citrate. To the reaction mixture were added immediately 4 units of peroxidase as well as 10 mM P-anisidine in a final volume of 1.0 ml. The reaction was allowed for 30 min and the absorbance was measured at 458 nm. Blanks did not contain the substrate citrate or Ga-citrate. The amount of peroxide produced was measured and quantified (P-anisidine ε =1.173 M-1cm-1, Munoz, C., 1997).
Superoxide measurement in cellular fraction exposed to gallium
The amount of superoxide was measured in the membrane fraction of control cells (3 mg), assayed in 25 mM Tris-HCl/5 mM MgCl2 buffer (pH=7.3), subjected to either 5 mM citrate or 5 mM Ga-citrate. INT (0.12 mM) was added to the reaction mixture immediately. The reaction was allowed to react for 5 hrs and the absorbance was measured at 485 nm. Blanks did not contain the substrate citrate or Ga-citrate. The amount of superoxide produced was measured using the extinction coefficient of 11 mM-1cm-1 for INT (Poinas, A., 2002).
Measurement of Fe-S cluster in proteins
The integrity of iron containing enzymes in the soluble fraction of control and Ga-stressed cells was monitored using UV/VIS scanning spectroscopy. Briefly, 1.0 mg of soluble protein equivalent topped off to 1.0ml with activity buffer was submitted to UV/VIS (200 nm-900 nm) scanning. The integrity of iron containing proteins attributable to a band in the 395-415 nm region was monitored (Soum, E., 2003)
Measuring enzyme activity in CFE from P. fluorescens
The CFE from Pseudomonas fluorescens were isolated as previously indicated in Figure 22. The protein content of each fraction was measured by the method of Bradford MACROBUTTON HtmlResAnchor (Bradford, 1976) using the kit supplied by BioRad. The methods utilized to monitor the various enzymatic activities are described below. The mean of specific activities and the standard deviation were calculated for each enzyme.
Catalase activity
The activity of catalase (EC 1.11.1.6) was measured with the aid of P-anisidine and the absorbance at 458 nm was monitored. Briefly, 200 µg of control or Ga-stressed soluble proteins obtained at various times of growth were incubated with 15mM hydrogen peroxide. 10 mM P-anisidine was added immediately in a final volume of 1.0 ml and the absorbance was measured after 60 min. Blanks were prepared similarly, however the addition of hydrogen peroxide was omitted (Igamberdiev, A., 1995)
Superoxide dismutase (SOD) activity
The activity of SOD (EC 1.15.1.1) was measured with the aid of INT at 485 nm (ε=11mM-1cm-1). The enzyme was assayed by a modified method from Beyer (Beyer, W. 1987). Briefly, 200 µg of control and Ga-stressed membrane proteins obtained at various times of growth were incubated with 5 mM menadione, a superoxide generating compound. 15 ul of INT (4mg/ml) was added for a final volume of 1.0ml and the absorbance was measured after 1-4 hrs. Blanks were prepared similarly, however the addition of menadione was omitted. Menadione was used to obtain a standard curve.
Assay for aldehydes and ketoacids
Levels of aldehydes and ketoacids were measured using 2,4-dinitrophenylhydrazine (DNPH) as reported by Katsuki (Katsuki, H., 1971) and modified by Romonov (Romanov, R, 1999). DNPH reacts readily with almost all aldehydes and ketones to yield 2,4-dinitrophenylhydrazones. Initially the reaction with the ketoacids takes place under acidic conditions (5mM DNPH in 2N HCl), and then addition of base (1N NaOH) was used to deprotonate and colourise the 2,4-dinitrophenylhydrazones. The absorbance was then monitored at 450 nm (ε = 16,000 M-1 cm-1).
All enzymatic reactions were performed in a final volume of 1 ml 50 mM Tris buffer, pH 7.3 containing 5 mM MgCl2. Just prior to stopping the reaction the samples were divided into 2 x 0.5 ml fractions at which time 0.1 ml 2,4-DNPH (5 mM in 2 N HCl) was added to stop the reaction. The samples were allowed to stand at room temperature for 15 min. The sample was diluted to 1 ml with water and 1 ml of NaOH (1N) was added. The absorbance at 450 nm arising from dinitrophenylhydrazone was measured within 10 min. Appropriate controls were utilized for each enzymatic assay. The respective keto acids and aldehydes served as standards.
Isocitrate Lyase (ICL) activity
ICL (EC 4.1.3.1) activity was assayed in 25 mM Tris-HCl buffer (pH 7.3) containing 5 mM MgCl2, 2 mM isocitrate, and approximately 0.1 mg ml-1 soluble protein. Blanks and controls were prepared in a similar manner except the substrate, isocitrate, was omitted. Enzyme activity was determined spectrophotometrically by monitoring the production of glyoxylate with 2,4-DNPH at 450 nm (Romanov, V., 1999). The increase in intensity of color is proportional to the glyoxylate produced. Glyoxylate was used as a standard.
ICL
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Isocitrate dehydrogenase (ICDH-NAD+) activity
ICDH-NAD+ (EC 1.1.1.41) catalyzes the oxidative decarboxylation of isocitrate to form (- ketoglutarate. ICDH-NAD+ activity was assayed in 25 mM Tris-HCl/5 mM MgCl2 buffer (pH 7.3) containing 4 mM isocitrate and 0.5 mM NAD+, 8 mM malonate (to inhibit any contaminating ICL activity) and approximately 0.4 mg ml-1 membrane protein. Blanks were prepared in a similar manner except the substrate, isocitrate, was omitted. ICDH-NAD+ activity was determined by measuring the formation of α- ketoglutarate. The amount of ketoacid produced was determined spectrophotometrically using 2, 4-DNPH (Romanov, V., 1999) and α- ketoglutarate served as the standard.
ICDH
[pic]
α-Ketoglutarate dehydrogenase (α-KGDH) activity
(-KGDH (EC 1.2.4.2) activity was assayed in 25 mM Tris-HCl buffer (pH=7.3) containing 5 mM MgCl2. To the buffer was added 0.3 mM (-ketoglutarate, 0.1 mM Coenzyme A and 0.5 mM NAD+ and approximately 0.2 mg ml-1 membrane protein. The disappearance of α-ketoglutarate was followed colorimetrically with the aid of DNPH (Romanov, V., 1999). Blanks contained 25 mM Tris-HCl/5 mM MgCl2 buffer (pH 7.3), 0.1 mM Coenzyme A and 0.5 mM NAD+ and approximately 0.2 mg ml-1 membrane protein but no substrate. A solution of 1.0ml containing 0.3 mM α-ketoglutarate served as standard.
[pic]
CoA independent α-Ketoglutarate decarboxylase (α-KGD) activity
α-KGD (EC 4.1.1.71) is able to decarboxylate α-ketoglutarate to succinate in a CoA independent fashion. This enzyme was assayed in both the soluble and membrane fraction of control and Ga-stressed cells. The enzyme was probed according to Laura Green (Green, L. 2000) with the following modification; the reaction mixture consisted of 25 mM tris/5 mM MgCl2 buffer supplemented with 0.4 mg of protein, 0.3 mM α-ketoglutarate in a final volume of 1.0 ml. The disappearance of α-ketoglutarate was followed colorimetrically with the aid of DNPH (Romanov, V., 1999). Blanks contained 25 mM Tris-HCl buffer (pH 7.3) and approximately 0.4 mg ml-1 membrane/soluble protein but no substrate. A solution of 1.0ml containing 0.3 mM α-ketoglutarate served as standard.
α-Ketoglutarate reductase (α-KGR) activity
The activity of α-KGR (EC 1.1.99.2) catalyzes the reduction of α-ketoglutarate to hydroxyglutaric acid and utilizes NADH as a cofactor. This enzyme was probed in both the soluble and membrane fraction of control and Ga-stressed cells. The enzyme was assayed according to Zhao (Zhao, G. 1996) with the following modification; the reaction mixture consisted of 25 mM tris/5 mM MgCl2 buffer supplemented with 0.4 mg of protein, 0.3 mM α-ketoglutarate and 0.5 mM NADH in a final volume of 1.0 ml. The disappearance of α- ketoglutarate was followed colorimetrically with the aid of DNPH (Romanov, V., 1999). Blanks contained 25 mM Tris-HCl buffer (pH 7.3), approximately 0.4 mg ml-1 membrane/soluble protein and NADH but no substrate. A solution of 1.0ml containing 0.3 mM α-ketoglutarate served as standard.
Glutamate dehydrogenase (GDH) activity
The enzyme GDH (EC 1.4.1.2) catalyzes the oxidative deamination of glutamate with the aid of NAD+ to α-ketoglutarate. This enzyme was monitored in the membrane fraction of control and Ga-stressed bacteria. Briefly in activity buffer (25 mM Tris/5 mM MgCl2) was added 2.0 mM glutamate, 0.5 mM NAD+ and 0.2 mg of membrane protein equivalent in a final volume of 1.0 ml. The formation of α-ketoglutarate was followed colorimetrically with the aid of DNPH (Romanov, V., 1999). Blanks contained 25 mM Tris-HCl/5 mM MgCl2 buffer (pH 7.3), approximately 0.2 mg ml-1 membrane and 0.5 mM NAD+ but no substrate. A solution of 1.0ml containing 0.3 mM α-ketoglutarate served as standard.
Citrate synthase (CS) activity
CS (EC 2.3.3.1) mediates the condensation of oxaloacetate and acetyl-CoA to produce citric acid. For the measurement of this reaction 0.2 mg ml-1 protein equivalent was incubated with 25 mM Tris-HCl/5 mM MgCl2 buffer (pH 7.3) containing, oxaloacetic acid (1 mM), acetyl-CoA (0.1 mM), and DTNB (0.1 mM). The increase in absorbance from the formation of free thionitrobenzoate, an ion produced by the reaction of DTNB with HSCoA, a product formed enzymatically, was monitored at 10 sec intervals for 10 min at A412 (ε = 13.6 mM-1 cm-1) (Williams, A., 1998).
[pic]
Malate synthase (MS) activity
MS (EC 2.3.3.9) activity was determined spectrophotometrically by monitoring the disappearance of coenzyme A in the presence of dithiobenzoic acid (Williams, A., 1998). In this method 0.2 mg ml-1 of soluble protein equivalent was incubated with glyoxylate (1 mM), acetyl-CoA (0.1 mM), DTNB (0.1 mM) in 25 mM Tris-HCl/5 mM MgCl2 buffer (pH 7.3). The increase in absorbance from the formation of free thionitrobenzoate ion was monitored at 10 sec intervals for 10 min at A412 (ε = 13.6 mM-1 cm-1) (Williams, A., 1998).
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Pyruvate dehydrogenase (PDH) activity
PDH (EC 1.2.4.1) catalyzes the oxidative decarboxylation of pyruvate to acetyl-CoA with the concomitant release of NADH from NAD+. The activity of this enzyme was monitored in the membrane fraction of CFE by measuring the consumption of pyruvate with the aid of DNPH (Romanov, V., 1999). Membrane protein (0.4 mg ml-1) were incubated with pyruvate (0.2 mM), CoA (0.1 mM), and NAD+ (0.5 mM) in a final volume of 1.0 ml. The absorbance at A450 was measured and pyruvate was used as the standard. Blanks consisted of the above mixture in which the substrate pyruvate was omitted.
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Succinate Dehydrogenase (SDH) activity
SDH (EC 1.3.5.1) catalyzes the oxidation of succinate to fumarate. Flavine adenine dinucleotide (FAD) is covalently bound to SDH. For the enzyme to complete its catalytic cycle, the electrons from the reduced flavin cofactor are normally passed on to the electron transport chain. 2,6-Dichlorophenol indophenol (DCPIP) was utilized as an artificial electron acceptor. DCPIP absorbs strongly at 600 nm (ε = 22,000 M-1 cm-1), when oxidized and becomes colorless in its reduced state. The decrease in intensity of the color measured at 600 nm is proportional to the measure of SDH activity. SDH activity was assayed according to the method as described by Maklashina and Cecchini, (Maklashina, 1999), with the following modifications; in 1.0 ml, the assay consisted of 25 mM Tris–HCl, 5 mM MgCl2, 10 mM succinate, 12.5 mg ml-1 DCPIP, 5 mM KCN ( to block the electron transport chain). The reaction was initiated by the addition of 0.2 mg ml-1 membrane protein equivalent and A600 was monitored at 10 second intervals over 100 sec.
[pic]
Malic enzyme (ME) activity
The oxidative decarboxylation of malate to pyruvate is catalyzed by ME (EC 1.1.1.38), an enzyme that uses NADP+ as a co-substrate. The reduction of NADP+ was determined by monitoring the formation of NADPH at A340. The assay was carried as previously described (Wynn, J., 1997) with the following modifications: the reaction was carried out at 26 °C, pH 7.3 and consisted of 25 mM Tris–HCl, 5 mM MgCl2, 2 mM malate, 0.5 mM NADP+, and 0.2 mg ml-1 soluble protein for a final volume of 1.0 ml. The absorbance at 340 nm was plotted over 200 seconds at 10 second intervals. The specific activity was calculated using the molar extinction coefficient for NADPH (6.22 mmol/L for a path length of 1.0 cm). The formation of pyruvate with the aid of DNPH was also monitored.
