INTRODUCTION - Laurentian University

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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 25.2 Metabolic Turnover and Cellular
ATP Production
Figure 1: An Overview of metabolism
Copyright © 2004 Pearson Education, Inc., publishing as Benjamin Cummings
Figure 25.2
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
1
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)
glutamine
nucleic
acids
2
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
3
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
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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
Genotype
Genomics
Expressional/Functional Genomics
Transcriptomics
Metabolic Network
Metabolic Nodes
Proteomics
Functional Proteomics
Regulatory
Metabolic
Metabolomics/Metobolome
“Phenotype”
Metabolites
Does the living/what you see
“Foundation” of Life
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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.
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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).
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Figure 4: Physiological conditions that trigger the generation of ROS
UV and ionizing radiations
Inflammation
Neurodegeneration
Ischemia-Reperfusion Injury
Tobacco smoke
Hypercholesterolemia
Phagocytosis
Hypertension
ROS
Atherosclerosis
Diabetes
Ammonia toxicity
Aging
Anticancer drugs
Congestive Heart Failure
Carcinogenesis
Alzheimer’s disease
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
8
(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
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
9
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
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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
Five reactions of the
form, dihydrolipoamide, is modified following H2O2 or O2-· exposure in both yeast and
pyruvate dehydrogenase multienzyme complex
E-Coli (Figure 6).
6: Different subunits involved in α-keto acid dehydrogenases (Taken from
- EFigure
1 (pyruvate dehydrogenase) contains TPP and catalyzes reactions 1 and 2.
- EVoet,
transacetylase) contains lipoamide; it catalyzes reaction 3.
1990)
2 (dihydrolipoyl
-E3 (dihydrolipoyl dehydrogenase) contains FAD and a redox-active disulfide
and catalyzes reactions 4 and 5.
10
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
11
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
12
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-
13
phosphate dehydrogenase (G6PDH), malic enzyme (ME) and isocitrate dehydrogenase
(ICDH).
Figure 7: Oxidative and antioxidative systems (Taken from Carri, M., 2003)
Pentose phosphate cycle
Glucose-6-Phosphate
OH- + ClPeroxynitrous ONOOH
acid
Fe+3
NADP+
Fe+2
NADPH
Glutathione
reductase
HOCL Hypochlorous acid
Peroxynitrite ONOO-
H2O +1/2 O2
NO Cl-
iNOS
GSSG
Se
GSH Glutathione
Catalase
Haem oxygenase
HO-1
peroxidase
SOD
O2-· + O2-·
H2O
H2O2
Fe+2
HO·
Polyunsaturated
Fatty acids
β-carotene
Damage to protein,
Carbohydrate and DNA
β-caratenoid
radical
RO·
RO2·
Lipid
hydroperoxides
α-tocopherol
OH
α-tocopherol
O·
Dehydroascorbic
acid
Ascorbate
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
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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 BenYoseph, O., 1996)
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
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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
Step 3: Generation of CO2 by an NAD+-linked dehydrogenase
involved
in the production
of NADPH utilized for both lipid biosynthesis and the
--isocitrate
dehydrogenase
--oxidative
decarboxylation
neutralization
of hydroxyperoxides,
its role in ROS defense is now being fully
understood.
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 upregulated 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.
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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-4phenyl-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
17
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 αKGDH 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).
18
Figure 9: Reaction between HNE and the lipoic residue of α-KGDH (Taken from
Humphries, K. 1998)
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
19
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)
2. Aconitase
31
20
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).
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Figure 12: Pathways of iron homeostasis in hepatocytes (Taken from Goswami, T.,
2002)
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.
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 βpeptide, 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
23
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)
It has been postulated that a decrease in iron stores may retard the onset of agerelated 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.
24
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)
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
25
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:
O2-·
+ Cu+2
GSH
Ascorbic acid
O2 + Cu+
Cu+ + H2O2
Cu+2 + OH- + OH·
Figure 15: Cytotoxic mechanisms of copper Taken from Pourahmad, J., 2001)
26
Protein and Cu
endocytosis
Autophagic
Vacuole
Cathepsins
Cystine
Lysosome
H2O2
Cu+1
O2
Autophagocytosed
mitochondria
O2-·
Cysteine
Cu+2
OH- + HO·
Membrane
disruption
Lipid peroxidation
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)
27
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
28
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).
