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5.4: Oxidative Phosphorylation - Biology

5.4: Oxidative Phosphorylation - Biology


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Oxidative phosphorylation denotes the phosphorylation of ADP into ATP, utilizing the energy from successive electron transports (hence the “oxidative”). The basic concept is that oxidation of NADH, being highly exergonic, can generate the energy needed to phosphorylate ADP. Since oxidation of NADH by oxygen can potentially release 52 kCal/mol (218 kJ/mol), and the energy needed to phosphorylate ATP is approximately 7.5 kCal/mol (30.5 kJ/mol), we should be able to expect the formation of several ATP per oxidized NADH. Indeed, this is what happens, although not directly. As noted with the breakdown of glucose, a one-step oxidation would generate too much energy for cellular processes to handle, and most would be wasted. So instead of oxidizing NADH directly with O2, the electrons are transferred to a series of gradually lower-energy carriers until finally reaching oxygen. This sequence is the electron transport chain.

The electron transport chain is based on the activity of four major enzyme complexes (conveniently called complexes I-IV) embedded in the inner mitochondrial membrane, along with some small easily diffusible electron carriers to move the electrons from one complex to the next. These complexes are present in extremely high numbers as befits their necessity in generating energy, comprising nearly 75% of the inner membrane mass (in comparison, the plasma membrane of an average eukaryotic cell has a protein concentration closer to 50%). An overview of the process is shown in Figure (PageIndex{6}): as previously noted, electrons are stripped from NADH, and eventually end up on oxygen. As the electrons are moved to lower-energy carriers, energy is released and used to pump protons from the mitochondrial matrix into the intermembrane space.

Complex I is an NADH dehydrogenase. Shown in yellow in Figure (PageIndex{6}), its purpose is to remove a pair of electrons from NADH and transfer them onto ubiquinone (Coenzyme Q or CoQ), a small hydrophobic electron carrier that can then carry the electrons to complex III. This is a multistep process that involves first transferring the electrons onto an associated flavin mononucleotide (FMN) molecule, which then transfers the electrons to a set of iron-sulfur moieties connected to the enzyme complex itself (structure in Figure (PageIndex{7})). Finally, the electrons are moved onto ubiquinone. As these transfers occur, the energy that is released during these transfers powers the pumping of 4 H+ ions across the inner mitochondrial membrane. Complex I is inhibited by rotenone, a pesticide used primarily against insects and shes.

We’ll take a mental pass on complex II for now and hit it at the end of this roll call. The reasons will be apparent then.

Complex III is also known as the cytochrome bc1 complex (Figure (PageIndex{6}), purple). The purpose of this complex is to pass the electrons from ubiquinone onto cytochrome c. The use of ubiquinone is important here, because it is stable with either two, or just one, extra electron. Cytochrome c, on the other hand, can only carry one electron. So, this complex docks ubiquinone, and holds it until it has passed its first electron onto cytochome c, which then moves onto complex IV, and then its second electron onto another cytochrome c. With each transfer, two protons are pumped across the membrane.

Finally, cytochrome c drops the electron off to complex IV, cytochrome c oxidase (Figure (PageIndex{6}), red). Cytochrome c oxidase accomplishes the final step: transferring electrons onto oxygen atoms to make water. The really interesting thing about this process is that the enzyme must hold onto the electrons as they are transferred one at a time from cytochrome c, until it holds four electrons. Then, it can transfer one pair to each of the oxygen atoms in molecular oxygen (O2). It is very important to do this because transferring any less than all four electrons would lead to the creation of reactive oxygen species (ROS) that could cause damage to the enzymes and membranes of the mitochondria.

In fact, some well known poisons act at exactly this point. Both cyanide and carbon monoxide can bind with higher affinity than oxygen at the heme in complex IV. Since neither can accept electrons, the effect is just as though no oxygen was available.

Although cytochrome c oxidase is sometimes abbreviated COX, it is not the target of the COX-2 inhibitors that are used pharmaceutically in pain management, e.g. Bextra, Celebrex, or Vioxx. That refers to a family of enzymes known as the cyclooxygenases.

Oxygen is absolutely required. If oxygen is not available, there is no place to transfer the electrons, and very quickly, the electron transport chain is halted and carriers such as cytochrome c and CoQ cannot release their electrons and eventually there are no more available carriers. Similarly, when that happens, NAD+ is not regenerated, so the TCA cycle is also stuck. This leaves only the anaerobic non-oxygen-requiring glycolysis- fermentation cycle for generating ATP.

We now return to complex II (see Figure (PageIndex{10})). We mentioned complex II as succinate dehydrogenase when discussing the TCA cycle. It also participates in the electron transport chain by passing electrons to ubiquinone. However, rather than transferring electrons that originated from NADH like the other three complexes of the electron transport chain, the electrons originate from the covalently bound electron carrier FADH2 (flavin adenine dinucleotide), which received the electrons from succinate, as described in the TCA cycle section. Once the electrons have been passed to ubiquinone, it then moves on to complex III to drop off those electrons to cytochrome c, and the rest of the electron transport chain continues. FAD, the oxidized form of FADH2, is then ready to participate in the next redox cycle.

The purpose of this electron transport chain, with respect to ATP generation, is the pumping of H+ from the mitochondrial matrix into the intermembranous space. Since the concentration of protons is higher in the intermembrane space, it will take energy to move them against the concentration gradient, which is where our high-energy electrons come into the picture. As they move from one carrier to the next, they are moving from a higher to a lower energy state. This implies that some energy is lost from the electron, and some of that energy is tapped by the enzymes of the electron transport chain to move protons from the matrix to the intermembrane space.

There are two methods by which the protons are moved: the redox loop, and the proton pump. The proton pump, which is the method by which complex IV moves protons, is the easier to understand: H+ is bound on the matrix side of the enzyme in its reduced state (after it has received an electron), and a conformational shift occurs upon reoxidation to open the enzyme up to the intermembrane side, and the H+ is released. The redox loop, which occurs in complex I, and in complex III in a variation called the Q cycle, essentially posits that an initial redox center requires the binding of both the high energy electron and a proton from the matrix side of the membrane. When the electron is transferred to the next redox center in the chain, a proton is released to the intermembrane space.

Whatever the mechanism, what is the point of all this proton pumping? As you might suspect, using up energy to pump an ion against its concentration gradient isn’t done for the fun of it. Rather, this generates significant potential energy across the inner mitochondrial membrane. And, it so happens that there is an enzyme that can convert that energy into the physiologically useful chemical form of ATP. This enzyme is, not surprisingly, named ATP synthase (Figure (PageIndex{8})). It is also referred to in some texts as the F1F0-ATPase, based on its reverse activity (at the expense of ATP, it can pump protons), and the fact that it can be broken down into two major functional units: F1 which can hydrolyze but not synthesize ATP and is a soluble protein, and F0 which is an insoluble transmembrane protein.