Glucose-6-phosphate dehydrogenase (G6PDH) activity
The oxidation of glucose-6-phosphate to form phosphoglucono-δ- lactone is catalyzed by G6PDH (EC 1.1.1.49), an enzyme that uses NADP+ as a co-substrate. The reduction of NADP+ was determined by monitoring the A340 according to the method described by Wynn (Wynn, J., 1997). The following modification were performed; the assay consisted of 25 mM Tris–HCl, 5 mM MgCl2 buffer (pH 7.3) with 1 mM glucose-6-phosphate, 0.5 mM NADP+, and 0.2 mg ml-1 soluble protein for a final volume of 1.0 ml. The absorbance at 340 nm was recorded over 5 min at 10 second intervals. The specific activity was calculated using the molar extinction coefficient for NADPH (6.22 mmol/L for a path length of 1.0 cm).
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6-Phosphogluconate dehydrogenase (6PGDH) activity
6-PGDH (EC 1.1.1.44) catalyzes the oxidative decarboxylation of 6-phospho-D-gluconate to form D-ribulose 5-phosphate using NADP+ as a co-substrate. The reduction of NADP+ was determined by monitoring the A340 according to the method described by Wynn (Wynn, J., 1997) with the following modifications; the assay consisted of 25 mM Tris–HCl, 5 mM MgCl2 buffer (pH 7.3) to which was added 1 mM 6-Phosphogluconate, 0.5 mM NADP+, and 0.2 mg ml-1 soluble protein for a final volume of 1.0 ml. The absorbance at 340 nm was plotted over 10 min at 10 second intervals. The specific activity was calculated using the molar extinction coefficient for NADPH (6.22 mmol/L for a path length of 1.0 cm).
Isocitrate dehydrogenase (ICDH-NADP+) activity
The oxidative decarboxylation of isocitrate to form α- ketoglutarate using NADP+ as a co-substrate is catalyzed by ICDH-NADP+ (EC 1.1.1.44). The reduction of NADP+ was determined by monitoring the formation of NADPH at A340 as described by Plaut (Plaut, G., 1983). The following modifications were performed: the assay consisted of 25 mM Tris–HCl (pH 7.3), 5 mM MgCl2 , 2 mM isocitrate, 0.5 mM NADP+, and 0.1 mg ml-1 soluble protein. The reaction was also performed in the presence of 4 mM malonate (to inhibit ICL). The absorbance at 340 nm was plotted over 100 seconds at 10 second intervals. The specific activity was calculated using the molar extinction coefficient for NADPH (6.22 mmol/L for a path length of 1.0 cm). α-ketoglutarate formation was also recorded via the DNPH assay.
Malate dehydrogenase (MDH) activity
The oxidation of malate to oxaloacetate using NAD+ as cofactor is catalyzed by MDH (EC 1.1.1.37). 2,4-DNPH at 450 nm (Romanov, V., 1999) was utilized to monitor the formation of oxaloacetate. The assay consisted of 25 mM Tris-HCl. (pH 7.3), 5 mM MgCl2 1 mM malate, 0.5 mM NAD+ and 0.2 mg ml-1 membrane protein equivalent over 7 min. Oxaloacetate served as the standard. The omission of malate in the above mixture served as blanks.
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Fumarase (FUM) activity
The conversion of fumarate to malate is catalyzed by FUM (EC 4.2.1.2). 2,4-DNPH at 450 nm MACROBUTTON HtmlResAnchor (Romanov et al., 1999) was utilized to monitor the formation of oxaloacetate. The assay consisted of 25 mM Tris-HCl/5 mM MgCl2 (pH 7.3), 1 mM fumarate, 0.5 mM NAD+ and 0.2 mg ml-1 membrane protein equivalent over 7 min. Oxaloacetate served as the standard. The omission of fumarate in the above mixture served as blanks. The activity of FUM was calculated taking in consideration MDH activity.
Aconitase (ACN) activity
The activity of ACN (EC 4.2.1.3) was determined in the soluble fraction of CFE. Precautions were taken to increase the stability of ACN in the soluble fraction of CFE. Therefore, 10% tricarballylic acid was added to the whole cells prior to sonication (cell disruption). The assay consisted of 25 mM Tris–HCl, 5 mM MgCl2, 10 mM substrate (citrate) and 0.2 mg ml-1 soluble protein. The reaction was monitored at 240 nm for the formation of cis-aconitate as previously described MACROBUTTON HtmlResAnchor (Jordan et al., 1999). Aconitate served as the standard. Blanks were prepared in a similar fashion however, omitting the substrate from the mixture.
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Hexokinase activity
Hexokinase (EC 2.7.1.1) was assayed in 25 mM Tris/5mM MgCl2 activity buffer (pH 7.3) containing 0.4 mg/ml equivalent of soluble proteins, 2 mM glucose, 2 mM ATP and 0.5 mM NADP+ in a final volume of 1.0 ml. The reaction was allowed to react for 20 min and the specific activity was calculated using the molar extinction coefficient for NADPH (6.22 mmol/L for a path length of 1.0 cm) (Wynn, J., 1997). The activity of G6PDH was taken into consideration before obtaining activity values for hexokinase
Aspartate Transaminase (AST) activity
AST (EC 2.6.1.1) was assayed in 25 mM Tris/5mM MgCl2 activity buffer. In a final volume of 1.0ml was added in the following order 5 mM aspartate, 0.5 mM NADH, 2 µl MDH (1 unit/0.12 µl), 0.2 mg of soluble proteins and finally, 5mM of α-ketoglutarate. The consumption of NADH by MDH was monitored at 340 nm and plotted over 5 min. The specific activity was calculated using the molar extinction coefficient for NADH (6.22 mmol/L for a path length of 1.0 cm) (Wynn, J., 1997).
Glucose-6-phosphate phosphatase (G6PP) activity
G6PP (EC 3.1.3.69) was assayed in 25 mM Tris/5mM MgCl2 activity buffer. In a final volume of 1.0 ml was added 0.4 mg of proteins, and 2.0 mM glucose-6-phosphate. The reaction was allowed for 30 min and the reaction samples were heated for 3 min in a water bath to terminate the reaction. The samples were then spun at 10 000 × g for 20 min to remove protein debris. Thereafter, 20 ul of each samples were diluted to 1.0 ml with activity buffer. According to the method described by Baykov (Baykov, A., 1988), 500 µl of working solution (10 ml of malachite green (0.44g in 1:5 ratio water:concentration H2SO4), 2.5 ml of 10% ammonium molybdate and 200 µl of 11% tween 20) was added to the 1.0 ml diluted samples for 10 min and the absorbance was recorded at 630 nm. G6P and buffer alone served as blank.
Pyruvate Carboxylase (PC) activity
PC (EC 6.4.1.1) was assayed in 25 mM Tris/5mM MgCl2 activity buffer. In a final volume of 1.0 ml was added 0.2 mg of proteins, 10 mM HCO3, 1 mM ATP and 2.0 mM pyruvate. The reaction was allowed for 30 min and the reaction samples were heated for 3 min in a water bath to terminate the reaction. The samples were then spun at 10 000 × g for 20 min to remove protein debris. Thereafter, 40 ul of each samples were diluted to 1.0 ml with activity buffer. According to the method described by Baykov (Baykov, A., 1988), 500 µl of working solution (10 ml of malachite green, 2.5 ml of 10% ammonium molybdate and 200 µl of 11% tween 20) was added to the 1.0 ml diluted samples for 10 min and the absorbance was recorded at 630 nm. Standards were obtained as mentioned above, however proteins were omitted.
LIST OF BUFFERS
Cell storage buffer
50 mM Tris-HCl
1 mM EDTA
1 mM PMSF
1 mM DTT
15 mM MgCl2
pH 7.3 at Room Temperature
1H NMR Buffer Activity Buffer
10 mM sodium phosphate 25 mM Tris HCl
5 mM MgCl2 5 mM MgCl2
pH 7.3 at Room Temperature pH 7.3 at Room Temperature
Statistical Analyses
The student t test value was calculated to determine the significance of the difference in specific activity of various enzymes in control compared to Ga-stressed grown bacteria.
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If the calculated t value exceeds the tabulated value of 2.35 for n=3 then the means are significantly different and p is said to be ≤ 0.05 (Zar, J., 1999)
Enzymatic activities at various growth intervals
Pseudomonas fluorescens were grown on medium containing Ga-citrate or citrate (control) as the only carbon source prepared as indicated in the previous section. At specified timed intervals the bacterial cells were harvested and the CFE were isolated as described before. A Bradford assay was performed to determine the protein content from CFE MACROBUTTON HtmlResAnchor (Bradford, 1976) and the specific activities of different enzymes were monitored as indicated above.
Enzyme activities as a function of gallium in the growth media
Media were prepared as mentioned in the Media and Growth conditions section with the following modifications: The gallium content was varied from media not containing gallium (control cultures) to media containing 0.1 mM Ga and cells were harvested at early stationary phase (40hrs). The CFE were isolated and the specific activities of various enzymes were determined as previously described.
Gallium, ROS and the modulation of enzymatic activities
Pseudomonas fluorescens was grown in Ga-citrate for 65 hrs and 10 mg of whole cell protein equivalent was transferred to a medium of citrate (100 ml), to a medium of citrate supplemented with 15mM H2O2 or 1mM menadione. These cultured were allowed to grow for 6 hrs. The CFE were assessed for activities of various enzymes.
Influence of Ga(NO3)3 on α-KGDH activity
The membrane fraction of control cells (0.5mg) was allowed to interact with various amount of Ga(NO3)3 (0.1 mM – 0.5 mM) for 15 min in the refrigerator. The specific activity of α-KGDH was measured as previously described.
Electrophoresis
Blue Native Polyacrylamide Gel Electrophoresis (BN PAGE) (gradient gels 4% - 16%)
1 mm spacers were used to make small gels for the BioRad MiniProtean™ 2 system. The final volume of one separating gel was 5.8 ml, therefore 2.9 ml 4% acrylamide and 2.9 ml 16% acrylamide solutions per gel where used to create a linear gradient (4%-16%) using a gradient former (BioRad) for a broad range separation.
| |4% |16% |Upper Gel (sample loading) |
|Acryl-Bis mix (49.5 %T, 1.5% C) |234 |937 |273 |
|3x buffer |967 |967 |1136 |
|water |1699 |223 |2000 |
|75% glycerol |--- |773 |-- |
|10%APS |9.7 |7.6 |30 |
|TEMED |1.0 |0.8 |2.5 |
(all values in are in microliters)
After pouring of the gel the sample wells were dried with filter paper and samples were applied and carefully overlaid with the blue cathode buffer. The rest of the inner chamber was filled with the blue cathode buffer. The anode buffer already is at its place. 50 V was used for running of the gel. Once the proteins reached the separating gel the voltage was increased to 80 V or a constant current of 15 mA. After the running-front is at the middle of the separating gel, the blue cathode buffer was exchanged with a colourless one and the voltage increased to 150 V. The chamber was not washed so that a small amount of Coomassie still remained. Electrophoresis was stopped before the running front moved out of the gel.
BN PAGE Buffers
Blue Cathode Buffer (1L) Colourless Cathode Buffer
8.96g Tricine (50 mM) 8.96g Tricine (50 mM)
3.138g BisTris (15 mM) 3.138g BisTris (15 mM)
0.2g Coomasssie blue G 250 pH 7.0 at 4°C
pH 7.0 at 4°C
3X Gel Buffer (50ml) Anode Buffer (1L)
9.84g aminocaproic acid (1.5 M) 10.45g BisTris (50 mM)
1.567g BisTris (150 mM) pH 7.0 at 4°C
pH 7.0 at 4°C
Activity stain in Blue Native gels for soluble enzymes
Soluble fraction from CFE was isolated from Pseudomonas fluorescens grown in citrate (control) and Ga-citrate medium at various growth times. Samples were prepared by diluting the soluble fraction with 3X Blue Native (BN) buffer and water to a final concentration of 3mg/ml protein equivalent and 1X BN buffer (50 mM BisTris, 500 mM ε-amino-n-caproic acid, pH 7.0) respectively. To each lane 60 µg of protein (20 µl of sample) were loaded per lane and electrophoresed under Blue native conditions. Following BN-PAGE the gels were incubated in equilibration buffer (25 mM Tris-HCl, pH 7.3, 5 mM MgCl2) for 15 min. The gels were then placed in the appropriate activity buffer (equilibration buffer with the desired substrate, cofactor, and/or enzymes for coupled reactions) and incubated for various times. The activity in the gels was visualized using phenazine methosulfate (PMS) and iodonitrotetrazolium violet (INT). Enzymatic reactions that require NAD+ and/or NADP+, which is converted to NADH and NADPH respectively are easily stained within the gel using a yellow soluble tetrazolium salt (INT), which is converted to an insoluble pink substance (formazan) in the presence of the electron donor (ex: NADH). Under such condition care was taken to avoid exposing the staining solution to the light, as this will result in a high background and thus, the reactions were performed in the dark. This reaction proceeds rapidly in the presence of PMS, which acts as an intermediary catalyst; for example ICDH:
[pic]
NADPH producing enzymes: ICDH, ME, G6PDH.