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 βamyloid 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
29
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).
30
Figure 18: Biochemical targets of zinc (Taken from Dineley, K., 2003)
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
31
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
Aluminum
(Environment/Diet
Amyloid deposition
Accumulation of
Aluminum in the brain
Oxidative stress
Lipid peroxidation
ROS formation
Depletion of
Anitoxidant defense
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.
32
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
33
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 (AlAoukaty, 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+3transferrin 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)
34
Activity
Specificity
Redox
Catalytic
Dinuclear iron center
Tyrosine free radical
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.
35
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.
36
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
37
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.
38
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)
39
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)
40
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)
41
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)
42
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
43
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
44
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
45
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
Culture
Pellet
(cells)
Pellet
(cell)
Pellet
(cells)
Centrifuge
10,000 x g 15
min
Resuspend
In 0.85% NaCl
Centrifuge
10,000 x g 15
Resuspend
In 0.85% NaCl
Centrifuge
10,000 x g 15
Supernatant
(discard)
Spent
fluid
(discard)
Supernatant
(discard)
46
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
47
Isolated cells
Sonicate
4 × 15 seconds
Cell Free Extract
Centrifuge
180,000 × g 60 min
Supernatant
(crude soluble
fraction)
Pellet
(membrane)
Centrifuge
180,000 × g 120 min
Pellet
(discard)
Supernatant
(Soluble fraction)
Isolation of spheroplasts of P. fluorescens
The inner membrane of P. fluorescens was isolated by a modified version of the
method described by Mizuno (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
48
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
Bacteria
Wash with 20%
sucrose
Resuspend cells in
isotonic buffer and
lysozyme and
Incubate at 37°C for
60 minutes
Centrifuge
10,000 xg 15
minutes
Supernatant
(outer membrane and
periplasm)
Spheroplast
Resuspend in
Lysis buffer
Pellet
(spheroplast)
Centrifuge
100,000 xg 30
minutes
Supernant
(cytoplasm)
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%
49
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
50
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.
TBARS
532 nm
Protein carbonyl measurement
51
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
Measurement of total carbonyls
ratio three times.
The final precipitate was dissolved in 1.0ml
of 6M guanidine
and the
(Spectrophotometric
DNPH
assay)
which the pelleted proteins were washed with a solution of ethylacetate:ethanol in a 1:1
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.
DNP
O
protein
H2O2 Fe
activated
neutrophil
oxidized
protein
DNPH
DNPprotein
Absorbance
at 370 nm
e.g. arg --->-glutamylsemialdehyde
Dinitrophenylhydrazone-protein
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
52
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 mM1
cm-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 Gastressed 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
53
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
(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
54
Levels of aldehydes and ketoacids were measured using 2,4dinitrophenylhydrazine (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
55
Isocitrate
Glyoxylate + Succinate
Glyoxylate + 2,4-DNPH
2,4-dinitrophenylhydrazone
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 αketoglutarate. The amount of ketoacid produced was determined spectrophotometrically
using 2, 4-DNPH (Romanov, V., 1999) and α- ketoglutarate served as the standard.
ICDH
Isocitrate + NAD+
-ketoglutarate + NADH + H+ + CO2

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.
56
-ketoglutarate + NAD+
-KGDH
Succinate + NADH + H+ + CO2
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 αketoglutarate 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
57
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).
Oxaloacetate + Acetyl-CoA
HSCoA + DTNB
CS
Citrate + HSCoA
TNB-SCoA +TNB-
58
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).
MS
Glyoxylate + Acetyl-CoA
HSCoA + DTNB
Malate + HSCoA
TNB-SCoA +TNB-
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 ml1
) 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.
Pyruvate + CoA + NAD+
PDH
Acetyl-CoA + CO2 + NADH + H+
59
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.
Succinate + FAD
SDH
Fumarate + FADH2
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.
60
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).
G6P + NADP+
G6PDH
phosphoglucono-lactone + NADPH + H+
6-Phosphogluconate dehydrogenase (6PGDH) activity
61
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
62
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.
Malate + NAD+
MDH
Oxaloacetate + NADH+ H+
Fumarase (FUM) activity
The conversion of fumarate to malate is catalyzed by FUM (EC 4.2.1.2).
2,4-DNPH at 450 nm (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 (Jordan et al., 1999). Aconitate served
63
as the standard. Blanks were prepared in a similar fashion however, omitting the substrate
from the mixture.
