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About the mechanism of coupled reaction/metabolism with ATP

About the mechanism of coupled reaction/metabolism with ATP


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I am not in the field of biochemistry so this may be a rookie question or misconception.

I heard occasionally about the energy "released" from ATP hydrolysis fueling (endergonic) biological reactions. I am concerned about how does the energy "released" actually "fuel" another reaction mechanistically. In chemistry and physics, you can say that the fact that energy is released is a sign that the process (ATP hydrolysis) is energetically favorable (more stable bonds are formed as some weaker ones are broken). Subsequently, when the released energy is heat for instance, this heat can be used to do (or "fuel") mechanical work (like in a car engine). This is intuitive to me but the same cannot be said with ATP hydrolysis. How does the release of energy from ATP-hydrolysis actually "fuel" other reactions? or is it a misuse?

Thank you so much.


Ethanol metabolism

Ethanol, an alcohol found in nature and in alcoholic drinks, is metabolized through a complex catabolic metabolic pathway. In humans, several enzymes are involved in processing ethanol first into acetaldehyde and further into acetic acid and acetyl-CoA. Once acetyl-CoA is formed, it becomes a substrate for the citric acid cycle ultimately producing cellular energy and releasing water and carbon dioxide. Due to differences in enzyme presence and availability, human adults and fetuses process ethanol through different pathways. Gene variation in these enzymes can lead to variation in catalytic efficiency between individuals. The liver is the major organ that metabolizes ethanol due to its high concentration of these enzymes.


Contents

Complex I is the first enzyme of the mitochondrial electron transport chain. There are three energy-transducing enzymes in the electron transport chain - NADH:ubiquinone oxidoreductase (complex I), Coenzyme Q – cytochrome c reductase (complex III), and cytochrome c oxidase (complex IV). [1] Complex I is the largest and most complicated enzyme of the electron transport chain. [2]

The reaction catalyzed by complex I is:

NADH + H + + CoQ + 4H + in→ NAD + + CoQH2 + 4H + out

In this process, the complex translocates four protons across the inner membrane per molecule of oxidized NADH, [3] [4] [5] helping to build the electrochemical potential difference used to produce ATP. Escherichia coli complex I (NADH dehydrogenase) is capable of proton translocation in the same direction to the established Δψ, showing that in the tested conditions, the coupling ion is H + . [6] Na + transport in the opposite direction was observed, and although Na + was not necessary for the catalytic or proton transport activities, its presence increased the latter. H + was translocated by the Paracoccus denitrificans complex I, but in this case, H + transport was not influenced by Na + , and Na + transport was not observed. Possibly, the E. coli complex I has two energy coupling sites (one Na + independent and the other Na + dependent), as observed for the Rhodothermus marinus complex I, whereas the coupling mechanism of the P. denitrificans enzyme is completely Na + independent. It is also possible that another transporter catalyzes the uptake of Na + . Complex I energy transduction by proton pumping may not be exclusive to the R. marinus enzyme. The Na + /H + antiport activity seems not to be a general property of complex I. [6] However, the existence of Na + -translocating activity of the complex I is still in question.

The reaction can be reversed – referred to as aerobic succinate-supported NAD + reduction by ubiquinol – in the presence of a high membrane potential, but the exact catalytic mechanism remains unknown. Driving force of this reaction is a potential across the membrane which can be maintained either by ATP-hydrolysis or by complexes III and IV during succinate oxidation. [7]

Complex I may have a role in triggering apoptosis. [8] In fact, there has been shown to be a correlation between mitochondrial activities and programmed cell death (PCD) during somatic embryo development. [9]

Complex I is not homologous to Na + -translocating NADH Dehydrogenase (NDH) Family (TC# 3.D.1), a member of the Na + transporting Mrp superfamily.

As a result of a two NADH molecule being oxidized to NAD+, three molecules of ATP can be produced by Complex IV downstream in the respiratory chain.

Overall mechanism Edit

All redox reactions take place in the hydrophilic domain of complex I. NADH initially binds to complex I, and transfers two electrons to the flavin mononucleotide (FMN) prosthetic group of the enzyme, creating FMNH2. The electron acceptor – the isoalloxazine ring – of FMN is identical to that of FAD. The electrons are then transferred through the FMN via a series of iron-sulfur (Fe-S) clusters, [10] and finally to coenzyme Q10 (ubiquinone). This electron flow changes the redox state of the protein, inducing conformational changes of the protein which alters the pK values of ionizable side chain, and causes four hydrogen ions to be pumped out of the mitochondrial matrix. [11] Ubiquinone (CoQ) accepts two electrons to be reduced to ubiquinol (CoQH2). [1]

Electron transfer mechanism Edit

The proposed pathway for electron transport prior to ubiquinone reduction is as follows: NADH – FMN – N3 – N1b – N4 – N5 – N6a – N6b – N2 – Q, where Nx is a labelling convention for iron sulfur clusters. [10] The high reduction potential of the N2 cluster and the relative proximity of the other clusters in the chain enable efficient electron transfer over long distance in the protein (with transfer rates from NADH to N2 iron-sulfur cluster of about 100 μs). [12] [13]

The equilibrium dynamics of Complex I are primarily driven by the quinone redox cycle. In conditions of high proton motive force (and accordingly, a ubiquinol-concentrated pool), the enzyme runs in the reverse direction. Ubiquinol is oxidized to ubiquinone, and the resulting released protons reduce the proton motive force. [14]