The activity of these enzymes was visualized using INT. As indicated above, INT, a tetrazolium is readily reduced by NADH or NADPH in the presence of PMS to form an insoluble formazan localized at the site of enzymatic activity. The gels were placed in equilibration buffer (25 mM Tris-HCl, pH 7.3, 5 mM MgCl2) plus 0.4 mg/ml PMS, 0.4 mg/ml INT, 0.1 mM NADP+ and the following substrate depending on the respective enzyme to be detected: 1 mM isocitrate (for ICDH-NADP+ activity), 5 mM malate (for ME activity) and 5 mM glucose-6-phosphate (for G6PDH activity). The total volume of the activity buffer was 1.5 ml per lane. Upon visualization of a pink precipitate at the site of enzyme catalysis the gel(s) was placed in destaining solution (50% methanol, 10% acetic acid). This stopped the reaction and served in removing the Coomassie G 250 from the gel leaving a clear gel and pink band(s) at the site of enzyme activity.
Detection of Catalase activity in BN PAGE
The in-gel activity of catalase was visualized using P-anisidine. The gels were placed in equilibration buffer (25 mM Tris-HCl, pH 7.3, 5 mM MgCl2) plus 10mM P-anisidine and 35mM hydrogen peroxide. The total volume of the mixture was 1.5 ml/lane. The degradation product of the catalase reaction reacts with P-anisidine and a pinkish color develops where the enzyme is located. Upon visualization of a pink precipitate at the site of enzyme catalysis (60 min) the gel was scanned.
Detection of ACN activity in BN PAGE
The in-gel activity of ACN was visualized using ICDH from porcine heart. The gels were placed in equilibration buffer (25 mM Tris-HCl, pH 7.3, 5 mM MgCl2) containing 60 units of ICDH, 0.5 mM NADP+, 10 mM citrate plus 0.4 mg/ml PMS and 0.4 mg/ml INT. The total volume was 1.5 ml/lane. The band corresponding to ACN was apparent after approximately 10 min of incubation.
AST in-gel activity detection
The detection of AST was possible with the addition of malic dehydrogenase from porcine heart (160 units). The gel was incubated in a volume of 1.5 ml/ lane consisting of activity buffer, 10 mM aspartate, 10 mM α-ketoglutarate, 0.5 mM NADH, 0.4 mg/ml INT and DCPIP 0.5 mg/ml. DCPIP is the recipient of the electrons transferred during malic dehydrogenase substrate turnover and reduce INT to form a formazan precipitate where the enzyme is situated on the gel. The detection of AST was evident after 15 min of incubation. Standard with NADH, DCPIP and INT alone were also performed.
Activity stain in Blue Native gels for membrane enzymes
Following the isolation of membranes as described above, the proteins were solubilized using dodecylmaltoside: Samples were prepared by diluting the membrane fraction with 3X Blue Native (BN) buffer, 10 % dodecylmaltoside, and water to give a final concentration of 4mg/ml protein equivalent, 1X Blue Native (50 mM BisTris, 500 mM aminocaproic acid, pH 7.0), and 1% dodecylmaltoside respectively. The samples were incubated on ice for 60 min with intermittent mixing. To each lane of a mini slab gel (BioRad), 60 µg of protein were loaded unless specified otherwise. Following BN-PAGE the gels were incubated in equilibration buffer (25 mM Tris-HCl, pH 7.3, 5 mM MgCl2 ) for 15 min. In an effort to identify the nature of these enzymes, the gels were placed in equilibration buffer plus 5 mM substrate, 0.4 mg/ml PMS, 0.4 mg/ml INT, 0.5 mM NAD+ unless specified otherwise. Upon visualization of a pink precipitate at the site of enzyme catalysis the gel(s) was placed in destaining solution (50% methanol, 10% acetic acid). This stopped the reaction and served in removing the Coomassie G 250 from the gel leaving a clear gel and pink band(s) at the site of enzyme activity.
SOD in-gel activity detection
For the detection of SOD, the gel was incubated in a volume of 1.5 ml/lane consisting of activity buffer, 0.5 mg/ml INT, 15 mM menadione. The detection of SOD was evident after 12 hrs and appeared as an achromatic (colorless) band whereas the remaining gel was deeply colored. The gels were scanned as is.
α-KGDH in-gel activity detection
For the detection of α-KGDH, the gel was incubated in a volume of 1.5 ml/lane consisting of acitivity buffer, 0.1mM CoA, 0.5 mM NAD+, 5 mM α-ketoglutarate, 0.4 mg/ml of PMS and INT. The detection of the α-KGDH was evident after 20 min. To prevent a dark background, the level of Coenzyme A may be decreased.
MDH in-gel activity detection
For the detection of MDH, the gel was incubated in a volume of 1.5 ml/lane consisting of acitivity buffer, 0.5 mM NAD+, 5 mM malate, 0.4 mg/ml of PMS and INT. The detection of the MDH was evident after 20 min.
GDH in-gel activity detection
For the detection of GDH, the gel was incubated in a volume of 1.5 ml/lane consisting of acitivity buffer, 0.5 mM NAD+ , 0.4 mg/ml of PMS and INT and 5 mM glutamate. The detection of the GDH was evident after 30 min.
SDH in-gel activity detection
For the detection of SDH, the gel was incubated in a volume of 1.5ml/lane consisting of activity buffer containing 5mM KCN, 10mM succinate, and INT (0.4mg/ml). KCN increased the rate of the reaction and the appearance of the formazan precipitate. The detection of SDH was evident after approximately 10 min.
FUM in-gel activity detection
The detection of FUM was possible with the addition of malic dehydrogenase from porcine heart (160 units). The gel was incubated in a volume of 1.5ml/lane consisting of activity buffer, 10mM fumarate, 0.5mM NAD+, 0.4mg/ml PMS and INT. The detection of fumarase A was evident after 1 hr whereas the detection of fumarase C was evident after 4 hrs.
2D BN PAGE
The protein of interest (ICDH-NADP+ or α-KGDH) was first detected catalytically in the first two lanes (citrate and Ga-citrate fractions) of the first dimension BN PAGE. The corresponding band in the next two lanes were cut in the first dimension BN PAGE and inserted between the glass plates of another 4-16% polyacrylamide gel. The stacking gel was subsequently added. To avoid the formation of air pockets, the gel casting may be tilted to one side. After polymerization, the gels were runned as previously described. Enzyme activity was detected as previously described.
Protein levels
Slab-gels were fixed and stained with 10% acetic acid, 50% methanol, and 0.2% Coomassie Brilliant Blue R-250. The gels were left in the staining solution overnight. The gels were destained in a solution of 10% acetic acid and 50% methanol.
Sodium Dodecyl Sulfate Polyacrylamide Gel Electrophoresis of 2D α-KGDH
The band corresponding to α-KGDH activity as visualized by formazan precipitation was cut in the 2D BN gel, incubated in electrophoresis buffer for 30 min and inserted between two plates containing the SDS gel. The gel slabs were kept wet to facilitate the insertion between the two plates. The stacking gel was subsequently added avoiding any air pockets by tilting the gel on either side. Electrophoresis using a discontinuous buffer system was performed according to the method of Laemmli MACROBUTTON HtmlResAnchor (Laemmli, 1970), with the following modifications. The concentrations in the separation gel were ; 10% T and 0.8% C, 0.375 M Tri-HCl (pH 8.8), 0.1% SDS, 0.06% TEMED, and 0.03% APS. The concentrations in the stacking gel were; 4% T and 0.8% C, 0.1% SDS, 0.625 M Tris-HCl (pH 6.8) , 0.06% TEMED, and 0.03% APS. The electrode buffer (pH 8.3) contained 0.025 M Tris, 0.192 M glycine, and 0.1% SDS. Electrophoresis was carried out with a constant voltage of 200 V until the Coomassie blue marker from the 2D BN PAGE reached the bottom of the gel.
30% Acrylamide Stock Solution 4X Tris/SDS pH 8.8
(30% T, 0.8% C) (100ml)
1.5 M Tris Base
29.2g Acrylamide 0.4% SDS
0.8g Bisacrylamide pH adjusted to 8.8 with 11 N HCl
4X Tris/SDS pH 6.8 5X Electrophoresis Buffer (1L)
1.5 M Tris Base 15.1g Tris Base
0.4% SDS 72.0g Glycine
pH adjusted to 68 with 11 N HCl 5.0g SDS
Coomassie Blue Destaining Solution
Staining Solution
50% Methanol
50% Methanol 10% Acetic acid
10% Acetic acid
0.2% Brilliant Blue R 250
Immunoblotting of Proteins Separated by SDS-PAGE
Following SDS-PAGE, the stacking gel was removed and the orientation of the resolving gel was marked by cutting out a corner. The gel was then soaked in the protein transfer buffer for at least 10-20 min. Hybond™- P (PVDF membrane) was pre-wet by placing in 100% methanol for 10 seconds and washed in distilled water for 5 min before the membrane was equilibrated in the protein transfer buffer for at least 10 min. The electroblotting cassette was assembled according to the instructions provided by BioRad laboratories. The proteins were transferred overnight at 4 oC with a constant voltage of 20 V. The PVDF membranes were then removed and non-specific binding sites were blocked by soaking the membranes in 5% Blotto (5% skim milk in TTBS: 20 mM Tris HCl, 0.8% NaCl, 1%Tween 20, pH 7.6). Following 60 min incubation, the membranes were washed, 1x 5 min, with an excess volume of TTBS. The blot was then incubated for 60 min with the primary antibody at the optimized dilution of 1/5000 for α-KGDH in 5% Blotto. The antiserum to bovine heart PDH was raised in rabbits. The membranes were then briefly washed with excess TTBS followed by 2 x 5 min washes in the same buffer. Following 60 min incubation with the appropriate dilution of the secondary antibody (1/10 000) in 5% Blotto, the blots were washed 1 x 15 min and 4 x 5 min with excess volume of TTBS.
LIST OF BUFFERS
Protein Transfer Buffer (1L) Tris Buffered Saline (TBS) (1L)
3.03 g Tris-base 2.42 g Tris-base
14.4 g glycine 8 g NaCl
200 ml Methanol adjust pH to 7.6 with 2N HCl
store at 2-8°C store at 2-8°C
Tween Tris Buffered 5% Blotto
Saline (TTBS) 5% (w/v) dried skim milk
Dilute required volume of in TTBS
Tween™ 20 in TBS to give
A 0.1% (v/v) solution
store at 2-8°C
Chemiluminescence Detection
The detection of the desired proteins was achieved with the ECL Plus system (Amersham Pharmacia Biotech). The detection reagents, Solution A (ECL Plus substrate solution) and Solution B (Acridan solution in dioxane and ethanol) were allowed to equilibrate to room temperature. The detection solutions A and B were mixed in a ratio of 40:1 (for example, 2 ml Solution A and 50 µl Solution B) and pipetted on to the membranes, protein side up. Following 5 min incubation at room temperature, the blots were visualized with autoradiography film, Hyperfilm™ ECL (Amersham Pharmacia Biotech).
Quantification of bands
Bands were quantified using Scion Image V. 4.0.2 (Scion Corporation, USA)
RESULTS
Growth profile of P. fluorescens exposed to gallium
When P. fluorescens was stressed with 1 mM gallium in the growth media, microbial multiplication was characterized with a lengthy adaptation phase. No significant cellular growth was observed after 40 hrs of incubation. However, after reaching the stationary phase at 65 hrs, the cell yield was 500 µg/ml, a value similar to that observed in the control culture at 24 hrs of growth (Figure 24). These results suggest that the bacteria were able to adapt to the trivalent metal.
[pic]
Figure 24: Influence of Ga+3 on the growth profile of P. fluorescens. Growth profiles were performed in triplicate and the mean values were taken.
■: control culture ∆: culture with 1 mM gallium/20 µM iron
▲: culture with 1mM gallium □: control low phosphate
○: low phosphate media supplemented with 1mM gallium/20 µM iron
(Al-Aoukaty, A. et al., Gallium toxicity and adaptation in Pseudomonas fluorescens, FEMS Microbiology Letters, 92, 1992, 265-272)
It was important to determine the fate of gallium during the course of bacterial growth. Therefore the gallium content of the supernatant fraction was monitored at various stage of growth. It was observed by X-ray fluorescence spectroscopy that the bacterium internalized gallium (Figure 25). As the organism multiplied, gallium was incorporated into the cells.