Citrate
ACN
cis-aconitate
cis-aconitase
Isocitrate
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 αketoglutarate. 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
64
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
65
1 mM EDTA
1 mM PMSF
1 mM DTT
15 mM MgCl2
pH 7.3 at Room Temperature
1H
NMR Buffer
10 mM sodium phosphate
5 mM MgCl2
pH 7.3 at Room Temperature
Activity Buffer
25 mM Tris HCl
5 mM MgCl2
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.
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)
66
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 (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
67
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.
Acryl-Bis mix (49.5
%T, 1.5% C)
3x buffer
water
75% glycerol
10%APS
TEMED
4%
16%
Upper Gel (sample
234
937
loading)
273
967
1699
--9.7
1.0
967
223
773
7.6
0.8
1136
2000
-30
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.
68
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)
8.96g Tricine (50 mM)
3.138g BisTris (15 mM)
0.2g Coomasssie blue G 250
pH 7.0 at 4°C
Colourless Cathode Buffer
8.96g Tricine (50 mM)
3.138g BisTris (15 mM)
pH 7.0 at 4°C
3X Gel Buffer (50ml)
Anode Buffer (1L)
9.84g aminocaproic acid (1.5 M)
1.567g BisTris (150 mM)
pH 7.0 at 4°C
10.45g BisTris (50 mM)
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
69
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:
isocitrate
NADPH
INT
PMS
α-ketoglutarate
NADP+
Formazan
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
70
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 Panisidine 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
71
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
72
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
73
(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
74
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
(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
(30% T, 0.8% C) (100ml)
29.2g Acrylamide
4X Tris/SDS pH 8.8
1.5 M Tris Base
0.4% SDS
75
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
0.4% SDS
pH adjusted to 68 with 11 N HCl
15.1g Tris Base
72.0g Glycine
5.0g SDS
Coomassie Blue
Staining Solution
Destaining Solution
50% Methanol
10% Acetic acid
0.2% Brilliant Blue R 250
50% Methanol
10% Acetic acid
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
76
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)
3.03 g Tris-base
14.4 g glycine
200 ml Methanol
store at 2-8°C
Tris Buffered Saline (TBS) (1L)
2.42 g Tris-base
8 g NaCl
adjust pH to 7.6 with 2N HCl
store at 2-8°C
Tween Tris Buffered
Saline (TTBS)
Dilute required volume of
Tween™ 20 in TBS to give
A 0.1% (v/v) solution
store at 2-8°C
5% Blotto
5% (w/v) dried skim milk
in TTBS
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
77
Bands were quantified using Scion Image V. 4.0.2 (Scion Corporation, USA)
RESULTS
78
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.
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)
79
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.
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)
80
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.
C NMR data and chemical analysis revealed fingerprint characteristic of β-
13
hydroxyaspartate (Figure 26).
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)
81
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 Gastressed cultures (Figure 27). However, more oxidized lipids were evident in the cells
subjected to gallium.
82
600
*
0.5
500
*
0.4
400
0.3
300
0.2
200
0.1
100
0
µg of protein/ml of culture
TBARS (nmol/mg of protein)
0.6
0
0
10
12
15
18
20
23
25
30
36
50
52
55
60
65
67
70
75
80
Time of growth (hrs)
TBARS control membrane
growth profile of control culture
TBARS Ga-stressed membrane
growth profile of Ga-stressed culture
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.
83
Oxidative stress marker: oxidized proteins
The oxidized proteins were visualized with the aid of DNPH and in 1 mM Ga-
0.05
600
*
0.045
500
0.04
0.035
400
0.03
0.025
300
0.02
200
0.015
0.01
100
µg of protein/ml of culture
pmol of carbonyl/mg of protein
stressed cells an increase in oxidized proteins was recorded (Figure 28).
0.005
0
0
0
10
12
15
18
20
23
25
30
36
50
52
55
60
65
67
70
72
75
80
Time of growth (hrs)
protein carbonyl in control cytoplasm
growth profile of control culture
protein carbonyl in gallium cytoplasm
growth profile of Ga-stressed culture
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.
84
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).