The ATP synthase is an extraordinary example of an enzyme that transforms the energy inherent in a concentration gradient across a membrane into mechanical energy, and finally into chemical bond energy. It is descriptively called a “rotary engine” because the very generalized sequence of events is as follows: protons ow down their gradient through a proton channel subunit of the ATP synthase, in owing down the gradient, energy is released, this energy causes rotation of a multisubunit “wheel”-like subunit attached to a spindle/axle (g subunit) which also spins. The spinning of this asymmetrically shaped spindle unit causes conformational changes in the catalytic subunit (made of the a and b subunits) it is attached to, changing an ADP+Pi binding site to a catalytic site that can “squeeze” the molecules together into an ATP, and then finally open up to release the ATP (Figure (PageIndex{9})).

Of course, it isn’t quite that simple (Figure (PageIndex{8})). Starting with the initial movement of pro- tons, as they move from the intermembrane space into the ATP synthase, they enter a small hydrophilic channel (a) and then bind onto one of the c-subunits of the “water wheel” c-ring. Binding of the H+ to the c-subunit causes it to lose affinity for the a- subunit, allowing it to spin, and simultaneously causes a conformational change that essentially pushes off against the a-subunit, initiating the movement. Once it has spun around almost a complete turn, the H+ is positioned by another channel (b), which funnels it from the c-subunit into the matrix. The c-subunit structure is connected to an asymmetric spindle that is itself connected to the catalytic subunits.


The oxygen dependence of mitochondrial oxidative phosphorylation was measured in suspensions of isolated rat liver mitochondria using recently developed methods for measuring oxygen and cytochrome c reduction. Cytochrome-c oxidase (energy conservation site 3) activity of the mitochondrial respiratory chain was measured using an artificial electron donor (N,N,N′,N′-tetramethyl-p-phenylenediamine) and ascorbate to directly reduce the cytochrome c, bypassing sites 1 and 2. For mitochondrial suspensions with added ATP, metabolic conditions approximating those in intact cells and decreasing oxygen pressure both increased reduction of cytochrome c and decreased respiratory rate. The kinetic parameters [KM and maximal rate (VM)] for oxygen were determined from the respiratory rates calculated for 100% reduction of cytochrome c. At 22°C, the KM for oxygen is near 3 Torr (5 μM), 12 Torr (22 μM), and 18 Torr (32 μM) at pH 6.9, 7.4, and 7.9, respectively, and VM corresponds to a turnover number for cytochrome c at 100% reduction of near 80/s and is independent of pH. Uncoupling oxidative phosphorylation increased the respiratory rate at saturating oxygen pressures by twofold and decreased the KM for oxygen to <2 Torr at all tested pH values. Mitochondrial oxidative phosphorylation is an important oxygen sensor for regulation of metabolism, nutrient delivery to tissues, and cardiopulmonary function. The decrease in KM for oxygen with acidification of the cellular environment impacts many tissue functions and may give transformed cells a significant survival advantage over normal cells at low-pH, oxygen-limited environment in growing tumors.

the oxygen partial pressure (P o 2) and pH are interrelated parameters in the cellular environment that strongly influence many functions of the organism. These range from metabolic homeostasis and gene expression in individual cells to whole body functions, such as breathing and cardiovascular performance. There is also a wide variety of pathological conditions for which tissue oxygenation and pH are critically important, such as hypoxic-ischemic injury, tumor growth, peripheral vascular disease, and diabetic retinopathy. It is the metabolism of individual cells that first responds to change in oxygen pressure, and this response is then translated into metabolic and physiological changes in tissue biology. In most cells, mitochondrial oxidative phosphorylation is central to metabolism, serving as the principal source of ATP (>95%) and accounting for >95% of the total oxygen consumed. Any perturbation in the function of oxidative phosphorylation has an immediate impact on cellular metabolism. It follows that knowledge of the response of this metabolic pathway to alterations in oxygen pressure in the cellular environment is essential to understanding cellular and tissue physiology.

The literature on the oxygen dependence of mitochondrial oxidative phosphorylation provides two different and completely incompatible sets of data and paradigms. One, based largely on the studies by Chance and coworkers (5, 9, 19, 20, 25), is that the critical oxygen concentration for bioenergetic function of mitochondria is ∼0.05 Torr (0.08 μM). An important corollary of this lack of oxygen dependence >0.05 Torr is that oxidative phosphorylation does not contribute to oxygen-dependent regulation of cellular metabolism or tissue function under physiological conditions. The second is based on the data showing that changes in cytochrome c reduction and energy metabolism respond to P o 2 changes at oxygen pressures as high as 30 Torr (15, 35–37, 40, 41). This value is near the mean P o 2 in the interstitial space of tissue and the mixed venous oxygen pressure. The corollary of the latter observation is that the mitochondrial oxidative phosphorylation, in addition to being the metabolic “power plant” of the cell, is highly responsive to changes in oxygen pressure under physiological conditions and, therefore, has a key role in regulation of metabolism and of the nutrient delivery system.

The reported experimental differences may have occurred, in large part, due to technical limitations of measurements of oxygen and reduction of the cytochromes. In many of the original experiments, “work arounds” were used to compensate for the technical limitations, which likely introduced sources of systematic errors that, in some cases, led to erroneous conclusions. In the present paper, we report measurements using most up to date technology for determination of oxygen pressures and cytochrome c reduction. Our results support the second paradigm: that the oxygen dependence of oxidative phosphorylation extends as high as to the mean oxygen pressures in tissue at a physiological pH of 7.4. Furthermore, we show that the oxygen dependence becomes lower at more acidic pH. The latter observation has far-reaching consequences, one of which is that the transformed cells in solid tumors may have significant metabolic advantage over the neighboring normal cells by being able to out-compete them for the limited supply of oxygen.


Oxidative phosphorylation in Zymomonas mobilis

The obligately fermentative aerotolerant bacterium Zymomonas mobilis was shown to possess oxidative phosphorylation activity. Increased intracellular ATP levels were observed in aerated starved cell suspension in the presence of ethanol or acetaldehyde. Ethanolconsuming Z. mobilis generated a transmembrane pH gradient. ATP synthesis in starved Z. mobilis cells could be induced by external medium acidification of 3.5–4.0 pH units. Membrane vesicles of Z. mobilis coupled ATP synthesis to NADH oxidation. ATP synthesis was sensitive to the protonophoric uncoupler CCCP both in starved cells and in membrane vesicles. The H + -ATPase inhibitor DCCD was shown to inhibit the NADH-coupled ATP synthesis in membrane vesicles. The physiological role of oxidative phosphorylation in this obligately fermentative bacterium is discussed.