[pic]
Figure 25: Gallium distribution profile during the course of P. fluorescens growth
A: Gallium in supernatant at 0 incubation time in 1mM Ga-enriched medium. B: Gallium in supernatant at logarithmic phase of growth in 1mM Ga-enriched medium. C: Gallium in supernatant at stationary phase of growth in 1mM Ga-enriched medium.
(Al Aoukaty, A. et al., Gallium toxicity and adaptation in Pseudomonas fluorescens, FEMS Microbiology Letters, 92, 1992, 265-27)
Gallium peptide
The analysis of the cell free extracts and the supernatant revealed that gallium was predominantly localized in the supernatant and the metal was associated with a hydroxyaspartate containing metabolite with a molecular mass of approximately 500 Da. 13C NMR data and chemical analysis revealed fingerprint characteristic of β-hydroxyaspartate (Figure 26).
[pic]
PPM
Figure 26: 13C NMR of Ga-metabolite isolated at stationary phase of growth and purified by Biogel P2 chromatography. (M.Santani, BSC Hons.thesis, 1992)
Gallium and oxidative stress
Thus, it became clear that although the metal affected cellular growth profile, the organism was able to survive the stress. And as gallium is known to interfere with iron metabolism and contribute to the generation of ROS, the oxidative status of the two potential targets of ROS namely proteins and lipids was monitored.
A TBARS (thiobarbituric acid reactive species) assay was performed in the membrane fraction of control and Ga-stressed cells in order to verify the prooxidative property of gallium. It was determined that the bacterial concentration of TBARS was two-fold greater in Ga-stressed cells compared to cells without the test metal. The amount of lipids that were oxidized increased over time in both control and 1 mM Ga-stressed cultures (Figure 27). However, more oxidized lipids were evident in the cells subjected to gallium.
[pic]
Figure 27: Oxidized lipids in control and 1 mM Ga-stressed P. fluorescens at various growth intervals (n=3). Values are means ± S.D.. TBARS values differ significantly from control (*) p≤0.05.
Oxidative stress marker: oxidized proteins
The oxidized proteins were visualized with the aid of DNPH and in 1 mM Ga-stressed cells an increase in oxidized proteins was recorded (Figure 28).
[pic]
Figure 28: Oxidized proteins in control and 1 mM Ga-stressed P. fluorescens at stationary phase (n=3). Values are means ± S.D.. Protein carbonyl value differs significantly from control (*) p≤0.05.
In vitro H2O2 measurement in cellular fractions exposed to gallium
As gallium has been shown to enter the bacteria with the concomitant increase in oxidized proteins and lipids, it was therefore important to determine the nature of the oxidative species involved in this process. Membrane fractions were subjected to 5mM Ga-citrate and the amount of H2O2 generated was monitored immediately with the aid of peroxidase and P-anisidine. The total amount of H2O2 recorded was 33 µmol/mg of protein equivalent of membranes utilized. On the other hand, when these same membranes were subjected to citrate as the substrate, no H2O2 was detected. This observation suggests a role of gallium in the formation of H2O2 in P. fluorescens (Figure 29).
[pic]
Figure 29: Peroxide production by the membrane fractions from P. fluorescens (n=3)
In vitro superoxide measurement in cellular fractions exposed to gallium
The ability of gallium to generate superoxide in the membrane fraction of control cells was also probed. Membrane fractions were subjected to 5mM Ga-citrate and the amount of O2∙- generated was monitored with the aid of INT. No O2∙- was recorded when Ga-citrate was incorporated with the membrane fraction of control cells. On the other hand, when these same membranes were subjected to citrate as the substrate, O2∙- was detected. This observation appears to indicate that gallium may bind O2∙- to create a more powerful oxidant in P. fluorescens (Figure 30).
[pic]
Figure 30: Superoxide production by the membrane fractions from P. fluorescens (n=3)
Catalase
Since ROS were generated as a consequence of gallium stress, it was critical to determine the ability of the organism to deal with the oxidative stress. As catalase is known to detoxify peroxide, experiments were designed in an effort to monitor this enzyme. The activity of catalase was higher in the control cultures compared to the Ga-stressed cultures. At logarithmic phase of growth, a 4-fold decrease was observed in the soluble fraction isolated from the Ga-stressed bacteria (Figure 31).
[pic]
Figure 31: Activity of catalase (H2O2 decomposition) in control and 1 mM Ga-stressed P. fluorescens (n=3). Values are means ± S.D.. Catalase activity values differ significantly from controls (*) (p≤0.05). 100% = 8.93 umol of H2O2* mg-1min-1
BN PAGE analysis of Catalase activity
This observation was further confirmed by BN PAGE analysis. In this instance, the enzyme was stained in the gel with P-anisidine. A more intense band was observed in the soluble fraction of control bacteria (Figure 32).
1 2 3 4 5 6 7 8
[pic]
[pic]
Figure 32: Panel A: BN PAGE analysis of catalase activity. Lanes 1, 2, 3 and 4 correspond to control cells harvested after 15, 20, 25 and 30 hrs respectively. Lanes 5, 6, 7 and 8 correspond to 1 mM Ga-stressed cells harvested after 55, 60, 65 and 70 hrs respectively. 50 µg of protein were loaded in each lane. Intensities were quantified using scion image softwasre. Relative areas of the bands are given.
Superoxide dismutase
Superoxide dismutase (SOD), another enzyme commonly known to be utilized in ROS defense was also probed. This enzyme mediates the conversion of O2-∙ into H2O2. The activity of this enzyme was detected by INT using menadione as the supplier of O2-∙ (Figure 33).
[pic]
Figure 33: Comparative study of SOD activity in control and in 1 mM Ga-stressed P. fluorescens (n=3). Values are means ± S.D.. SOD activity values differ significantly from controls (*) (p≤0.05).
BN PAGE analysis of SOD activity
The activity of SOD was confirmed by a BN PAGE analysis. The membrane fraction of cell free extract (CFE) from cells grown in Ga-citrate or in Ga-free media (control) was subjected to a non-denaturing BN PAGE. The activity of SOD was detected by the addition of menadione. The achromatic band was increased in Ga-stressed cells thus indicating enhanced activity. (Figure 34).
1 2 [pic]
Figure 34: BN PAGE activity of SOD in control and 1 mM Ga-stressed P. fluorescens at logarithmic growth phase.
Lane 1: cells grown in citrate medium (control). Lane 2: cells grown in Ga-citrate medium. 30 µg of protein were loaded in each lane
Thus, the data indicated that gallium is able to evoke ROS production in P. fluorescens. However, catalase, a key enzyme involved in the detoxification of H2O2 is markedly diminished while the activity of SOD experiences an increase. Hence, it was important to evaluate how the cell may be dealing with the H2O2 generated as a consequence of gallium toxicity. The membrane fraction obtained from the control cells was subjected to either 5mM citrate or 5 mM Ga-citrate in the presence of NAD+ for 60 min and the 13C NMR spectra were recorded (Figure 35). When citrate was the substrate, a peak at 32 ppm indicative of a CH2 from α-ketoglutarate was evident. In the presence of Ga-citrate, peaks attributable to succinate were discerned. However, a peak with a chemical shift of 130 ppm chacarcteristic of aconitate was present in the incubation mixture with citrate as substrate and absent when Ga-citrate was the substrate. These disparate 13C NMR clearly depicted a marked variation in citrate and Ga-citrate metabolism.
13C NMR analyses of citrate metabolism in CFE
A
[pic]
ppm
B
[pic]
ppm
C
[pic]
D ppm
[pic]
ppm
Figure 35: 13C NMR spectra of A: membrane from control cells incubated with labeled citrate and NAD+ for 1 hr B: membrane from control cells incubated with labeled Ga-citrate and NAD+ for 1 hr C: membrane from Ga-stressed cells incubated with labeled citrate and NAD+ for 1hr D: membrane from Ga-stressed cells incubated with labeled Ga-citrate for 1 hr.
Hence, the activity of various enzymes involved in cellular metabolism was monitored in control and Ga-stressed P. fluorescens with aim of identifying the metabolic network that enables the microbe to survive the production of ROS and the depletion of iron evoked by gallium stress.
Table 1: Activities of metabolic enzymes in control and Ga-stressed cells
|Enzymes |Specific activity in CFE (nmol/mg of |Specific activity in CFE (nmol/mg of |
| |protein/min) of control cells (n=3) |protein/min) of 1 mM Ga-stressed cells |
| | |(n=3) |
|Glucose-6-Phosphate phosphatase |0.016±0.002 |0.011±0.003 |
|Pyruvate carboxylase |0.01±0.0009 |0.002±0.0007 |
|Hexokinase |11.8±1.6 |13.9±0.9 |
|6-Phosphogluconate dehydrogenase |2.0±0.3 |2.6±0.5 |
|Succinate dehydrogenase |41±2.2 |38±3.1 |
|Fumarase |30±1.3 |41±4.2 |
|Malate dehydrogenase |62±4 |77±4 |
|Aspartate transaminase |108±1.5 |121±8.1 |
|Glutamate dehydrogenase |15±0.8 |18±0.5 |
|Malic enzyme |172±8 |158±16 |
|Aconitase |106±4.5 |75±2.5 |
|Isocitrate Lyase |10±2.1 |15±1.4 |
|Isocitrate dehydrogenase (NAD+) |1.47±0.23 |2.15±0.16 |
|Isocitrate dehydrogenase (NADP+) |911±76 |1248±23 |
|Glucose-6-Phosphate dehydrogenase |24±1.7 |39±4.5 |
|α-Ketoglutarate dehydrogenase |29±0.6 |10±3.5 |
|Malate synthase |84±9 |23±11 |
|Citrate Synthase |29±6 |51±8 |
|Pyruvate dehydrogenase |3.3±0.2 |2.0±0.2 |
Table 2: A comparative evaluation of various enzymatic activities in control and Ga-stressed P. fluorescens.
|Enzymes | Variation compared to control cells |
|6-Phosphogluconate dehydrogenase |↑30% |
|Fumarase |↑37% |
|Aconitase |↓30% |
|Isocitrate Lyase |↑50% |
|Isocitrate dehydrogenase (NAD+) |↑45% |
|Isocitrate dehydrogenase (NADP+) |↑64% |
|Glucose-6-Phosphate dehydrogenase |↑63% |
|α-Ketoglutarate dehydrogenase |3 X ↓ |
|Malate synthase |3 X ↓ |
|Citrate Synthase |↑76% |
|Pyruvate dehydrogenase |↓40% |
*** control is taken as 100% or 1
ICDH-NADP+
In many organisms, an oxidative environment leads to the induction of enzymes that are capable of generating NADPH. Since the enzyme catalase was inhibited by gallium, it was essential to determine if NADPH generating enzymes were involved in the detoxification of ROS. NADPH is an essential cofactor in enzymes such as glutathione peroxidase, catalase and glutathione. Hence, studies were designed to investigate the role of ME, G6PDH and ICDH. This latter enzyme was found to be significantly increased in Ga-stressed cells. This NADPH producing enzyme is also important in the metabolism of citrate, the sole source of carbon in this study. The ICDH localized in the cytoplasm was dependent on NADP+. The activity of this enzyme was first determined by monitoring the disappearance of NADP+ at 340 nm, a cofactor necessary for the conversion of isocitrate to α-ketoglutarate (Figure 36).
[pic]
Figure 36: Rate of reaction illustrating the appearance of NADPH when isocitrate is the substrate in control and in 1 mM Ga-stressed soluble fraction.
It became important to examine the activity of the soluble ICDH-NADP+ at various growth intervals in control and 1 mM Ga-stressed cells. The activity of this enzyme was higher in Ga-stressed bacteria compared to control bacteria (Figure 37).
[pic]
Figure 37: Specific activity of ICDH-NADP+ enzyme in control and in 1 mM Ga-stressed cells at various growth intervals (n=3). Values are means ± S.D.. ICDH-NADP+ differ significantly from controls (*) (p≤0.05).
BN PAGE analysis of ICDH-NADP+
BN PAGE was utilized to determine the activity of the soluble ICDH-NADP+. The soluble fraction of cell free extract (CFE) from cells grown in Ga-citrate or in control media obtained at various intervals of growth was subjected to a non-denaturing BN PAGE. The activity of ICDH-NADP+ was detected by the addition of isocitrate as well as the cofactor NADP+. The NADPH produced help precipitate INT. The activity of this enzyme was markedly increased in Ga-stressed bacteria and an isoenzyme was also detected. This isoenzyme (A) was more prominent in 1 mM Ga-stressed cells (Figure 38).
1 2 3 4 5 6
[pic]
Band A
[pic]
Band B
[pic]
Figure 38: BN PAGE analysis of ICDH-NADP+ activity in control and 1 mM Ga-stressed P. fluorescens at various growth intervals.