35
Amount of peroxide (umol/mg of protein)
H2O2of(µmol/mg
of protein)
Amount
of protein)
H2O2 (umol/mg
40
30
25
20
15
10
5
0
Citrate
Ga-citrate
Substrate (5mM)
Substrate 5mM
Figure 29: Peroxide production by the membrane fractions from P. fluorescens
(n=3)
85
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
14
12
Amount of superoxide generated (nmol/mg of proteins)
Amount of superoxide generated (nmol/mg of protein)
Superoxide generated (nmol/mg of protein)
powerful oxidant in P. fluorescens (Figure 30).
10
8
6
4
2
0
Citrate
Ga-citrate
Substrate (5mM)
Substrate (5mM)
Figure 30: Superoxide production by the membrane fractions from P. fluorescens
(n=3)
86
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-
Antioxidative system operative in
Ga-stressed cells: Catalase activity
stressed cultures. At logarithmic phase of growth, a 4-fold decrease was observed in the
120
600
100
500
80
400
*
60
300
40
200
*
*
20
100
0
µg of protein/ml of culture
Activity
catalase
(%)
Activity ofof
catalase
(%)
soluble fraction isolated from the Ga-stressed bacteria (Figure 31).
0
0
10
12
15
18
20
23
25
30
36
50
52
55
60
65
67
70
72
75
80
Time of growth (hrs)
activity of catalase in control cytoplasm
growth profile of control culture
activity of catalase in gallium cytoplasm
growth profile of gallium culture
Activity of catalase (H2O2 decomposition) in control and Ga-stressed P. fluorescens (n=3). Values
Figure 31: Activity of catalase (H2O2 decomposition) in control and 1 mM GaAre means ± S.D.. Catalase values differ significantly from controls (p≤0.05).
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
87
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
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
88
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).
600
*
140
500
120
400
100
80
300
*
60
200
40
*
20
100
*
0
µg of protein/ml of culture
Activity of SOD (100%)
160
0
0
10
12
15
18
20
23
25
30
36
50
52
55
60
65
67
70
72
75
80
Time of growth (HRS)
activity of SOD in control membrane
growth profile of control culture
activity of SOD in gallium membrane
growth profile of gallium culture
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
89
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 Gastressed cells thus indicating enhanced activity. (Figure 34).
1
2
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
90
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.
91
13C NMR analyses of citrate metabolism in CFE
A
43
α-ketoglutarate
Aconitate
32
29
36
54
130
ppm
succinate
B
succinate
54 44
181
α-ketoglutarate
33 32
ppm
92
C
43
α-ketoglutarate
Aconitate
32
54
130
D
36
29
ppm
succinate
succinate
54
181
33
32
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.
93
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.
94
Table 1: Activities of metabolic enzymes in control and Ga-stressed cells
Enzymes
Glucose-6-Phosphate phosphatase
Pyruvate carboxylase
Specific activity in CFE
Specific activity in CFE
(nmol/mg of protein/min) of (nmol/mg of protein/min) of
control cells (n=3)
1 mM Ga-stressed cells (n=3)
0.016±0.002
0.011±0.003
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
3.3±0.2
2.0±0.2
Pyruvate dehydrogenase
95
Table 2: A comparative evaluation of various enzymatic activities in control and Gastressed 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
3X↓
Malate synthase
3X↓
Citrate Synthase
↑76%
Pyruvate dehydrogenase
↓40%
*** control is taken as 100% or 1
96
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).
97
2
1.8
1.6
1.4
Absorbance
1.2
1
0.8
0.6
0.4
0.2
0
0
5
10
15
20
25
30
35
40
45
50
55
60
65
70
75
80
85
90
95
100 105
Time (sec)
kinetic of ICDH in control soluble fraction
kinetic of ICDH in Ga-stressed soluble fraction
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.
98
α-Ketoglutarate synthesis: ICDHNADP+ activity
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
1600
600
*
1400
*
1200
500
*
400
1000
800
300
600
200
400
100
200
0
µg of protein/ml of culture
Specific activity (nmol/mg of protein/min)
enzyme was higher in Ga-stressed bacteria compared to control bacteria (Figure 37).
0
0
10 12 15 18 20 23 25 30 36 50 52 55 60 65 67 70 72 75 80
Time of growth (hrs)
specific activity of ICDH in control soluble
specific activity of ICDH in gallium soluble
growth profile of control culture
growth profile of gallium culture
Figure 37: Specific activity of ICDH-NADP+ enzyme in control and in 1 mM Gastressed cells at various growth intervals (n=3). Values are means ± S.D.. ICDH-NADP+
differ significantly from controls (*) (p≤0.05).