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Contents

Metabolites Edit

The matrix is host to a wide variety of metabolites involved in processes within the matrix. The citric acid cycle involves acyl-CoA, pyruvate, acetyl-CoA, citrate, isocitrate, α-ketoglutarate, succinyl-CoA, fumarate, succinate, L-malate, and oxaloacetate. [2] The urea cycle makes use of [[ornithineL-ornithine]], carbamoyl phosphate, and L-citrulline. [4] The electron transport chain oxidizes coenzymes NADH and FADH2. Protein synthesis makes use of mitochondrial DNA, RNA, and tRNA. [5] Regulation of processes makes use of ions (Ca 2+ /K + /Mg + ). [6] Additional metabolites present in the matrix are CO2, H2O, O2, ATP, ADP, and Pi. [1]

Enzymes Edit

Inner membrane components Edit

The inner membrane is a phospholipid bilayer that contains the complexes of oxidative phosphorylation. which contains the electron transport chain that is found on the cristae of the inner membrane and consists of four protein complexes and ATP synthase. These complexes are complex I (NADH:coenzyme Q oxidoreductase), complex II (succinate:coenzyme Q oxidoreductase), complex III (coenzyme Q: cytochrome c oxidoreductase), and complex IV (cytochrome c oxidase). [6]

Inner membrane control over matrix composition Edit

The electron transport chain is responsible for establishing a pH and electrochemical gradient that facilitates the production of ATP through the pumping of protons. The gradient also provides control of the concentration of ions such as Ca 2+ driven by the mitochondrial membrane potential. [1] The membrane only allows nonpolar molecules such as CO2 and O2 and small non charged polar molecules such as H2O to enter the matrix. Molecules enter and exit the mitochondrial matrix through transport proteins and ion transporters. Molecules are then able to leave the mitochondria through porin. [9] These attributed characteristics allow for control over concentrations of ions and metabolites necessary for regulation and determines the rate of ATP production. [10] [11]

Citric acid cycle Edit

Following glycolysis, the citric acid cycle is activated by the production of acetyl-CoA. The oxidation of pyruvate by pyruvate dehydrogenase in the matrix produces CO2, acetyl-CoA, and NADH. Beta oxidation of fatty acids serves as an alternate catabolic pathway that produces acetyl-CoA, NADH, and FADH2. [1] The production of acetyl-CoA begins the citric acid cycle while the co-enzymes produced are used in the electron transport chain. [11]

All of the enzymes for the citric acid cycle are in the matrix (e.g. citrate synthase, isocitrate dehydrogenase, α-ketoglutarate dehydrogenase, fumarase, and malate dehydrogenase) except for succinate dehydrogenase which is on the inner membrane and is part of protein complex II in the electron transport chain. The cycle produces coenzymes NADH and FADH2 through the oxidation of carbons in two cycles. The oxidation of NADH and FADH2 produces GTP from succinyl-CoA synthetase. [2]

Oxidative phosphorylation Edit

NADH and FADH2 are produced in the matrix or transported in through porin and transport proteins in order to undergo oxidation through oxidative phosphorylation. [1] NADH and FADH2 undergo oxidation in the electron transport chain by transferring an electrons to regenerate NAD + and FAD. Protons are pulled into the intermembrane space by the energy of the electrons going through the electron transport chain. Four electrons are finally accepted by oxygen in the matrix to complete the electron transport chain. The protons return to the mitochondrial matrix through the protein ATP synthase. The energy is used in order to rotate ATP synthase which facilitates the passage of a proton, producing ATP. A pH difference between the matrix and intermembrane space creates an electrochemical gradient by which ATP synthase can pass a proton into the matrix favorably. [6]

Urea cycle Edit

The first two steps of the urea cycle take place within the mitochondrial matrix of liver and kidney cells. In the first step ammonia is converted into carbamoyl phosphate through the investment of two ATP molecules. This step is facilitated by carbamoyl phosphate synthetase I. The second step facilitated by ornithine transcarbamylase converts carbamoyl phosphate and ornithine into citrulline. After these initial steps the urea cycle continues in the inner membrane space until ornithine once again enters the matrix through a transport channel to continue the first to steps within matrix. [12]

Transamination Edit

α-Ketoglutarate and oxaloacetate can be converted into amino acids within the matrix through the process of transamination. These reactions are facilitated by transaminases in order to produce aspartate and asparagine from oxaloacetate. Transamination of α-ketoglutarate produces glutamate, proline, and arginine. These amino acids are then used either within the matrix or transported into the cytosol to produce proteins. [7] [13]

Regulation Edit

Regulation within the matrix is primarily controlled by ion concentration, metabolite concentration and energy charge. Availability of ions such as Ca 2+ control various functions of the citric acid cycle. in the matrix activates pyruvate dehydrogenase, isocitrate dehydrogenase, and α-ketoglutarate dehydrogenase which increases the reaction rate in the cycle. [14] Concentration of intermediates and coenzymes in the matrix also increase or decrease the rate of ATP production due to anaplerotic and cataplerotic effects. NADH can act as an inhibitor for α-ketoglutarate, isocitrate dehydrogenase, citrate synthase, and pyruvate dehydrogenase. The concentration of oxaloacetate in particular is kept low, so any fluctuations in this concentrations serve to drive the citric acid cycle forward. [2] The production of ATP also serves as a means of regulation by acting as an inhibitor for isocitrate dehydrogenase, pyruvate dehydrogenase, the electron transport chain protein complexes, and ATP synthase. ADP acts as an activator. [1]

Protein synthesis Edit

The mitochondria contains its own set of DNA used to produce proteins found in the electron transport chain. The mitochondrial DNA only codes for about thirteen proteins that are used in processing mitochondrial transcripts, ribosomal proteins, ribosomal RNA, transfer RNA, and protein subunits found in the protein complexes of the electron transport chain. [15] [16]


5.4: Oxidative Phosphorylation - Biology

Human body is continuously exposed to different types of agents that results in the production of reactive species called as free radicals (ROS/RNS) which by the transfer of their free unpaired electron causes the oxidation of cellular machinery. In order to encounter the deleterious effects of such species, body has got endogenous antioxidant systems or it obtains exogenous antioxidants from diet that neutralizes such species and keeps the homeostasis of body. Any imbalance between the RS and antioxidants leads to produce a condition known as “oxidative stress” that results in the development of pathological condition among which one is diabetes. Most of the studies reveal the inference of oxidative stress in diabetes pathogenesis by the alteration in enzymatic systems, lipid peroxidation, impaired Glutathione metabolism and decreased Vitamin C levels. Lipids, proteins, DNA damage, Glutathione, catalane and superoxide dismutase are various biomarkers of oxidative stress in diabetes mellitus. Oxidative stress induced complications of diabetes may include stroke, neuropathy, retinopathy and nephropathy. The basic aim of this review was to summarize the basics of oxidative stress in diabetes mellitus.


Materials and methods

To identify orthologous OXPHOS genes and their duplications in D. pseudoobscura and A. gambiae, contigs from BCM [13] and scaffolds from AnoBase [21] were searched using TBLASTN with the D. melanogaster OXPHOS peptides listed in the MitoDrome database [19] as queries.

Amino-acid sequence identity and similarity values were obtained from pairwise alignments using the Needleman-Wunsch global alignment algorithm at the EMBL-EBI server [60]. Multiple sequence alignments of the OXPHOS amino-acid and coding sequences and visualization of the dendrograms were obtained using the MultAlin 5.4.1 software [61] from MultAlin server [62].