Lane 1,2,3: cells grown in citrate medium (control) for 15, 25 and 30 hrs respectively. Lane 4,5,6: cells grown in Ga-citrate medium for 55, 65 and 70 hrs respectively. 30 µg of protein were loaded in each lane. Intensities were quantified using scion image software. Relative areas of the bands are given.
2D BN PAGE analysis of ICDH-NADP+
The enzyme ICDH-NADP+ (B) was further studied by two dimension BN PAGE analysis. The activity of this enzyme was markedly increased in 1 mM Ga-stressed bacteria (Figure 39).
1 2
[pic]
Figure 39: 2D BN PAGE analysis of ICDH-NADP+ activity in control and 1 mM Ga-stressed cells.
Lane 1: cells obtained in citrate medium (control). Lane 2: cells obtained in Ga-citrate medium. 60 µg were loaded in each lane.
Modulation of ICDH-NADP+ activity: influence of ROS
To determine the nature of the expression of ICDH-NADP+ and its influence on ROS, Ga-stressed cells were subjected to control media supplemented with H2O2 or menadione. In the H2O2 and menadione supplemented medium, the ICDH-NADP+ activity was as high as in the Ga-stressed cultures, while in the control media, a decline in activity was observed (Figure 40).
[pic]
Figure 40: Influence of different effectors on ICDH-NADP+ activity in P. fluorescens (n=3).
(A) Cells grown in 1 mM Ga-citrate for 65 hrs. 10 mg of these whole cells were transferred for 6 hrs to media B (control media), C (control media + menadione) D (control media + H2O2). The specific activity in A is the mean ± S.D. of three independent experiments to which are compared the specific activity of B, C and D. ICDH-NADP+ activity values differ significantly from control (*) (p≤0.05). 100%= 1250 nmol/mg of protein/min
BN PAGE of ICDH-NADP+ regulation
This regulatory pathway was also confirmed by BN PAGE. It was determined that the activity of this enzyme remained high in gallium cells transferred to a control media supplemented with 15 mM H2O2. The isoenzyme A was also higher (Figure 41).
1 2 3 4 5
[pic]
Figure 41: BN PAGE analysis of ICDH-NADP+ activity in various media. Influence of H2O2 in the growth media
Lane 1: cells grown in citrate medium (control). Lane 2: cells grown in Ga-citrate medium. Lane 3: 1 mM Ga-stressed cells transferred to control media for 6 hrs. Lane 4: 1 mM Ga-stressed cells transferred to control media containing 1 mM menadione for 6 hrs. Lane 5: 1 mM Ga-stressed cells transferred to control media containing 15 mM H2O2 for 6 hrs. 30 µg of protein were loaded in each lane.
Malic enzyme (ME)
This enzyme is involved in the generation of pyruvate and NADPH from the substrate malate. Kinetic studies showed that the enzyme was slightly slower in the Ga-stressed cells compared to control cells (Table 1). To further investigate the properties of this enzyme, a BN PAGE analysis was performed from the soluble fraction from the control and Ga-stressed cells (Figure 42).
1 2 3
[pic]
Figure 42: BN PAGE analysis of ME activity in the soluble fraction. Lane 1: control cells at stationary phase (24 hrs). Lane 2: 1 mM Ga-stressed cells at stationary phase (65 hrs). Lane 3: 1 mM Ga-stressed cells introduced in a control medium for 6 hrs. 60 µg of protein were loaded in each lane.
Glucose 6-Phosphate dehydrogenase
Amongst the enzymes that are induced in situations of oxidative stress is G6PDH. This enzyme is involved in the pentose phosphate pathway and generates NADPH for various enzymatic and nonenzymatic antioxidative systems. In P. fluorescens subjected to 1mM gallium, a 60% increase in G6PDH activity is observed compared to control cells (Table 1). This observation was further confirmed with a BN PAGE analysis of the soluble fraction of control and Ga-stressed bacteria obtained at logarithmic phase of growth (Figure 43).
1 2
[pic]
Figure 43: BN PAGE analysis of G6PDH activity in the soluble fraction. Lane 1: control cells at stationary phase (24 hrs). Lane 2: 1 mM Ga-stressed cells at stationary phase (65 hrs). 60 µg of protein were loaded in each lane.
ICDH-NAD+
In biological systems, the synthesis of α-ketoglutarate is also mediated by ICDH-NAD+. This enzyme is localized in the membrane and generates NADH for ATP production. A DNPH assay was performed in order to measure the amount of α-ketoglutarate generated by this enzyme in control and Ga-stressed cells. It was determined that the activity of this enzyme in Ga-stressed bacteria was increased 40% compared to control (Figure 44). This increase in activity was dependent on the gallium present in the media.
[pic]
Figure 44: Influence of gallium on the ICDH-NAD+ specific activities (formation of α-ketoglutarate) in control, 1 mM Ga-stressed bacteria and 1 mM Ga-stressed cells transferred to control media for 6 hrs (n=3). Values are means ± S.D.. ICDH-NAD+ differ significantly from control (*) (p≤0.05).
α-KGDH
It appeared that when P. fluorescens was grown in a media containing 1 mM gallium, the membrane and soluble form of ICDH were upregulated in an effort to increase the generation of α-ketoglutarate. Hence, to elucidate the biochemical significance associated with high levels of α-ketoglutarate in the Ga-stressed cells, it was important to determine the fate of α-ketoglutarate in this system
Of all the tricarboxylic acid cycle enzymes studied, α-KGDH exhibited the most variation between control and Ga-grown cells. The study of this enzyme was therefore crucial since its activity was markedly diminished (p≤0.05) in Ga-stressed P. fluorescens. At least three fold decrease in activity was observed in bacteria cultured in the gallium medium compared to those isolated from the control medium at identical phase of growth (Figure 45).
[pic]
Figure 45: α-KGDH specific activities (decomposition of α-ketoglutarate) in control and 1 mM Ga-stressed cells at logarithmic phase of growth (n=3) p≤0.05.
The increase activity of ICDH-NADP+ and the expression of a novel isoenzyme suggest that the metabolism of the organism stressed with 1mM gallium is geared towards the generation of α-ketoglutarate. However, the activity of the downstream enzyme α-KGDH was significantly diminished. The 13C NMR data pointed to the decomposition of α-ketoglutarate into succinate when Ga-citrate was the substrate. It was therefore essential to determine if P. fluorescens subjected to 1 mM gallium expressed a novel α-ketoglutarate metabolizing enzyme. Thus, this enzyme was assayed in the presence of different co-factors in the Ga-stressed membrane cell free extracts. It became evident that α-KGDH was the only discernable enzyme capable of utilizing α-ketoglutarate.
Table 3: Influence of various cofactors on α-ketoglutarate metabolism
|Substrate |Cofactor(s) |Specific activity (nmol/mg of protein/min) n=3 |
| | | |
|α-Ketoglutarate |NIL |NIL |
|α-Ketoglutarate |NAD+ |NIL |
|α-Ketoglutarate |NAD+ + CoA |10±3.6 |
|α-Ketoglutarate |NAD+ + CoA + TPP |11±2.6 |
|α-Ketoglutarate |NADH |NIL |
Confirmation of the decreased activity of α-KGDH in the membrane fraction of CFE of control and Ga-stressed P fluorescens.
1H NMR was utilized to determine the presence/absence of α-KGDH activity in control and Ga-stressed membrane fraction. When coenzyme A and NAD+ were added in conjunction with α-ketoglutarate in the membrane fraction of control cells, a peak at 1.98 ppm indicative of succinate was discernible whereas the peak corresponding to α-ketoglutarate were absent (Figure 46A). However, when the same substrates/cofactors were added to the membrane fraction of Ga-stressed membrane fraction, only peaks corresponding to α-ketoglutarate are evident, thus confirming the decreased α-KGDH activity in the gallium cell free extract (Figure 46B).
A
[pic]
ppm
B
[pic]
ppm
Figure 46: Panel A: 1H NMR of α-KGDH activity. Membrane fraction from control cell free extract (0.5mg protein equivalent) was incubated with α-ketoglutarate (2mM), Coenzyme A (0.1mM) and NAD+ (0.1mM) for 30 min (Note: the succinate peak). Panel B: 1H NMR of α-KGDH. Membrane fraction from Ga-citrate cell free extract (0.5mg protein equivalent) was incubated with α-ketoglutarate (2mM), Coenzyme A (0.1mM) and NAD+ (0.1mM) for 30 min (Note: the α-ketoglutarate peak)
Expression of α-KGDH in control and Ga-stressed P. fluorescens
BN PAGE analysis
The membrane fractions of the cell free extract (CFE) from cells grown in Ga-citrate or in Ga-free media (control) were subjected to a non-denaturing BN PAGE. The α-KGDH activity was detected due to its ability to produce NADH upon incubation with nicotinamide (NAD+) and Coenzyme A. The formation of NADH was visualized by the precipitation of formazan. The band corresponding to the enzymatic activity was markedly intense in control cells (Figure 47).
1 2
[pic]
Figure 47: BN PAGE detection of α-KGDH activity in control and 1 mM Ga-stressed cells
Lane 1: cells grown in citrate medium (control). Lane 2: cells grown in Ga-citrate medium. 60 µg of protein were loaded in each lane
2D BN PAGE analysis of α-KGDH
Subsequently, a two dimension BN PAGE was performed on the band corresponding to α-KGDH. The α-KGDH activity was once again visualized by the precipitation of formazan. The activity of α-KGDH was decreased in Ga-citrate grown bacteria. The intensity of this protein was visualized by Coomassie blue staining and was more abundant in the cells grown in citrate (Figure 48).
A 1 2
[pic]
B
[pic]
Figure 48: Panel A: 2D BN PAGE activity staining of α-KGDH: Lane 1: cells grown in citrate medium. Lane 2: cells grown in Ga-citrate medium. Panel B: Coomassie Blue staining
Western Blot analysis of α-KGDH
Western Blot analysis was performed on the blot corresponding to the E2 subunit of α-KGDH. The enzyme α-KGDH was visualized with autoradiography film, Hyperfilm™ ECL. The amount of protein was decreased in Ga-citrate grown bacteria (Figure 49).
1 2
[pic]
Figure 49: Western Blot analysis of the E2 subunit of α-KGDH obtained from control cells (lane 1) and 1 mM Ga-stressed cells (lane 2). Intensities were quantified using scion image software. Relative areas of the bands are given.
α-KGDH activity in P. fluorescens at various stages of growth in citrate and Ga-citrate medium.
The ability to selectively decrease the activity of α-KGDH enabled this organism to survive milimolar amounts of gallium. This enzyme had low activity throughout the growth phase of Ga-stressed bacteria. On the other hand, this same enzyme was unaffected and active in control cells throughout the incubation period analyzed (Figure 50).
[pic]
Figure 50: α-KGDH specific activities (α-ketoglutarate decomposition) in control and 1 mM Ga-stressed bacteria harvested at various growth stages and 1 mM Ga-stressed cells transferred to a control medium for 6 hrs (n=3) Values are means ± S.D.. α-KGDH values differ significantly from controls (*) (p≤0.05)
BN PAGE of α-KGDH activity in P. fluorescens at various stages of growth in citrate and Ga-supplemented citrate medium.
The purpose of this experiment was to confirm and visualize the decrease in α-KGDH activity along different stages of growth. The membrane fractions of cell free extract from citrate and Ga-citrate grown bacteria were subjected to BN PAGE. This procedure allowed visualizing the activity of α-KGDH at different stages of growth (Figure 51). These results confirmed the data obtained in the spectrophotometric assay utilized to determine the activity of α-KGDH.
1 2 3 4 5 6 7 8
[pic]
Figure 51: BN PAGE of α-KGDH activity of citrate and Ga-citrate grown bacteria at various intervals of growth. Lanes 1,2,3,4 correspond to control cells harvested after 15, 20, 25 and 30 hrs respectively. Lanes 5,6,7,8 correspond to 1 mM Ga-stressed cells harvested after 55, 60, 65 and 70 hrs respectively.
α-KGDH activity in various metal stressed media
It became important to determine whether this decrease in α-KGDH was specific to the gallium stress and its ability to generate ROS. Therefore, it was important to measure the activity of this enzyme in other systems containing different metal stress. Calcium, a nonoxidative metal, aluminum and iron known for their prooxidant properties were utilized as stressors. When the bacterium was faced with calcium, α-KGDH was not inhibited. A significant reduction was observed with aluminum or iron.