99
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
A
B
Band A
100
Band B
Figure 38: BN PAGE analysis of ICDH-NADP+ activity in control and 1 mM Gastressed 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.
101
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
Figure 39: 2D BN PAGE analysis of ICDH-NADP+ activity in control and 1 mM Gastressed cells.
Lane 1: cells obtained in citrate medium (control). Lane 2: cells obtained in Ga-citrate
medium. 60 µg were loaded in each lane.
102
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).
105
*
*
100
Activity of ICDH (NADP+) (%)
Activity of ICDH-NADP+ (%)
110
*
95
90
85
80
75
A
B
C
D
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
103
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
A
B
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.
104
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 Gastressed 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
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.
105
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
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.
106
ICDH-NAD+
In biological systems, the synthesis of α-ketoglutarate is also mediated by ICDHNAD+. 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 αketoglutarate 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
*
2.5
600
500
2
400
1.5
300
1
200
0.5
100
0
µg of protein/ml of culture
Specific activity (nmol/mg of protein/min)
present in the media.
0
0 10 12 15 18 20 23 25 30 36 50 52 55 60 65 67 70 72 75 80 //
0
6
Growth (HRS)
Specific activity of ICDH in control membrane
Specific activity of ICDH in gallium membrane
Specific activity of ICDH im membrane of gallium cells introduced in a control media
grow th profile of control cells
grow th profile of gallium cells
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).
107
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).
Specific activity umol*min-1*mg protein -1
35
30
25
20
15
10
5
0
Control
Gallium
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.
108
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 αketoglutarate.
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
109
Confirmation of the decreased activity of α-KGDH in the membrane fraction of
CFE of control and Ga-stressed P fluorescens.
1
H 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 αketoglutarate 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).
110
A
SUCCINATE
ppm
B
α-Ketoglutarate
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)
111
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 Gacitrate 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
1
2
2
Figure 47: BN PAGE detection of α-KGDH activity in control and 1 mM Gastressed 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
112
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
B
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
113
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
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.
114
α-KGDH activity in P. fluorescens at various stages of growth in citrate and Gacitrate 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).
600
30
500
25
400
*
20
*
15
300
*
*
200
10
100
5
0
µg of protein/ml of culture
Specific activity (nmol/mg of protein/min)
35
0
0
10
12
15
18
20
23
25
30
36
50
52
55
60
65
67
70
72
75
80
//
0
6
Time of growth (hrs)
Specific activity of KGDH in control membrane
Specific activity of KGDH in gallium membrane
Specific activity of KGDH in membrane of gallium cells introduced in a control media
growth profile of control cells
growth profile of gallium cells
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 Gastressed 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)
115
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 αKGDH 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
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.
116
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.
Specific activity (nmol*min-1*mg protein-1
Metal
(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
117
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
35
30
Specific activity (µmol/mg of protein/min)
Specific activity (nmol/mg of protein/min)
directly or indirectly in the decrease of α-KGDH activity.
25
20
*
15
10
5
0
0
100
200
500
Amount of Ga(NO3)3 µM
Amount of Ga(NO3)3 (µM)
Figure 52: Modulation of α-KGDH specific activities (decomposition of αketoglutarate) 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).
118
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
α-ketoglutarate dehydrogenaseb
α-ketoglutarate reductasea
α-ketoglutarate reductaseb
CoA independent α-ketoglutarate
dehydrogenaseb
CoA independent α-ketoglutarate
dehydrogenasea
Glutamate dehydrogenasea
Glutamate dehydrogenaseb
Aspartate transaminaseb
Specific activity (nmol*min-1*mg protein-1)
Control cultures
Ga-stressed cultures
Nil
Nil
Nil
Nil
Nil
Nil
Nil
Nil
Nil
14.6±0.8
nil
108±1.5
Nil
17.5±0.5
Nil
122±8.1
a = membrane fraction of cell free extract
b = soluble fraction of cell free extract
119
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).
120
25
Specific activity of a-KGDH (umol/mg of protein/min)
Specific activity (nmol/mg of protein/min)
30
20
*
*
15
10
5
0
A
B
C
D
Media
Figure 53: Regulation of α-KGDH specific activities (decomposition of αketoglutarate) 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).
121
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
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.
122
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 αKGDH 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).