The genomic sequence of each gene was manually searched for intron-exon boundaries and the predicted mRNA sequence reconstructed in silico. A. gambiae mRNAs were assembled by overlapping ESTs extracted from AnoBase [21].

We have named each newly identified A. gambiae gene with the four-letter code 'agEG' followed by the last four or five digits of its Ensembl [36] gene number, excluding the multiple zeros of the prefix the D. pseudoobscura genes were named with the code 'DpseCG' followed by the Celera number of their D. melanogaster counterparts.

The D. pseudoobscura OXPHOS genes investigated here were assigned a chromosomal location where possible, using the putative chromosomal assignments available at BCM [13] for the majority of the large D. pseudoobscura contigs. We also utilized the Ensembl mosquito genome server [36] to identify and visualize the chromosomal location of the A. gambiae annotated OXPHOS DNA sequences.

The D. melanogaster EST database, available from the National Center for Biotechnology Information (NCBI) contains ESTs from cDNA libraries obtained from different developmental stages and body parts. The relative abundance of the transcripts of duplicate or triplicate D. melanogaster OXPHOS genes was defined by counting their cognate ESTs in non-normalized cDNA libraries generated by the Berkeley Drosophila Genome Project (BDGP) [43] from embryos (LD), larvae/pupae (LP), and adult ovary (GM), head (GH) and testes (AT), and also the ESTs from adult testes generated at the National Institute of Diabetes and Digestive and Kidney Diseases (NIDDK) [63]. ESTs from BDGP normalized EST libraries generated from head (RH) and embryos (RE) were also considered. The relative abundance of the transcripts of duplicate or triplicate A. gambiae OXPHOS genes was defined by counting their cognate ESTs in all libraries recovered from the Anobase server [21]. Since the number of sequences in the EST databases changes as new EST sequences are added, our values are calculated on the EST sequences present in the databases as of July 2004.

The list of D. melanogaster P-insertion OXPHOS mutants is reported in the MitoComp website [22] and was mostly compiled using information from FlyBase [42] and from the BDGP P-Element Gene Disruption Project [43].


Oxidative stress-induced phosphorylation, degradation and aggregation of α-synuclein are linked to upregulated CK2 and cathepsin D

Intracellular accumulation of α-synuclein (α-Syn) as filamentous aggregates is a pathological feature shared by Parkinson's disease, dementia with Lewy bodies and multiple system atrophy, referred to as synucleinopathies. To understand the mechanisms underlying α-Syn aggregation, we established a tetracycline-off inducible transfectant (3D5) of neuronal lineage overexpressing human wild-type α-Syn. α-Syn aggregation was initiated by exposure of 3D5 cells to FeCl2. The exposure led to formation of α-Syn inclusions and oligomers of 34, 54, 68 kDa and higher molecular weights. The oligomers displayed immunoreactivity with antibodies to the amino-, but not to the carboxyl(C)-, terminus of α-Syn, indicating that C-terminally truncated α-Syn is a major component of oligomers. FeCl2 exposure also promoted accumulation of S129 phosphorylated monomeric α-Syn (Pα-Syn) and casein kinase 2 (CK2) however, G-protein-coupled receptor kinase 2 was reduced. Treatment of FeCl2-exposed cells with CK2 inhibitors (DRB or TBB) led to decreased formation of α-Syn inclusions, oligomers and Pα-Syn. FeCl2 exposure also enhanced the activity/level of cathepsin D. Treatment of the FeCl2-exposed cells with pepstatin A or NH4Cl led to reduced formation of oligomers/inclusions as well as of ∼ 10 and 12 kDa truncated α-Syn. Our results indicate that α-Syn phosphorylation caused by FeCl2 is due to CK2 upregulation, and that lysosomal proteases may have a role in producing truncated α-Syn for oligomer assembly.

Fig. S1. FeCl2 treatment inhibits proteasomes. (A) Immunoblotting of cell lysates with anti-ubiquitin antibody (Mab 1510, Chemicon). FeCl2-treated samples displayed more high-molecular-weight (HMW) proteins with ubiquitin (ubiq) immunoreactivities. The arrow indicates ubiquitin monomer. (B) Quantitative analysis of ubiquitin-immunoreactive proteins of molecular weight > 75 kDa. Student?s t-test, N = 6, *P < 0.05. (C) Proteasomal activity assay using fluorogenic peptides as substrate. Chymotrypsine-like activity was lower in Fe(+) than Fe(?) controls, and higher in Syn(+) than Syn(?) cells without the FeCl2 treatment, N = 6, *P < 0.05. The specificity of activity assay was verified by analysis of lysates in the presence of the proteasome inhibitor, epoxomicin.

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Extended Data Fig. 1 Activity of PolD and PolK with 5′-OAP.

Shown are HPLC chromatograms at 260 nm for a full reaction with PolK, a full reaction with PolD, a control without enzyme, and a control without α-KG. No consumption of 5′-OAP was detected. The results were reproducible in at least two independent assays.

Extended Data Fig. 2 Activity of PolQ2 with EP-UMP.

Shown are HPAEC chromatograms at 260 nm for a control without the enzyme (trace i) and the complete reaction (trace ii). No consumption of EP-UMP is detected. The results were reproducible in at least two independent assays.

Extended Data Fig. 3 Activity of PolD, PolK, and NikM with OABP.

Shown are HPAEC chromatograms at 260 nm for a control without the enzyme (trace i), an assay with PolD (ii), an assay with PolK (iii), and an assay with NikM (iv). No consumption of OABP was observed after 24 hours. The results were reproducible in at least two independent assays.

Extended Data Fig. 4 Activity assays of PolJ, NikK, and PolK with 2’-HAP.

a. PolJ phosphatase (10 μM) was incubated with 2′-HAP (100 μM) in 50 mM Tris pH 8.0 supplemented with 10 mM MgCl2 at 25 °C for 18 hrs. b. NikK aminotransferase (30 μM) was incubated with 2′-HAP (200 μM) in 200 mM Tris pH 9.0 supplemented with 1 mM MgCl2 and 10 mM l -Glu at 25 °C for 2 hrs. c. PolK oxygenase (25 μM) was incubated with 2′-HAP (200 μM) in 150 mM NaCl, 50 mM Tris pH 7.5 supplemented with 1 mM Fe 2+ , 1 mM ascorbate, 200 μM α-KG at 25 °C for 18 hrs. No product formation was detected. The results were reproducible in at least two independent assays.

Extended Data Fig. 5 Activity assays of PolK with KOAP.

PolK (40 μM) was incubated with KOAP (100 μM) in the presence of 100 μM Fe 2+ , 2 mM ascorbate, and 1 mM α-KG for 20 hours at 25 °C. Even after prolonged incubation (20 hours), no product is observed. The results were reproducible in at least two independent assays.