Table 4: α-KGDH activity in P. fluorescens cultures grown in media supplemented with various metals. Membrane fraction from cell free extracts (0.2mg protein equivalent was incubated for 10 min with 0.3 mM α-ketoglutarate, 0.5mM Coenzyme A and 0.5mM NAD+. The utilization of α-ketoglutarate was monitored by the DNPH assay.
|Metal |Specific activity (nmol*min-1*mg protein-1 (n=3) |
| | |
|Control (no metal) |28.7±0.6 |
|1 mM Ga |10.0±3.6 |
|0.1 mM Ga |11.2±2.7 |
|1 mM Ga/20 µM Fe |5.1±3.8 |
|1 mM Ga/100 µM Fe |3.5±0.9 |
|1 mM Ca |27.1±1.1 |
|15 mM Al |15.1±0.39 |
Modulation of α-KGDH by Ga(NO3)3
The aforementioned results appeared to indicate that α-KGDH was in fact inhibited by gallium and other prooxidative metals. To evaluate the direct or indirect inhibition of α-KGDH activities, the control membrane fraction was subjected to various amount of gallium for 15 min and the specific activity of α-KGDH was determined. It was demonstrated that 500 µM of gallium inhibited α-KGDH by at least two-fold (Figure 52). This in vitro evidence substantiated an involvement of gallium either directly or indirectly in the decrease of α-KGDH activity.
[pic]
Figure 52: Modulation of α-KGDH specific activities (decomposition of α-ketoglutarate) by Ga(NO3)3 in the membrane fraction of control cells (n=3). Values are means ± S.D.. α-KGDH value differ significantly from control (*) (p≤0.05).
Fate of α-ketoglutarate in Ga-stressed P. fluorescens
The decreased activity of α-KGDH in Ga-stressed cells appeared to be necessary for the bacteria to adapt and survive. However, α-ketoglutarate was not being utilized in the tricarboxylic acid cycle as α-KGDH was downregulated. Thus it became important to decipher the apparent build-up of this α-ketoacid. Therefore, different enzymes other than the membrane associated α-KGDH capable of metabolizing α-ketoglutarate were studied (Table 5). Evidently, no enzyme was drastically increased in Ga-stressed cells that would be indicative of an increased utilization of α-ketoglutarate. Hence, the notion of a pool of α-ketoglutarate that might be involved in a non-enzymatic fashion became attractive.
Table 5: α-ketoglutarate utilizing enzymes in control and Ga-stressed cells (n=3)
|Enzymes |Specific activity (nmol*min-1*mg protein-1) |
| |Control cultures |Ga-stressed cultures |
| | | |
|α-ketoglutarate dehydrogenaseb |Nil |Nil |
|α-ketoglutarate reductasea |Nil |Nil |
|α-ketoglutarate reductaseb |Nil |Nil |
|CoA independent α-ketoglutarate dehydrogenaseb |Nil |Nil |
|CoA independent α-ketoglutarate dehydrogenasea |Nil |Nil |
|Glutamate dehydrogenasea |14.6±0.8 |17.5±0.5 |
|Glutamate dehydrogenaseb |nil |Nil |
|Aspartate transaminaseb |108±1.5 |122±8.1 |
a = membrane fraction of cell free extract
b = soluble fraction of cell free extract
Modulation of α-KGDH activity: influence of ROS
Not only did gallium inhibit the enzyme α-KGDH but this metal also generated H2O2 in P. fluorescens. It was therefore important to determine if the inhibition of this enzyme was an important strategy to survive the oxidative condition. Therefore, 1 mM Ga-stressed cells were grown to stationary phase where the specific activity of α-KGDH is low (10±3.6). These cells (10mg) were then reintroduced in a gallium media for 6 hrs, a control medium devoid of metal for 6 hrs and a control medium containing 1 mM menadione and a control medium containing 15mM H2O2 for 6 hrs. The specific activity of this enzyme obtained by the DNPH assay was markedly decreased in gallium, H2O2 and menadione containing media. There was 60% decrease in α-KGDH activity in the cells subjected to menadione, thus indicating a link between the enzyme and ROS tolerance (Figure 53).
[pic]
Figure 53: Regulation of α-KGDH specific activities (decomposition of α-ketoglutarate) in various media
Gallium cells (A) were grown for 65 hrs. 10 mg of these whole cells were transferred for 6 hrs to media B (control media), C (control media + H2O2), D (control media + menadione) (n=3). Values are means ± S.D.. α-KGDH values differ significantly from control (*) (p≤0.05).
BN PAGE analysis of α-KGDH in cells stressed with ROS
The membrane fractions of cell free extract (CFE) from cells grown in various media were subjected to a non-denaturing BN PAGE. The α-KGDH activity was detected by formazan precipitation. The band corresponding to the enzymatic activity was markedly intense in control cells compared to cells grown in a control culture containing 1mM menadione (Figure 54).
1 2 3 4
[pic]
[pic]
Figure 54: BN PAGE analysis of α-KGDH activity as a function of ROS.
Lane 1: control membrane fraction Lane 2: membrane from 1 mM Ga-stressed cells Lane 3: membrane from 1 mM Ga-stressed cells introduced in a control medium for 6hrs Lane 4: membrane from 1 mM Ga-stressed cells introduced in a control medium containing 1mM menadione for 6 hrs. Intensities were quantitated using scion image software. Relative areas of the bands are given.
Another attractive possibility for the fate of α-ketoglutarate is its ability to scavenge H2O2 in vivo. It was shown that more reactive oxygen species like H2O2 were generated in Ga-stressed cells either due to the increase in lipid/protein oxidation or the increase in labile iron. And, as the enzyme catalase was inhibited in Ga-stressed bacteria, the organism must utilize an alternative to detoxify H2O2. Could the inactivation of α-KGDH serve as a means to increase the pool of α-ketoglutarate in order to rid the cell of H2O2? This might explain the non enzymatic conversion of α-ketoglutarate to succinate observed in the NMR experiment when labeled Ga-citrate was the substrate. Indeed, in vitro experiments did show the ability of α-ketoglutarate to scavenge H2O2 (Figure 55).
[pic]
Figure 55: The decomposition of H2O2 by α-ketoglutarate. Values are means ± S.D..
The ability of α-ketoglutarate to effectively scavenge H2O2 was confirmed by NMR. When α-ketoglutarate (5 distinct peaks) was allowed to react with H2O2, the result was the appearance of two peaks indicative of succinate (Figure 56).
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Figure 56: Panel A: 13C NMR spectra corresponding to α-ketoglutarate. Panel B: 13C NMR spectra corresponding to succinate. Panel C: 13C NMR spectra depicting the reaction between α-ketoglutarate and H2O2.
Influence of gallium on the iron homeostasis in P. fluorescens
It has widely been reported in literature that gallium can substitute for iron in iron containing proteins. In fact, the therapeutical feature of gallium is due to its ability to mimic iron. The liberation of iron from proteins would greatly increase the labile iron pool and explain the rise in oxidative species as well as the inhibition of iron containing enzymes. It was therefore important to determine the integrity of iron proteins in Ga-stressed bacteria. Evidently, the integrity of iron in iron-containing proteins attributed to the band in the 395-415 nm region was markedly perturbed in the Ga-stressed cytoplasm (Figure 57)
A
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B
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Figure 57: Detection of iron sulfur (Fe-S) clusters in; A: control soluble fraction and B: 1 mM Ga-stressed soluble fraction
Iron-deprivation induced by Ga-stress
Aconitase (ACN)
If in fact gallium is able to displace iron in biomolecules, iron containing enzymes such as ACN, FUM, SDH and catalase must be affected by the trivalent metal. Indeed, catalase was markedly lower even though the level of H2O2 was higher in Ga-stressed cells. The first enzyme that was assayed was ACN. In numerous organisms, ACN serves as an iron sensor that is inhibited when the levels of intracellular iron are low. Trivalent metal ions such as Ga+3 may also displace the iron constituting the iron sulfur cluster and inhibit the enzyme. It was determined by kinetic reaction that the activity of the enzyme ACN is decreased significantly in Ga-stressed cells. An experiment was also performed in order to determine if gallium was the real causative agent in the observed decreased activity of ACN (Figure 58). The result demonstrated that when gallium cells were reintroduced in the control media for 6 hrs, the specific activity of ACN returned back to control level.
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Figure 58: Specific activities of ACN (cis-aconitate formation) in control cells, 1 mM Ga-stressed cells and 1 mM Ga-stressed cells transferred to a control medium (n=3) Values are means ± S.D.. Aconitase value differs significantly from control (*) (p≤0.05)
BN PAGE analysis of ACN
The soluble fractions of cell free extract (CFE) from cells grown in Ga-citrate or in Ga-free media (control) were subjected to a non-denaturing BN PAGE. The ACN activity was detected by monitoring isocitrate formation from citrate. The NADPH generated by ICDH-NADP+ was visualized by formazan precipitation. The band corresponding to the enzymatic activity was more intense in control cells (Figure 59).
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Figure 59: BN PAGE detection of ACN in control and 1 mM Ga-stressed soluble fraction.
Lane 1: Cells grown in citrate medium (control). Lane 2: Cells grown in Ga-citrate medium. 60 µg of protein were loaded in each lane.
ACN activity as a function of ROS in the growth media
Not only does gallium inhibit the enzyme ACN but this metal also generates H2O2 in P. fluorescens. It was therefore important to determine if the inhibition of this enzyme was an important consequence of the oxidative environment. Therefore, gallium cells were grown till stationary phase where the activity of ACN was low (75 ± 2.5nmol/mg of protein/min). These cells (10mg) were then reintroduced in a gallium media for 6hrs, a control media devoid of metal for 6hrs and a control medium containing 1 mM menadione, a superoxide generating agent for 6 hrs. The specific activity of this enzyme obtained from a kinetic assay in these various media showed lower activities in gallium, H2O2 and menadione media (Figure 60). The difference between the specific activity of ACN in control medium and medium with menadione was 40% thus pointing to the inhibition of ACN as a consequence of ROS.
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Figure 60: ACN specific activity (formation of aconitate) in various media (n=3). Gallium cells (A) were grown for 65 hrs. 10 mg of these whole cells were transferred for 6 hrs to media B (control), C (control + H2O2), D (control + menadione). Values are means ± S.D.. Aconitase values differ significantly from control (*) (p≤0.05)
Fumarase (FUM)
Another iron containing enzyme involved in the TCA cycle is fumarase. The specific activity of this enzyme determined via a DNPH assay was found to be increased by 40% in Ga-stressed bacteria compared to control cells (Figure 61).
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Figure 61: Specific activities of FUM (oxaloacetate formation) in control cells, 1 mM Ga-stressed cells and 1 mM Ga-stressed cells transferred to a control medium for 6 hrs (n=3). Values are means ± S.D.. Fumarase value differs significantly from control (*) (p≤0.05)
BN PAGE analysis of FUM
The membrane fractions of cell free extract (CFE) from cells grown in Ga-citrate or in Ga-free media (control) were subjected to a non-denaturing BN PAGE. The activity of FUM was detected with the aid of malate dehydrogenase (MDH) from porcine heart. The formation of NADH was visualized by PMS and INT. The appearance of two bands suggested the induction of an isoenzyme that was only slightly expressed in the membrane fraction of control cells (Figure 62). The induction of this isoenzyme (A) would explain the increase specific activity detected with the DNPH assay in the Ga-stressed cells.
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Figure 62: BN PAGE detection of FUM in control and Ga-stressed cells. Lane 1,2: control membrane fraction obtained at 25 and 30 hrs of growth respectively. Lane 3,4: membrane fraction from 1 mM Ga-stressed cells obtained at 65 and 70 hrs of growth respectively.
Expression of the FUM (FUM C) isoenzyme as a function of ROS in the growth media
Gallium was shown to inhibit the activity of iron containing enzymes such as ACN and catalase but this metal also generates ROS in P. fluorescens. It was therefore important to determine if the expression of FUM C was due to the oxidative insult. Therefore, control cells were grown to stationary phase for 24 hrs. 10 mg of these cells were transferred to a control medium for 6 hrs and utilized as a critical control. Gallium cells were grown to stationary phase (65 hrs). These cells (10mg) were then reintroduced in a gallium media, a control media devoid of metal, a control media containing 1 mM menadione, and in a control media supplemented with 15mM H2O2 for 6 hrs. The visualization of the non-iron containing FUM C isoenzyme was accomplished with BN PAGE analysis. The appearance of the iron independent isoenzyme in H2O2 and menadione stressed cells suggests that this isoenzyme was expressed as a consequence of gallium and ROS stress respectively (Figure 63).
1 2 3 4 5
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Figure 63: BN PAGE analysis of the expression of the iron independent FUM C as a function of ROS in the growth media
Lane 1: cells grown in citrate medium (control). Lane 2: cells grown in Ga-citrate medium. Lane 3: 1 mM Ga-stressed cells introduced in control media for 6 hrs. Lane 4: 1 mM Ga-stressed cells introduced in control media containing 1 mM menadione for 6 hrs. Lane 5: 1 mM Ga-stressed cells introduced in control media containing 15 mM H2O2 for 6 hrs. 30 ug of protein were loaded in each lane.
Succinate dehydrogenase (SDH)
SDH is another iron containing enzyme involved in the TCA cycle. This enzyme is responsible for the conversion of succinate to fumarate. This enzyme is also involved in the production of energy by generating the reducing agent FADH2 for the oxidative phosphorylation. However, the specific activity of this enzyme determined with the DCPIP assay was found to be slightly higher in control bacteria compared to Ga-stressed cells (Figure 64).