250
Peroxide decomposed (umol)
200
150
100
50
0
1
2
3
Alpha-ketoglutarate concentration (mM)
Figure 55: The decomposition of H2O2 by α-ketoglutarate. Values are means ± S.D..
123
13C
NMR standards
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).
A
PPM
-ketoglutarate
B
PPM
succinate
124
H2O2 + -ketoglutarate =
C
PPM
Succinate
Figure 56: Panel A: 13C NMR spectra corresponding to α-ketoglutarate. Panel B: 13C
NMR spectra corresponding to succinate. Panel C:
13
C NMR spectra depicting the
reaction between α-ketoglutarate and H2O2.
125
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 Gastressed 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)
126
A
B
Figure 57: Detection of iron sulfur (Fe-S) clusters in; A: control soluble fraction and B:
1 mM Ga-stressed soluble fraction
127
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.
128
600
120
500
100
400
*
80
300
60
200
40
100
20
0
µg of protein/ml of culture
Specific activity (nmol/mg of protein/min)
140
0
0
10
12
15
18
20
23
25
30
36
50
52
55
60
65
67
70
72
75
80
//
0
6
Growth (hrs)
Specific activity of ACN in control soluble
Specific activity of ACN in gallium soluble
Specific activity of ACN in regulation control soluble
Growth profile of control culture
Growth profile of gallium culture
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)
129
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).
1
2
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.
130
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.
131
100
Specific activity (nmol/mg of protein/min)
Specific activity (nmol/mg of protein/min)
120
80
*
*
*
60
40
20
0
A
C
B
D
Media
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)
132
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
50
600
*
45
500
40
35
400
30
25
300
20
200
15
10
100
5
0
0
0 10 12 15 18 20 23 25 30 36 50 52 55 60 65 67 70 72 75 80 //
0
µg of protein/ml of culture
Specific activity (nmol/mg of protein/min)
by 40% in Ga-stressed bacteria compared to control cells (Figure 61).
6
Growth (hrs)
Specific activity of FUM in control membrane
Specific activity of FUM in gallium membrane
Specific activity of FUM in control regulation membrane
Growth profile of control culture
Growth profile of gallium culture
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)
133
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 Gastressed cells.
1
2
3
4
A
B
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.
134
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).
135
1
2
3
4
5
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.
136
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).
50
600
500
µg of protein/ml of culture
Specific activity (nmol/mg of protein/min)
45
40
35
400
30
25
300
20
200
15
10
100
5
0
0
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
Time of growth (hrs)
specific activity in control cells
growth profile of control culture
specific activity in Ga-stressed cells
growth profile of Ga-stressed culture
Figure 64: Specific activity of SDH (FADH2 formation) in control and 1 mM Gastressed 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
137
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).
1
2
Figure 65: BN PAGE detection of SDH in the membrane fraction of control and Gastressed 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
138
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)
139
In biological system, the synthesis of aspartate depends of AST, an enzyme
responsible for the transamination of oxaloacetate and glutamate to aspartate and αketoglutarate. 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)
140
600
120
500
100
400
80
300
60
200
40
20
100
0
0
µg of protein/ml of culture
Specific activity (nmol/mg of protein/min)
(Figure 66).
0 10 12 15 18 20 23 25 30 36 50 52 55 60 65 67 70 72 75 80
Growth (hrs)
Specific activity of AST in control soluble
Specific activity of AST in gallium soluble
Growth profile of control culture
Growth profile of gallium culture
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)
140
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 αketoglutarate is transaminated to aspartate by glutamate, the oxaloacetate produced is
converted to malate (Figure 67).
1
2
3
4
5
6
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
141
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
142
600
*
80
500
70
60
400
50
300
40
30
200
20
100
µg of protein/ml of culture
Specific activity (nmol/mg of protein/min)
90
10
0
0
0
10
12
15
18
20
23
25
30
36
50
52
55
60
65
67
70
72
75
80
Growth (hrs)
specific activity of MDH in control membrane
specific activity of MDH in gallium membrane
growth profile of control cells
growth profile of gallium cells
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).
143
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).
1
2
3
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.
144
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)
1
2
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.
145
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
Gallium
Toxicity
Interact with
lipids
Iron metabolism
ROS
Adaptation
Proteins e.g.