Extended Data Fig. 6 Activity of PolD with KOAP.

PolD (40 μM) was incubated with KOAP (100 μM) in the presence of 100 μM Fe(II), 1 mM ascorbate, and 2 mM α-KG for 2 hours at 25 °C. No product formation or KOAP consumption. For comparison, under similar conditions, PolD completed the conversion of AHOAP to AHAP in <15 minutes. The results were reproducible in at least two independent assays.

Extended Data Fig. 7 Stereochemistry of AHOAP.

Coupling constants for possible stereochemistry of C4′, C5′, C6′ and C7′. Experimental evidence of JH4’-H5’ = 3.5 Hz, JH5’-H6’ = 10.5 Hz and JH6’-H7’ = 5.6 Hz for AHOA is most consistent with the stereochemistry of 4′-S, 5′-S, 6′-S, 7′-R indicating that AHOA and AHOAP have 4′-S, 5′-S, 6′-S, 7′-R stereochemistry. Dihedral angles were calculated with ChemDraw Professional v19.0 and ChemDraw 3D v19.0 (PerkinElmer Informatics).

Extended Data Fig. 8 Comparison of the rates of reactions between PolD + AHOAP vs. PolD + 2′-OAP.

The PolD assays with AHOAP were performed with 10 μM PolD, 0.1mM Fe 2+ , 2 mM ascorbate, 100 μM AHOAP in 150 mM NaCl, 50 mM Tris pH 7.6, 10% glycerol. The PolD assays with 2′-OAP were performed with 30μM PolD, 1mM ascorbate, 0.5 mM Fe 2+ , 200 μM 2’-OAP in 150 mM NaCl, 50 mM Tris pH 7.6.

Extended Data Fig. 9 LC-HRMS analysis of culture media of S. cacaoi ΔpolQ2, wt, and ΔpolQ2 + polQ2.

Shown are EICs (calculated m/z for [M+H] + ± 5 ppm) for polyoxins A, B, D, F, G, H, I, and J. No polyoxin production was detected in ΔpolQ2 (a), polyoxin A, B, F, and G were found in the wt strain (b), and polyoxin A and F were detected in the ΔpolQ2 + polQ2 strain (c). The observations were reproducible for 2-3 different clones for each mutant strain. The culture was repeated twice for each clone.

Extended Data Fig. 10 Proposed divergent biosynthesis of antifungal nucleoside natural products.

Proposed tailoring of sugar size by oxidative C-C bond cleavage by PolD homologs.


MATERIALS AND METHODS

Materials

Rabbit polyclonal antibodies against KHK-A and KHK-C and KHK-A pS80 and p62 pS28 antibodies were obtained from Signalway Biotechnology (Pearland, TX). The antibody generation procedures were described previously (32). Normal mouse immunoglobulin G (IgG) (sc-2025), TRIM21 (sc-25351), GST (sc-138), Nrf2 (sc-13032), tubulin (sc-8035), ERK1/2 (sc-514302), pERK1/2 (Thr 202 /Tyr 204 ) (sc-16982), PRPS1/2 (sc-100822), and pan–KHK-A/C (sc-377411) antibodies were obtained from Santa Cruz Biotechnology (Santa Cruz, CA). Rabbit antibodies against mitogen-activated protein kinase-activated protein kinase 2 (MAPK/APK2) (#3042), MAPK/APK2 pT222 (#3316), c-Jun (#9165), c-Jun pS73 (#9164), Trx1 (#2429), AMPKα (#5831), AMPKα pT172 (#2535), ACC1 (#3676), ACC1 pS79 (#11818), LC3B (#3868), cleaved PARP (Asp 214 ) (#9541), and anti-rabbit IgG conformation-specific secondary antibody (L27A9) (#3678) were purchased from Cell Signaling Technology (Danvers, MA). BSO (B2515), NAC (A7250), Trolox (238813), MG132 (M7449), mouse monoclonal anti-Flag (F1804), rabbit anti-Flag (F7425), and anti-His (SAB1305538) antibodies were purchased from Sigma-Aldrich (St. Louis, MO). Anti-Flag M2 agarose beads and [γ- 32 P]ATP were purchased from MP Biochemicals (Santa Ana, CA). Active AMPK proteins (#P47-10H) were obtained from SignalChem (Richmond, BC, Canada). A769662 was purchased from Tocris Bioscience (Avonmouth, Bristol, UK). Rabbit anti-Ki67 antibody (#AB9260) and compound C were obtained from Millipore (Billerica, MA). Anti-Keap1 (ab226997) and anti-p62 (ab56416) antibodies were obtained from Abcam (Cambridge, MA). Ub-K63 antibody (HWA4C4) was purchased from Novus (Littleton, CO). Anti–proliferating cell nuclear antigen (PCNA) (#610665) and hypoxia-inducible factor 1α (#610958) antibodies were obtained from BD Biosciences (Bedford, MA). An anti-PRPS1 T225 phosphorylation antibody was generated as previously described (32). U0126, SP600125, and SB203580 were purchased from EMD Biosciences (San Diego, CA). A fluorescein isothiocyanate (FITC)–labeled antibody against 5-bromo-2′-deoxyuridine (BrdU) (BU20A, 11-5071) was purchased from eBioscience.

DNA construction and mutagenesis

PCR-amplified human p62 was cloned into pcDNA3.1/hygro(+)-Flag, pcDNA3-HA, or pColdI (His) vector. Human AMPKα1 was cloned into pcDNA3-HA vector. The construction of Flag/V5–rKHK-A or Flag/V5–rKHK-C, GST–KHK-A or GST–KHK-C, Flag-rKHK G257R, His-PRPS1, and KHK short hairpin RNA (shRNA) was described previously (32). Nrf2 CA (with the deletion of amino acids 1 to 89) was cloned into pcDNA3.1/puro(+)-HA. PINK1 and SQSTM1 shRNA were constructed via ligation of an oligonucleotide targeting human PINK1 (5′-GCTGGAGGAGTATCTGATAGG-3′) and p62 (5′-ACTGGACCCATCTGTCTTCAA-3′) into an Xho I–/Mlu I–digested pGIPZ vector, respectively. Flag-, V5-, or GST-tagged rKHK-A S80A, Flag–rKHK-A S80E, Flag-p62 S28E, Flag-p62 S28A, His–p62 S28A, Flag–p62 K7R, and all the shRNA-resistant p62 constructs containing nonsense mutations of C1017T, T1020C, and G1023A were constructed using the QuikChange Site-Directed Mutagenesis Kit (Stratagene, La Jolla, CA). Vectors expressing GFP-LC3, HA-ubiquitin-WT, HA-ubiquitin-K48, and HA-ubiquitin-K63 were purchased from Addgene.