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Figure 64: Specific activity of SDH (FADH2 formation) in control and 1 mM Ga-stressed P. fluorescens at stationary phase (n=3). Values are means ± S.D.. SDH value does not differ significantly from control (p≥0.05).
BN PAGE analysis of SDH
The membrane fractions from cells grown in Ga-citrate or in Ga-free media (control) were subjected to a non-denaturing BN PAGE. The SDH activity was visualized as FADH2 donated its electrons to INT. The band corresponding to the enzymatic activity was more intense in control cells (Figure 65).
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Figure 65: BN PAGE detection of SDH in the membrane fraction of control and Ga-stressed cells: Lane 1: cells grown in citrate medium (control). Lane 2: cells grown in Ga-citrate medium. 30 µg of protein were loaded in each lane
β-hydroxyaspartate residue
NMR evidence showed that gallium was detoxified via a β-hydroxyaspartate residue. These residues are often found in iron siderophores produced by microorganisms to acquire iron from their environment. It was important to determine how the reconfiguration of the cellular metabolism was providing the necessary precursors to generate this moiety.
Aspartate transaminase (AST)
In biological system, the synthesis of aspartate depends of AST, an enzyme responsible for the transamination of oxaloacetate and glutamate to aspartate and α-ketoglutarate. The activity of this enzyme was enhanced in Ga-stressed cells (121 ± 8 nmol/mg of protein/min) compared to control cells (108 ± 2 nmol/mg of protein/min) (Figure 66).
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Figure 66: Specific activities of AST (disappearance of NADH) in control and 1 mM Ga-stressed P. fluorescens at stationary phase (n=3) Values are means ± S.D.. AST value does not differ significantly from control (p≥0.05)
BN PAGE analysis of AST
The soluble fractions of cell free extract (CFE) from cells grown in Ga-citrate or in Ga-free media (control) were subjected to a non-denaturing BN PAGE. The activity of AST was detected with the aid of the enzyme malate dehydrogenase (MDH) When α-ketoglutarate is transaminated to aspartate by glutamate, the oxaloacetate produced is converted to malate (Figure 67).
1 2 3 4 5 6
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Figure 67: BN PAGE detection of AST in control and Ga-stressed cells at different intervals of growth
Lane 1, 3, 5 are control cells grown for 20, 25 and 30 hrs respectively
Lane 2, 4, 6 are 1 mM Ga-stressed cells grown for 60, 65 and 70 hrs respectively
Malate dehydrogenase (MDH)
This enzyme is responsible for the generation of oxaloacetate in biological systems. In Ga-stressed bacteria, the synthesis of a β-hydroxyaspartate may necessitate oxaloacetate, a metabolite that can be transaminated to aspartate in the presence of glutamate. Since the activity of ME was reduced in Ga-stressed cells, it was important to determine if malate was preferentially destined to oxaloacetate rather than pyruvate in order to synthesize the β-hydroxyaspartate residue. A DNPH assay was performed to determine the specific activity of MDH in control and Ga-stressed bacteria. It was determined that the activity of this enzyme was 20% higher in Ga-stressed cells compared to control cells (Figure 68). The increased activity could allow the metal stressed cells to generate a greater amount of aspartate for the detoxification of gallium
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Figure 68: Specific activities of MDH (oxaloacetate formation) in control and 1 mM Ga-stressed P. fluorescens at stationary phase (n=3) Values are means ± S.D.. MDH value differs significantly from control (p≤0.05).
BN PAGE analysis of MDH
The membrane fractions of cell free extract (CFE) from cells grown in Ga-citrate or in Ga-free media (control) as well as the membrane fraction of cell free extract from cells grown in a gallium media and transferred to control media devoid of metal for 6hrs were subjected to a non-denaturing BN PAGE. The MDH activity was detected due to its ability to produce NADH upon incubation with nicotinamide (NAD+) and the substrate malate. The formation of NADH was visualized by PMS and INT. The band corresponding to the enzymatic activity was markedly intense in Ga-stressed cells (Figure 69).
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Figure 69: In-gel activity of MDH by BN PAGE
Lane 1: cells grown in citrate medium (control). Lane 2: cells grown in Ga-citrate medium. Lane 3: 1 mM Ga-stressed cells transferred to control media for 6 hrs.
BN PAGE analysis of Glutamate dehydrogenase (GDH)
The synthesis of aspartate in biological systems also depends on the availability of glutamate. Therefore, the activity of GDH was determined by a DNPH assay in control cells (15 ± 1.1) and Ga-stressed cells (18 ± 1.5) at the stationary phase (Table 1). The slight increase in activity was also confirmed via BN PAGE analysis. The membrane fractions of the cell free extract (CFE) from cells grown in Ga-citrate or in Ga-free media (control) were subjected to a non-denaturing BN PAGE. The formation of NADH was visualized by PMS and INT. The band corresponding to the enzymatic activity was more intense in Ga-stressed cells (Figure 70)
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Figure 70: BN PAGE detection of GDH activity in control and Ga-stressed cells.
Lane 1: cells grown in citrate medium (control). Lane 2: cells grown in Ga-citrate medium.
DISCUSSION
The aforementioned results clearly indicate that gallium triggers ROS stress in P. fluorescens and the perturbations in iron metabolism are either caused or a consequence of the increase in oxidative damage. Hence, the homeostasis of iron and oxidative species are closely intertwined in organisms subjected to gallium toxicity.
Figure 71: Possible biochemical interaction of gallium in P. fluorescens
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Gallium was shown to increase the amount of oxidized lipids and the amount of oxidized proteins. The trivalent metal may interact directly with the membranes and increase the susceptibility of lipids for oxidation (Figure 72) (1). The increase in lipid peroxidation may also be driven by iron-dependent ROS formation since gallium increases the amount of labile iron that can generate free radicals (Kruszewski, M., 2003). It is known that when mitochondria are exposed to iron, they undergo lipid peroxidation (Gogvadze, V, 2002.). The hydroxyl radical generated when free iron reacts with peroxide gives rise to primary lipid peroxide that can trigger various stages of radical formation (2). In another scenario gallium may act as a prooxidant through the formation of a superoxide-metal complex (Exley, C., 2004). The latter would increase the iron driven oxidation of the lipid membrane and would also favor the reduction of Fe+3 to Fe+2 thus allowing the generation of the highly damaging hydroxyl radical (3). The ROS stress created by gallium is also evident by the inhibition of catalase (4). This enzyme is a heme containing enzyme and depends on NADPH to be in the active conformation. In a situation of ROS stress, the NADPH levels are greatly diminished thus inhibiting the enzyme. Peroxide and other oxidative species can also liberate iron from catalase rendering it inactive. Alternatively, gallium may also substitute for iron in catalase and inhibit the enzyme. Gallium was also shown to generate either directly or indirectly H2O2 in P. fluorescens. When Ga-citrate was incubated with the membrane fraction of bacteria grown in control conditions, the production of peroxide was detected. However, when citrate alone was allowed to react with the membrane fraction of control bacteria no H2O2 was discernable. Thus, it is not inconceivable that Ga-stressed cells subjected to elevated ROS and consequently, the organism will have to adapt to this stress if it is to survive.
Figure 72: Possible mechanisms of ROS generation by gallium
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Metabolic shift operative in Ga-stressed P. fluorescens
Metabolic shift is an important strategy the cell invokes in an effort to nullify this oxidative environment. An important metabolic rearrangement in P. fluorescens subjected to 1 mM gallium appears to be geared towards the formation of a pool of α-ketoglutarate. Indeed, a novel ICDH-NADP+ enzyme is expressed in Ga-stressed P. fluorescens. The activity of the wild type ICDH-NADP+ enzyme is also increased. Concomitantly, the activity of α-KGDH is reduced in these cells. The enzyme ICDH-NADP+ may be more ubiquitous then previously reported (Contreras-Shannon, V., 2004). The family of ICDH-NADP+ isoenzyme in yeast allows the formation of the products, α-ketoglutarate and NADPH in the mitochondria, cytosol and peroxisome. Hence, α-ketoglutarate is not a metabolite restricted to the mitochondria. Strikingly, the soluble ICDH-NADP+ (cytosol) was shown to be critical in detoxifying H2O2 in situations of oxidative stress in yeast (Contreras-Shannon, V., 2004). The net result of these modifications in P. fluorescens is the formation of a α-keto acid pool that has a crucial role in combating the oxidative condition resulting from the exposure to the trivalent metal. There are several strategies to increase the amount of a particular metabolite in biological systems. The activity of the anabolic enzyme may be increased i.e. ICDH (NAD+/NADP+ dependent) or the activity of the catabolic enzyme may be decreased i.e. α-KGDH (Figure 73)
Figure 73: Metabolic reconfiguration to combat the oxidative stress generated by gallium in P. fluorescens
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The metabolism of α-ketoglutarate in biological systems does not solely depend on the activity of α-KGDH. The enzyme aspartate transaminase is able to transaminate α-ketoglutarate to glutamate. This enzyme was found not to be significantly increased in Ga-stressed cells. Therefore, α-ketoglutarate is not preferentially transaminated to glutamate in Ga-stressed cells. Similarly, the enzyme glutamate dehydrogenase is only slightly more active in Ga-stressed cells, thereby suggesting that the α-ketoacid is not preferentially channeled towards glutamate. Another α-ketoglutarate consuming enzyme, α-ketoglutarate reductase was not found in our system and therefore, α-ketoglutarate is not channeled to 2-hydroxyglutaric acid. In some biological systems, the conversion of α-ketoglutarate to succinate is attained in a CoA and NAD+ independent fashion. An alternative pathway for the catabolism of α-ketoglutarate in a mutant of Bradyrhizobium japonicum lacking α-KGDH has been recently demonstrated (Green, S., 2000). These bacteria were able to suffice their ATP demand via the expression of a α-ketoglutarate decarboxylase. Thus, the organism was able to bypass the α-KGDH reaction and metabolize α-ketoglutarate to succinate semialdehyde in a CoA independent fashion. And as this enzyme was absent in Ga-stressed P. fluorescens, an alternative mechanism involved in the metabolism of α-ketoglutarate was not operative in these cells. Hence, the pool of α-ketoglutarate may be intended to act as an ROS scavenger.
Figure 74: Possible fate of α-ketoglutarate metabolism
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INHIBITION OF α-KGDH
How are these enzymes being modulated in P. fluorescens? It as been reported that gallium competes with magnesium in biological systems and the latter is also known to bind α-KGDH. Therefore, gallium may bind α-KGDH in P. fluorescens thus inhibiting the enzyme. Our result demonstrates that when gallium is allowed to interact with the membrane fraction of the control cells for 15 min, α-KGDH is inhibited by two-fold. However, it is not clear if the inhibition is direct or caused indirectly by the metal. Arsenite, As(III), a toxic metal was shown to inhibit the purified α-KGDH. Arsenite was found to bind to the lipoic acid residue on the E2 subunit thereby inhibiting the enzyme in an irreversible manner. One of the products of lipid peroxidation 4-hydroxy-2-nonenal (HNE) inhibits α-KGDH in vitro at much lower concentrations then those found in oxidative environments (Humphries, K. 1998). The lipoic acid residue is a target of HNE. Hydrophobic interactions occur between the hydrocarbon chain of lipoic acid and HNE. Since gallium causes the peroxidation of lipids it is possible that HNE does in fact inhibit α-KGDH. The sulfhydryl groups present in α-KGDH may also act as antioxidants and neutralize the peroxide produced in the bacteria subjected to gallium. The result would be the inhibition of the enzyme. This may be the trigger that leads to the decrease of α-KGDH expression as the level of the E2 subunit is lower in the Ga-stressed cells.
Figure 75: Inhibition of α-KGDH in Ga-stressed P. fluorescens
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The inhibition of α-KGDH may serve more then one purpose. Not only does it create a pool of α-ketoglutarate that can act as a scavenger of peroxide but it also reduces the generation of NADH. The cofactor NADH is a powerful reducing agent and is capable of regenerating Fe+2, the unsafe form of iron from Fe+3. This may also explain the inhibition of α-KGDH which ultimately results in a reduction of total cellular NADH. The increase in NADPH/NADH ratio as a strategy to limit the generation of ROS has been demonstrated in E.coli (Brumaghim, J.L., 2003). NADPH is a much slower reductant of Fe+3 than is NADH. The latter can also participate in the generation of ROS since it is a substrate of complex I of the ETC where the majority of superoxide is produced.