(NADH oxidase)
ETC components
Iron containing
enzymes
ROS
ROS
Detoxification
ROS
Siderophores
(β-hydroxyaspartate)
ROS scavengers
Diminished Iron
Requiring
metabolism
Iron binding protein
(storage)
ROS
146
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
147
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
GaO2˙+2 + 2H+
3
O2-˙
Ga+3
H2O2
1
2
HO-˙
Iron proteins
4
Fe+3 + H2O2
Catalase
NADPH
Fe+3
NADPH
H2O + O2
NADP+
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 αketoglutarate. 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.
148
Concomitantly, the activity of α-KGDH is reduced in these cells. The enzyme ICDHNADP+ 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, αketoglutarate and NADPH in the mitochondria, cytosol and peroxisome. Hence, αketoglutarate 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)
149
Figure 73: Metabolic reconfiguration to combat the oxidative stress generated by
gallium in P. fluorescens
isocitrate
ICDH
Ga
α-ketoglutarate + H2O2
α-KGDH
(inhibition
directly or
indirectly)
Isoenzyme
Increased protein
expression
Succinate + CO2
Decreased protein
expression
Decreased synthesis
Of NADH
Decrease ROS
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
150
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
Glutamate
(unchanged)
NAD+
Succinate
(increased)
CO2
H2O2
NH4+
NADH
HS-CoA
CO2
α-ketoglutarate
aspartate
NADH
Succinyl-CoA
(inhibited)
Glutamate
(unchanged)
NAD+
CO2
oxaloacetate
Succinic
Semialdehyde
(absent)
Hydroxyglutaric
Acid
(absent)
151
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 twofold. 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.
152
Figure 75: Inhibition of α-KGDH in Ga-stressed P. fluorescens
4-hydroxy-2-nonenal
Ga+3
H2O2
glutathionylation
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
153
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, αketoglutarate 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 αKGDH 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
Energy
Cofactor for
Hydroxylases
e.g.prolyl
hydroxylase
ROS
scavenger
α-ketoglutarate
DNA repair
e.g.AlkB
Gene
Regulator
e.g. ammonium
assimilation
154
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):
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
155
Labeled citrate NMR analysis
d citrate NMR a
N
HO
H
H
at 33 ppm and 181 ppm. The only possible explanation for this observation is that αketoglutarate 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.
ICDH
+
COOH
HO
ACN
COOH
COOH NAD NADH, H
+
COOH
COOH
COOH
ICDH
COOH HO COOH + +
COOH +
+
NADP
NADPH,
H
NAD
NADH, H
COOH
O
COOH
COOH
ICL
COOH
COOH
ICL
Ga
H 2O 2
NAD+ CoA
KGDH+
NADP
NADPH, H
NADH, CoA
H2O + CO2
Ga
H 2O 2
HOC COOH
COOH
+
COOH
H2O + CO2
HOC
= 13C-labelled
COOH
156
O
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. αKetoglultarate 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
157
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 αketoglutarate (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 NADPHgenerating 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
158
of this spatial arrangement would enable the α-ketoglutarate generated from the ICDHNADP+ 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
159
express a novel ICDH-NADP+ that may compensate for the lower activity of ACN in Gastressed 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
Gallium
Iron deprivation
Aconitase
inhibition
Catalase
inhibition
ICDH activity
Is increase
Induction of ICDH
isoenzyme
Inhibition of
Iron-containing
FUM A
Induction of iron-free
FUM C
Increase
α-ketoglutarate
generation for ROS
scavenging
Metabolome
effect
160
Gallium sequestration
Many bacteria including cyanobacteria and heterotrophic bacteria utilize
siderophores to bind iron in the environment as an uptake mechanism and to survive ironlimited 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.
161
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. αKetoglutarate 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 Gastressed 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.
162
Scheme 1: Ga-evoked metabolic shift operative in P. fluorescens
Gallium
Reduces
Bioavailable
iron
Increases ROS
Adaptation
ROS defense
Iron-limited
metabolism
Gallium
immobilization
α-ketoglutarate
β-Hydroxyaspartate
Detoxification
Glutamate
Aspartate
Citrate -Ga
Citrate
ROS
Oxaloacetate
cis-Aconitate
NADPH
X 1
2
α-ketoglutarate
Isocitrate
Malate
4
α-ketoglutarate
Fumarate
3
Succinate
6
Glutamate
X
H 2O 2
X
5
H 2O + O 2
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
163
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