Cell lines and cell culture conditions

LO2, Hep3B, and Huh7 cells were obtained from the American Type Culture Collection and were maintained in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% dialyzed bovine calf serum (HyClone). AMPKα1 and AMPKα2 double-knockout MEFs (a gift from B. Viollet, Institut Cochin, Paris, France) were maintained in DMEM supplemented with 10% fetal bovine serum (FBS). No cell lines used in this study were found in the database of commonly misidentified cell lines maintained by the International Cell Line Authentication Committee and National Center for Biotechnology Information BioSample. Cell lines were authenticated by short tandem repeat profiling and were routinely tested for mycoplasma contamination. Cells were plated at a density of 4 × 10 5 per 60-mm dish or 1 × 10 5 per well of a six-well plate 18 hours before transfection. The transfection procedure was performed as previously described (36).

Immunoprecipitation and immunoblotting analysis

The extraction of proteins using a modified buffer from cultured cells was followed by immunoprecipitation and immunoblotting using corresponding antibodies, as described previously (37).

GST pulldown assay

Equal amounts of purified His-tagged protein (200 ng per sample) were incubated with GST fusion proteins (100 ng) on glutathione (GSH) agarose beads in a modified binding buffer [50 mM tris-HCl at pH 7.5, 1% Triton X-100, 150 mM NaCl, 1 mM dithiothreitol (DTT), 0.5 mM EDTA, 100 μM phenylmethylsulfonyl fluoride, 100 μM leupeptin, 1 μM aprotinin, 100 μM sodium orthovanadate, 100 μM sodium pyrophosphate, and 1 mM sodium fluoride]. The glutathione agarose beads were then washed four times with binding buffer and then subjected to immunoblotting analysis, as previously described (38).

Purification of recombinant proteins

WT His-p62, His-p62 S80A, GST–KHK-A, GST–KHK-C, GST–KHK-A S80A, and GST–KHK-A G257R were expressed in bacteria and purified, as described previously (39).

In vitro kinase assay

The kinase reactions were performed, as described previously (40). Briefly, purified active AMPK proteins (including AMPKα1/β1/γ1) were incubated with purified GST-tagged KHK-A, KHK-C, or GST–KHK-A S80A mutant (100 ng) in 25 μl of kinase buffer <50 mM tris-HCl (pH 7.5), 100 mM KCl, 50 mM MgCl2, 1 mM Na3VO4, 1 mM DTT, 5% glycerol, 0.2 mM AMP, 0.5 mM ATP, and 10 μCi [γ- 32 P]ATP> at 25°C for 1 hour. To test p62 phosphorylation in vitro, KHK-A or KHK-C on the glutathione agarose beads was washed in phosphate-buffered saline (PBS) five times after in vitro AMPK kinase assay and then incubated with or without bacterially purified His-p62 or His–p62 S28A in the presence of [γ- 32 P]ATP. The reaction was terminated by adding SDS–polyacrylamide gel electrophoresis loading buffer and heating at 100°C for 5 min.

MS analysis

In vitro KHK-A phosphorylation by AMPK and p62 phosphorylation by KHK-A were performed. The protein samples were digested and analyzed by LC-MS/MS on an Orbitrap-Elite mass spectrometer (Thermo Fisher Scientific, Waltham, MA), as described previously (41).

CRISPR-Cas9–mediated genome editing

Genomic mutations were introduced into cells using the CRISPR-Cas9 system, as described previously (41). Single-guide RNAs (sgRNAs) were designed to target the genomic area adjacent to the KHK-A S80A and p62 S28A mutation sites using the CRISPR design tool (http://crispr.mit.edu/). The annealed guide RNA oligonucleotides were inserted into a PX458 vector (Addgene, Cambridge, MA) digested with the Bbs I restriction enzyme (42). Cells were seeded at 60% confluence, followed by cotransfection of sgRNAs (0.5 μg) and single-stranded donor oligonucleotide (ssODN) (10 pmol) as a template to introduce mutations. Twenty-four hours after transfection, cells were trypsinized and diluted so that cells in single were seeded into 96-well plates. Genomic DNA was extracted from GFP-positive cells, followed by sequencing of the PCR products spanning the mutation sites. sgRNA targeting sequence for KHK-A S80A: 5′-TGACCTCCGCCGCTATTCTG-3′, ssODN sequence for KHK-A S80A: 5′-CCTCCAGTCCCCAAAACCCTTGTCGAACTGCACCCCCTTCGGGTTACTCCGCCCTAGCAGTTTTGTCCTGGATGACCTCCGaCGaTAcgCTGTGGACCTACGCTACACAGTCTTTCAGACCACAGGCTCCGTCCCCATCGCCACGGTCATCATCAACGAGGCC-3′, sgRNA targeting sequence for p62 S28A: 5′-CTGCTGCAGCCCCGAGCCTG-3′, and ssODN sequence for p62 S28A: 5′-GCGTCGCTCACCGTGAAGGCCTACCTTCTGGGCAAGGAGGACGCGGCGCGCGAGATTCGCCGCTTCAGCTTCTGCTGCgcCCCaGAaCCaGAGGCGGAAGCCGAGGCTGCGGCGGGTCCGGGACCCTGCGAGCGGCTGCTGAGCCGGGTGGCCGCCCTGTTCC-3′.

Genotyping was performed by sequencing PCR products amplified from the following primers: KHK-A forward: 5′-TTAGGACTGGGAGGGACTGA-3′, KHK-A reverse: 5′-GTGAGGGAACTGATTGAGCC-3′, p62 forward: 5′-CGACCTAGCAGCCTCCTGA-3′, and p62 reverse: 5′-CCTTGGTCACCACTCCAGTCA-3′.

Measurement of intracellular NADPH and NADP + levels

The intracellular levels of NADPH and total NADP (which means NADPH and NADP + ) were measured according to previously described methods (43, 44). Briefly, 2 × 10 6 cells were lysed in 400 μl of extraction buffer containing 20 mM nicotinamide, 20 mM NaHCO3, and 100 mM Na2CO3 and centrifuged for 20 min using maximum speed. The supernatant was kept on ice in the dark until use. To extract NADPH, 150 μl of the supernatant was incubated at 60°C for 30 min because heating destroys the oxidized form (NADP + ) while having no effect on the reduced form (NADPH). Next, 160 μl of NADP-cycling buffer [100 mM tris-HCl (pH 8.0), 0.5 mM thiazolyl blue tetrazolium blue (MTT), 2 mM phenazine ethosulfate, and 5 mM EDTA] containing glucose-6-phosphate dehydrogenase (1.3 U) was added to a 96-well plate containing 20 μl of sample. After a 1-min incubation in the dark at 30°C, 20 μl of glucose 6-phosphate (10 mM) was added to the mixture, and the change in absorbance at 570 nm was measured every 30 s for 4 min at 30°C with a microplate reader. The concentration of NADP + was calculated by subtracting NADPH from the total NADP.