Recently, an in vivo model of heart ischemia-reperfusion in rats, demonstrated that α-KGDH as well as ACN were inactivated whereas other Krebs cycle enzymes were unaffected. The inhibition of these two enzymes would explain the reduced mitochondrial respiration observed. The inactivation of α-KGDH was accompanied with the loss of the lipoic residue (Sadek, H., 2002). Free radical events triggered by H2O2 inhibited α-KGDH in vitro as observed in P. fluorescens. The enzyme α-KGDH therefore appears to act as a redox sensitive enzyme as is ACN. More importantly, α-ketoglutarate is an important metabolite involved in DNA repair (Begley, T.J., 2003) and is known to regulate genes involved in the assimilation of ammonia (Galvez, S., 1999) (Figure 76). This may explain the lower levels of α-KGDH found in Ga-stressed P. fluorescens by Coomassie staining and western blot analysis. Indeed, the increased levels of α–ketoglutarate due to the inhibition of α-KGDH by gallium may destabilize the α-KGDH mRNA and decrease the expression of the corresponding enzyme. Also, the increased levels of α–ketoglutarate may induce the expression of the ICDH-NADP+ isoenzyme.
Figure 76: α-ketoglutarate, a metabolite with diverse biochemical functions
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Classical ROS detoxifying pathways may not be an important contributor in combating gallium stress when iron is a limiting factor. Catalase is not necessarily the main protector against ROS mediated cellular damage in vivo in humans. Subjects lacking catalase are not very sensitive to oxidative stress compared to humans lacking G6PDH (Brumaghim, J., 2003). However, increasing the level of ICDH-NADP+ does augment the resistance to oxidative stress in mice.
The role of pyruvate, a ketoacid in the Giardia intestinalis, a parasite that lacks catalase or other peroxidases has been shown. Pyruvate was effective in neutralizing the H2O2 induced generation of ROS (Giancarlo, A., 2001). For a hundred years now, the effectiveness of α-ketoacids as antioxidants is known. They are part of the nonenzymatic antioxidant system. In the presence of H2O2, α-ketoglutarate and pyruvate undergo a decarboxylation producing carbon dioxide and water (Velvizhi, S., 2002):
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This phenomenon may be occurring in Ga-stressed P. fluorescens and appears to be an important strategy the cell utilizes to combat ROS stress. When cell free extract from control cells are incubated with labeled Ga-citrate and NAD+, the α-ketoglutarate produced is nonenzymatically converted to succinate as detected by the 13C NMR peaks at 33 ppm and 181 ppm. The only possible explanation for this observation is that α-ketoglutarate is utilized to detoxify the peroxide generated by gallium. On the other hand, when the labeled substrate citrate and NAD+ are added to the cell free extract from control cells, the metabolism is halted at the level of α-ketoglutarate indicated by a peak at 29 ppm, attributable to the labeled CH2 group of this metabolite. The appearance of the succinate peaks are not an artifact and are not a consequence of the enzyme isocitrate lyase (ICL) that is also able to metabolize isocitrate to glyoxylate and succinate. In this case the succinate produced would be labeled only at the C2 position and arise at 33 ppm.
[pic]
[pic]= 13C-labelled
α-Ketoacids have been shown to successfully protect human breast cancer (MCF7) cells against menadione induced DNA damage and cytotoxicity. Menadione is a quinone that leads to dramatic increased levels of intracellular peroxide through intensive redox cycling in biological systems. Pyruvate protects cells against menadione-generated H2O2 and reduces H2O2 induced DNA damage thereby improving cellular growth in vivo (Nath, K. 1995). α-Ketoacids act as protector of oxidation-sensitive enzyme involved in metabolism. More significantly, inside cells, the levels of α-ketoacids are in the millimolar range whereas concentrations of H2O2 are in the nanomolar range. Therefore the role of α-ketoacids in H2O2 scavenging is as important in the antioxidative systems as catalase and glutathione peroxidase. Hence, the pathways leading to the biosynthesis of these α-ketoacids would play a critical role in situations of oxidative stress. α-Ketoglultarate is more powerful then vitamin C as an antioxidant. It is used as a cyanide poisoning antidote, as a ROS scavenger for ischemia/reperfussion during heart surgery and for hyperammonemia therapy (Velvizhi, S. 2002). α–Ketoglutarate may serve as an important mitochondrial H2O2 scavenger, preventing a vast array of metabolic diseases occurring when the mitochondrial DNA is altered. Recently, α-ketoacids have been suggested to inhibit the increased levels of ROS induced by H2O2 in erythrocytes and in cultured striatal neurons (Yamamoto, H., 2003). Therefore, the inhibition of α-KGDH observed in many neurodegenerative diseases such as Alzheimer’s disease may be the consequence of a cellular strategy to reduce the levels of oxidative stress and the propagation of the disease. When maleate, a thiol consuming agent that is known to increase the levels of α-ketoacids was administered to rats, the level of peroxide in the kidneys was significantly lower and the α-ketoacids were deemed excellent peroxide scavengers (Nath, K., 1995). Furthermore, this experiment demonstrates that glutathione and its peroxidase are not necessarily the first line of defense against peroxide induced oxidative stress since maleate also depletes the levels of GSH. In our study, when gallium cells were grown to stationary phase, α-KGDH had low activity. When these cells were transferred to a media devoid of gallium but supplemented with 1mM menadione, the activity of α-KGDH remained low compared to control bacteria. This metabolic strategy is most likely evoked by the microorganism to allow the generation of α-ketoglutarate with the aim of neutralising the menadione-generated H2O2. Thus, this suggests that the inhibition of α-KGDH observed in Ga-stressed cells may serve as a strategy to increase the α-ketoglutarate pool in an effort to fend an oxidative environment created by gallium.
Although DNA base oxidation, protein carbonyls level, oxidized amino acids and oxidized lipids are biomarkers of oxidative stress, the amount of decarboxylated α-ketoglutarate (succinate) may be an excellent gauge for an oxidative environment. Because they are non toxic and can be easily transported across membranes, α-ketoacids are excellent therapeutic agents in cases of elevated oxidative damage such as carcinogenesis, radiation injury or drug toxicity damage is involved. Also, α-ketoacids could be involved in the treatment of cataracts. Other enzymes such as G6PDH and ICDH-NADP+ can also play a pivotal role in ROS homeostasis. Indeed, these NADPH-generating enzymes are increased in Ga-stressed cells.
The TCA cycle depends on the close proximity of many enzymes that may form a metabolome and allow for funneling of metabolites to specific enzymes/sites. BN-PAGE analysis did reveal a close proximity between ACN and ICDH-NADP+. One advantage of this spatial arrangement would enable the α-ketoglutarate generated from the ICDH-NADP+ to protect ACN against ROS damage and allow ACN to function even under iron-deprived situations.
When P. fluorescens is grown in gallium, the growth of the bacteria is greatly retarded and an adaptive phase is evident. Gallium has been shown to reduce cellular proliferation due to its ability to block iron uptake and iron mediated processes. This may explain why Ga-stressed P. fluorescens has lower iron-content compared to control cells. Gallium has been shown to replace iron in various iron containing proteins (Figure 77). Due to this property, gallium bound to transferrin is toxic to tumors. It has been suggested that the internalization of gallium mimics the depletion of iron in leukemia cells in vitro. Gallium can also create a state of iron starvation in red blood cells (Seligman, P. 1992). Addition of iron may reverse the toxicity mediated by gallium. Hence, there exists an intimate relationship between iron and gallium in biological systems. Owing to its ability to mimic iron, it was not surprising to find an increase in labile iron pool and ROS in Ga- stressed cells. When peroxide reacts with iron sulfur cluster (4Fe-4S) of proteins such as ACN and FUM, the liberated iron ion joins what is called the iron labile pool. It is this iron that has proooxidative properties and that is so destructive. The inhibition of ACN does also indicate that gallium alters the homeostasis of iron. Gallium may substitute directly for iron in this enzyme. Alternatively, the peroxide produced when cells are exposed to gallium may also react with the iron present in ACN and inhibit the enzyme. In these situations, the overexpression of ACN may not be a good metabolic strategy since the more protein is expressed, the more iron is necessary to accommodate the enzyme (Kruszewski, M., 2003). A better strategy is to express a novel ICDH-NADP+ that may compensate for the lower activity of ACN in Ga-stressed bacteria and protect ACN via α-ketoglutarate. This isoenzyme would also increase the flux of citrate to the TCA cycle. Furthermore, iron-independent ACN is not known. On the other hand, fumarases do exist as iron-containing and iron-free enzymes (Benov, L., 2002). In this instance, the iron-free species (FUM C) is expressed to fend the iron-deprived conditions created by gallium. Furthermore, this enzyme would be recalcitrant to ROS attack. In fact, this enzyme is expressed under conditions of iron starvation and is a favorable metabolic strategy that ensures the continuity of the TCA cycle under less than favorable conditions.
Figure 77: Adaptation to iron deprivation evoked by gallium stress in P. fluorescens
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Gallium sequestration
Many bacteria including cyanobacteria and heterotrophic bacteria utilize siderophores to bind iron in the environment as an uptake mechanism and to survive iron-limited environment. Aquachelins are siderophores produced by heterotrophic bacteria to acquire Fe+3. Many iron siderophores contain the characteristic β-hydroxyaspartate residue. These β-hydroxyaspartate containing siderophores are often expressed in situations of iron stress (Barbeau, K., 2001).
Gallium is known to bind to bacterial iron siderophores (Olakanmi, O. 2000). It is possible that gallium binds to siderophores produces by P. fluorescens. In this instance, the β-hydroxyaspartate has also been shown in this moiety (Santani, 1992). Once saturated with gallium, these siderophores are excreted in the exocellular environment. The synthesis of a β-hydroxyaspartate residue would depend on the hydroxylation of the amino acid aspartate. In biological systems, this is assumed by aspartyl β-hydroxylase, a α-ketoglutarate-dependent dioxygenase. Thus enzymes like AST, GDH and MDH that will all promote the biosynthesis of aspartate appear to function effectively in Ga-stressed cells. Indeed most of the gallium is sequestered exocellularly in this hydroxyaspartate containing moiety.
CONCLUSION
It appears that the global cellular metabolism has been reconfigured with aim of circumventing the toxic influence of gallium. This metabolic reconfiguration allows the cell to deal with oxidative stress, iron deprivation and gallium immobilization. α-Ketoglutarate is key in this strategy and provides the ideal tool to combat ROS stress under iron-deprived situation. The decarboxylation of this moiety also limits the production of NADH, a generator of ROS. This shows the plasticity of the TCA cycle and extends its significance beyond its classical role as a provider of precursors for anabolic reactions and for oxidative phosphorylation. The shift towards an iron-limited metabolism is evident and this organism appears to have invoked this strategy. Decreased ACN activity supplemented by increased ICDH-NADP+ activity and expression of FUM C, an iron independent enzyme are pivotal for the survival of Ga-stressed P. fluorescens. Scheme 1 depicts the metabolic flux triggered by gallium in P. fluorescens. This study illustrates the significance of metabolism in cellular functions and demonstrates how metabolomic and proteomic approaches can help unravel the molecular workings of biological systems.
Scheme 1: Ga-evoked metabolic shift operative in P. fluorescens
[pic]
[pic]
Legend: Bold lines indicate enhanced activity whereas X indicates decreased activity. The critical enzymes governing the adaptive response are 1. ACN, 2. ICDH-NADP+, 3. α-KGDH, 4. FUM C, 5. Catalase, 6. Non-enzymatic H2O2 detoxification
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-----------------------
glutamine
nucleic
acids
ROS
Inflammation
Hypercholesterolemia
Hypertension
Diabetes
Ischemia-Reperfusion Injury
Atherosclerosis
Congestive Heart Failure
Ammonia toxicity
Aging
Carcinogenesis
Alzheimer’s disease
Phagocytosis
UV and ionizing radiations
Anticancer drugs
Neurodegeneration
Tobacco smoke
O2-· + Cu+2 O2 + Cu+
GSH
Ascorbic acid
Cu+ + H2O2 Cu+2 + OH- + OH·
Culture
Pellet
(cells)
Centrifuge
10,000 x g 15 min
Spent fluid
(discard)
Pellet
(cell)
Supernatant
(discard)
Pellet
(cells)
Supernatant
(discard)
Resuspend
In 0.85% NaCl
Centrifuge
10,000 x g 15 min
Resuspend
In 0.85% NaCl
Centrifuge
10,000 x g 15 min
TBARS
532 nm
Isocitrate
Glyoxylate + Succinate
Glyoxylate + 2,4-DNPH
2,4-dinitrophenylhydrazone
Isocitrate + NAD+
(-ketoglutarate + NADH + H+ + CO2
(-ketoglutarate + NAD+
Succinate + NADH + H+ + CO2
(-KGDH
Oxaloacetate + Acetyl-CoA Citrate + HSCoA
HSCoA + DTNB TNB-SCoA +TNB-
CS
Glyoxylate + Acetyl-CoA Malate + HSCoA
HSCoA + DTNB TNB-SCoA +TNB-
MS
Succinate + FAD
Fumarate + FADH2
SDH
succinate
Aconitate
[?]00.0/090>0@0A0B0D0E0G0H0J0K0M0N0T0U0V0X0Y0_0`0b0c0α-ketoglutarate
succinate
α-ketoglutarate
B
A
B
A
A
B
C
B
A
................
................
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