Measurement of intracellular ROS

The fluorescence probe 2′,7′-dichlorofluorescin diacetate (DCFDA) (Abcam) was used to measure intracellular ROS following the manufacturer’s instructions. Briefly, 2 × 10 4 cells were seeded in a clear-bottom 96-well plate. After treatment, cells were washed with Hanks’ balanced salt solution, followed by incubation with DCFDA (25 μM)–containing HBSS for 30 min at 37°C in the dark. Cells were washed, and green fluorescence was measured with a Synergy HT microplate reader (BioTek) at 485/535 nm.

GSH/GSSG detection

GSH and GSSG concentrations were measured by a GSH/GSSG detection kit (catalog no. S0053), according to the manufacturer’s instructions (Beyotime).

ARE-luciferase assay

Three copies of the ARE (5′-GTGACAAAGCAATCCCGTGACAAAGCAATCCCGTGACAAAGCAATA-3′) were cloned into a pGL3-basic luciferase reporter plasmid. The same amounts of ARE-luciferase reporter together with KHK plasmids were transfected into cells. Each transfection included the same amount of Renilla, which was used to normalize the transfection efficiency. Twenty-four hours after transfection, cells were treated with or without hypoxia for 6 hours. The luciferase activity in cell lysates was measured with the luciferase assay system as previously described (26).

Immunofluorescence analysis

Cells were fixed and incubated with primary antibodies at a dilution of 1:100, fluorescence dye–conjugated secondary antibodies, and 4′,6-diamidino-2-phenylindole (DAPI), according to standard protocols. Immunofluorescent microscopic images of the cells were obtained and viewed with an IX81 confocal microscope system (Olympus America).

Subcellular fractionation

Nuclear fractions were isolated from cells using a nuclei isolation kit (NUC201-1KT) according to the manufacturer’s instructions (Sigma-Aldrich).

Real-time PCR

Total RNA was extracted from cells and tissue specimens using TRIzol according to the manufacturer’s instructions (Invitrogen). A total of 1 mg per sample RNA was used for complementary DNA synthesis using a TaqMan Reverse Transcription Reagents kit (Applied Biosystems). Quantitative real-time PCR (qRT-PCR) was carried out in the 7500 real-time PCR system (Applied Biosystems) using an SYBR Premix Ex Taq kit (TAKARA). The following primers were used for qRT-PCR: GSTM1, 5′-TTTGTCCTGCCCACGTTTCT-3′ and 5′-TCAAAGTCGGGAGCGTCAC-3′ HO-1, 5′-GGTCAGGTGTCCAGAGAAGG-3′ and 5′-CTTCCAGGGCCGTGTAGATA-3′ PGD, 5′-AAAGATCCGGGACAGTGCT-3′ and 5′-CACCGAGCAAAGACAGCTT-3′ GCLC, 5′-GTGGACGAGTGCAGCAAG-3′ and 5′-GTCCAGGAAATACCCCTTCC-3′ NQO1, 5′-ATCCTGCCGAGTCTGTTCTG-3′ and 5′-AGGGACTCCAAACCACTGC-3′ UGT1A1, 5′-AACAAGGAGCTCATGGCCTCC-3′ and 5′-GTTCGCAAGATTCGATGGTCG-3′ and glyceraldehyde-3-phosphate dehydrogenase, 5′-AGCCACATCGCTCAGACAC-3′ and 5′-GCCCAATACGACCAATCC-3′.

DSP cross-linking

Cells with less than 80% confluence were washed twice with ice-cold washing buffer (10 mM Na2HPO4, 1.8 mM KH2PO4, 137 mM NaCl, 2.7 mM KCl, 0.1 mM CaCl2, and 1 mM MgCl2) and incubated in washing buffer with DSP (0.4 mg/ml) at 4°C for 2 hours. Cells were lysed in immunoprecipitation lysis buffer with 1% SDS and sonicated for 10 s. For immunoprecipitation, the lysate was diluted with immunoprecipitation lysis buffer at a ratio of 1:10 and subjected to immunoprecipitation assay.

KHK activity assay and 13 C-labeled F1P detection

KHK activity was measured by a method described previously (32). We added Roche cOmplete Protease Inhibitor Cocktail and Roche PhosSTOP to the reaction mixture containing 140 mg of protein of Huh7 cell lysates and incubated the mixture at 37°C for 3 hours. Spectrophotometric absorbance measurements of the remaining fructose were detected in 96-well plates at 515 nm using a SpectraMax plate reader (Molecular Devices).

The culture media of Huh7 cells were supplied with 13 C-labeled fructose for 6 hours, and then the cells were washed with PBS and whole-cell culture plates were snap-frozen on dry ice. Metabolites were extracted twice from plates with 70% ethanol (75°C). Targeted analysis 13 C-labeled F1P pattern was performed by ion-pairing, reverse-phase, ultra-performance LC-MS/MS on a Thermo Quantum Ultra instrument (Thermo Fisher Scientific), as previously described (32).

Apoptosis analysis

Cells (1 × 10 5 ) were seeded into a six-well plate overnight. Cells were then fixed by direct addition of 12% formaldehyde into the culture medium. After fixation, the cells were stained with DAPI (1 μg/ml) for 5 min and washed by PBS. The cells with condensed and/or fragmented chromatin were indicative of apoptosis and counted.

Cell viability analysis

Cells (2 × 10 5 ) were plated in DMEM with 10% FBS. The viable cells were stained with trypan blue (0.5%) and counted using Beckman Coulter.

Cell proliferation assay

Cells were incubated with BrdU (50 μM) for 1 hour before harvest. Collected cells were fixed in 70% ethanol at 4°C for 1 hour, followed by incubation with 2 N HCl/0.5% Triton X-100 for 30 min and 0.1 M borate sodium for 2 min. After incubation with an anti–BrdU-FITC antibody (eBioscience) and washing five times in PBS, the BrdU incorporation rate was analyzed by flow cytometry.

Terminal deoxynucleotidyl transferase–mediated deoxyuridine triphosphate nick end labeling assay

Mouse tumor tissues were sectioned at 5 μm thickness. Apoptotic cells were counted using the DeadEnd Colorimetric TUNEL System (Promega) according to the manufacturer’s instructions.

IHC analysis and histologic evaluation of human HCC specimens

Mouse tumor samples were fixed and prepared for staining. The samples were stained with Mayer’s hematoxylin and eosin (H&E) (BioGenex). The slides were then mounted using Universal Mount (Research Genetics).

Liquid nitrogen–frozen human HCC and adjacent matched nontumor tissue samples from 90 patients were obtained from Eastern Hepatobiliary Surgery Hospital in Shanghai, People’s Republic of China. The use of human HCC specimens and the database was approved by the Hospital’s Institutional Review Board. All tissue samples were collected in compliance with informed consent policy. Sections of paraffin-embedded human HCC tissue were stained with antibodies against Ki67, phospho–KHK-A S80, phospho-p62 S28, Nrf2, or nonspecific IgG as a negative control. The tissue sections were quantitatively scored according to the percentage of positive cells and staining intensity, as described previously (41). The following proportion scores were assigned to the sections: 0 if 0% of the tumor cells exhibited positive staining, 1 for 0 to 1%, 2 for 2 to 10%, 3 for 11 to 30%, 4 for 31 to 70%, and 5 for 71 to 100%. In addition, the staining intensity was rated on a scale of 0 to 3: 0, negative 1, weak 2, moderate and 3, strong. The proportion and intensity scores were then combined to obtain a total score (range, 0 to 8), as described previously (41). Scores were compared with overall survival duration, defined as the time from date of diagnosis to death or last known follow-up examination. All patients had received standard therapies after surgery.

Animal studies

One million Huh7 cells or Huh7 cells with knock-in of KHK-A S80A or p62 S28A were collected in 20 μl of DMEM with 33% Matrigel and intrahepatically injected into livers of 6-week-old male BALB/c athymic nude mice. The injections were performed as described previously (32). Seven mice per group in each experiment were used. Animals were euthanized 28 days after injection. The liver of each mouse was dissected, fixed in 4% formaldehyde, and embedded in paraffin. Tumor formation and phenotype were determined by histologic analysis of H&E-stained sections. The tumor volume was calculated using the formula V = 1/2a 2 b (V, volume a, shortest diameter b, longest diameter). The animals were treated in accordance with relevant institutional and national guidelines and regulations. The use of the animals was approved by the Institutional Review Board at MD Anderson Cancer Center. Animals arriving in our facility were randomly put into cages with five mice each. No statistical method was used to predetermine sample size. The investigators were not blinded to allocation during experiments and outcome assessment.

Statistics and reproducibility

The mean values obtained in the control and experimental groups were analyzed for significant differences. Pairwise comparisons were performed using a two-tailed Student t test. P values less than 0.05 were considered significant.


DISCUSSION

Although the manner of programmed cell death of the larval midgut has yet to be clearly established, it is known that mitochondria play a key role in type I (apoptosis) and type II (autophagy) programmed cell death (9, 34). Even as it is dying, the larval midgut must maintain a degree of structural and functional integrity until the pupal epithelium is formed. For this to occur, ATP production must be maintained, and, in this obligatorily aerobic tissue, this means maintaining mitochondrial function. Nevertheless, aerobic metabolism progressively degrades as metamorphosis proceeds. One day before wandering, the midgut's respiration rate falls, and isolated midgut mitochondria have a diminished capacity to oxidize succinate, although oxidation of palmitoyl carnitine is unimpaired (16). Coincident with wandering is an 80% decline in epithelial ion transport and further diminution of mitochondrial substrate oxidation (16). These findings have been confirmed in the present study, because rates of succinate oxidation are lower in mitochondria isolated from day 4 and day 5 (wandering) larvae than from day 2 larvae. In state 4, decreased respiration could be due to decreased activity in the substrate oxidation system and/or a decreased proton conductance of the inner mitochondrial membrane. Elasticity analysis revealed that the depressed state 4 rate seen in day 4 and day 5 mitochondria was not due to decreased proton permeability of the membrane but, rather, decreased activity of the substrate oxidation system. Although elasticity analysis does not identify the site(s) within the substrate oxidation subsystem that is(are) responsible for the decreased activity of the system, changes in the citric acid cycle may be important, because previous studies demonstrated that midgut mitochondrial citrate synthase activities fall between day 2 and day 5 (14).

The results of the present study show that the electron transport chain may be another site of modulation during metamorphosis. Mitochondria isolated from the midguts of postcommitment larvae have lower levels of cytochrome c, but, interestingly, it appears that the total concentration of cytochrome c is the same in the midguts of day 4 and day 2 larvae. These observations are consistent with the release of cytochrome c into the cytoplasm on or before day 4. The appearance of cytochrome c in the cytoplasm is an early event in many (28), but not all (27), cells undergoing programmed cell death. Loss of cytochrome c can lead to a decreased mitochondrial respiration rate, and, in some cases, reintroduction of cytochrome c can restore the respiration rate in mitochondria that have lost this protein (1, 20, 39). This restoration, however, appears to be effective only in the very early stages of cell death, and, as cell death proceeds, irreversible mitochondrial dysfunction is apparent (39, 45). Although it is not known whether midgut mitochondria can be rescued with the addition of exogenous cytochrome c, such a rescue might be unlikely in day 5 mitochondria, where there is a loss of cytochrome aa3. Additional studies are needed to determine whether this loss of cytochrome c oxidase, which is normally found at excess capacity in mitochondria (25), is large enough to impair mitochondrial respiration when exogenous cytochrome c is provided.

Although the activity of the substrate oxidation system declined after commitment to pupation, this subsystem conferred ∼90% of the control over state 3 respiration at all stages of development. This high level of control over respiration differs from that observed in mammalian mitochondria (29, 46) but is similar to that of plant mitochondria (33). The relatively low degree of control over state 3 respiration that is exerted by the phosphorylation system may account for the observation that the oxygen consumption of the intact midgut is apparently insensitive to the phosphorylation status of the cell. That is, inhibition of active ion transport, which should alter the ADP availability and the activity of the phosphorylation system, has no effect on mitochondrial respiration in intact midgut cells (36). In contrast, modulating the substrate oxidation system by presenting the midgut tissue with different metabolic substrates does elevate the tissue's respiration rate (36).

The proton conductance of the inner mitochondrial membrane increases with membrane potential therefore, comparisons of proton leak rates or proton conductances among treatment groups or species must be done at the same membrane potential. In addition, such comparisons must be done at the same temperature. The present study is the first to report proton conductances in insect mitochondria, but the value at 160 mV (∼0.3 nmol H + ·min −1 ·mg protein −1 ·mV −1 ) is similar to that of frog muscle mitochondria (0.26 nmol H + ·min −1 ·mg protein −1 ·mV −1 ) measured at 25°C and 158.9 mV (49). Although the maximal state 4 membrane potential was lower in day 5 than in day 2 mitochondria, the proton conductance was higher. Proton permeability is affected by membrane area (43), lipid composition (32), and the presence and types of uncoupling proteins (50), but it is not known whether any of these factors change during tobacco hornworm development.

The proton leak confers about half the control over state 4 respiration, but this declines to near zero in state 3. The proton leak also confers little control over the phosphorylation system. In contrast, the phosphorylation system confers a substantial negative control over the proton leak. These patterns are similar to those observed in mammalian (22, 29, 30, 46) and plant (33) mitochondria.

In summary, the substrate oxidation system confers substantial control over oxidative phosphorylation in midgut mitochondria, and it is activity of this subsystem that declines during the early stages of metamorphosis. The large decline in the kinetics of the substrate oxidation system is reminiscent of those changes seen during metabolic depression in other organisms. Studies on hibernating ground squirrels (2), hibernating frogs (49), and estivating snails (3) show a decrease in the activity of the substrate oxidation system with no change in the proton leak. Whether the specific sites within the substrate oxidation system that are targeted during programmed cell death are the same as those targeted during metabolic depression remains to be determined.


Watch the video: Substrate level vs. Oxidative Phosphorylation (June 2022).


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