Proton translocation mechanism Edit

The coupling of proton translocation and electron transport in Complex I is currently proposed as being indirect (long range conformational changes) as opposed to direct (redox intermediates in the hydrogen pumps as in heme groups of Complexes III and IV). [10] The architecture of the hydrophobic region of complex I shows multiple proton transporters that are mechanically interlinked. The three central components believed to contribute to this long-range conformational change event are the pH-coupled N2 iron-sulfur cluster, the quinone reduction, and the transmembrane helix subunits of the membrane arm. Transduction of conformational changes to drive the transmembrane transporters linked by a 'connecting rod' during the reduction of ubiquinone can account for two or three of the four protons pumped per NADH oxidized. The remaining proton must be pumped by direct coupling at the ubiquinone-binding site. It is proposed that direct and indirect coupling mechanisms account for the pumping of the four protons. [15]

The N2 cluster's proximity to a nearby cysteine residue results in a conformational change upon reduction in the nearby helices, leading to small but important changes in the overall protein conformation. [16] Further electron paramagnetic resonance studies of the electron transfer have demonstrated that most of the energy that is released during the subsequent CoQ reduction is on the final ubiquinol formation step from semiquinone, providing evidence for the "single stroke" H + translocation mechanism (i.e. all four protons move across the membrane at the same time). [14] [17] Alternative theories suggest a "two stroke mechanism" where each reduction step (semiquinone and ubiquinol) results in a stroke of two protons entering the intermembrane space. [18] [19]

The resulting ubiquinol localized to the membrane domain interacts with negatively charged residues in the membrane arm, stabilizing conformational changes. [10] An antiporter mechanism (Na + /H + swap) has been proposed using evidence of conserved Asp residues in the membrane arm. [20] The presence of Lys, Glu, and His residues enable for proton gating (a protonation followed by deprotonation event across the membrane) driven by the pKa of the residues. [10]

NADH:ubiquinone oxidoreductase is the largest of the respiratory complexes. In mammals, the enzyme contains 44 separate water-soluble peripheral membrane proteins, which are anchored to the integral membrane constituents. Of particular functional importance are the flavin prosthetic group (FMN) and eight iron-sulfur clusters (FeS). Of the 44 subunits, seven are encoded by the mitochondrial genome. [21] [22] [23]

The structure is an "L" shape with a long membrane domain (with around 60 trans-membrane helices) and a hydrophilic (or peripheral) domain, which includes all the known redox centres and the NADH binding site. [24] All thirteen of the E. coli proteins, which comprise NADH dehydrogenase I, are encoded within the nuo operon, and are homologous to mitochondrial complex I subunits. The antiporter-like subunits NuoL/M/N each contains 14 conserved transmembrane (TM) helices. Two of them are discontinuous, but subunit NuoL contains a 110 Å long amphipathic α-helix, spanning the entire length of the domain. The subunit, NuoL, is related to Na + / H + antiporters of TC# 2.A.63.1.1 (PhaA and PhaD).

Three of the conserved, membrane-bound subunits in NADH dehydrogenase are related to each other, and to Mrp sodium-proton antiporters. Structural analysis of two prokaryotic complexes I revealed that the three subunits each contain fourteen transmembrane helices that overlay in structural alignments: the translocation of three protons may be coordinated by a lateral helix connecting them. [25]

Complex I contains a ubiquinone binding pocket at the interface of the 49-kDa and PSST subunits. Close to iron-sulfur cluster N2, the proposed immediate electron donor for ubiquinone, a highly conserved tyrosine constitutes a critical element of the quinone reduction site. A possible quinone exchange path leads from cluster N2 to the N-terminal beta-sheet of the 49-kDa subunit. [26] All 45 subunits of the bovine NDHI have been sequenced. [27] [28] Each complex contains noncovalently bound FMN, coenzyme Q and several iron-sulfur centers. The bacterial NDHs have 8-9 iron-sulfur centers.

A recent study used electron paramagnetic resonance (EPR) spectra and double electron-electron resonance (DEER) to determine the path of electron transfer through the iron-sulfur complexes, which are located in the hydrophilic domain. Seven of these clusters form a chain from the flavin to the quinone binding sites the eighth cluster is located on the other side of the flavin, and its function is unknown. The EPR and DEER results suggest an alternating or “roller-coaster” potential energy profile for the electron transfer between the active sites and along the iron-sulfur clusters, which can optimize the rate of electron travel and allow efficient energy conversion in complex I. [29]

Conserved subunits of Complex I [30]
# Human/Bovine subunit Human protein Protein description (UniProt) Pfam family with Human protein
Core Subunits a
1 NDUFS7 / PSST / NUKM NDUS7_HUMAN NADH dehydrogenase [ubiquinone] iron-sulfur protein 7, mitochondrial EC 1.6.5.3 EC 1.6.99.3 Pfam PF01058
2 NDUFS8 / TYKY / NUIM NDUS8_HUMAN NADH dehydrogenase [ubiquinone] iron-sulfur protein 8, mitochondrial EC 1.6.5.3 EC 1.6.99.3 Pfam PF12838
3 NDUFV2 / 24kD / NUHM c NDUV2_HUMAN NADH dehydrogenase [ubiquinone] flavoprotein 2, mitochondrial EC 1.6.5.3 EC 1.6.99.3 Pfam PF01257
4 NDUFS3 / 30kD / NUGM NDUS3_HUMAN NADH dehydrogenase [ubiquinone] iron-sulfur protein 3, mitochondrial EC 1.6.5.3 EC 1.6.99.3 Pfam PF00329
5 NDUFS2 / 49kD / NUCM NDUS2_HUMAN NADH dehydrogenase [ubiquinone] iron-sulfur protein 2, mitochondrial EC 1.6.5.3 EC 1.6.99.3 Pfam PF00346
6 NDUFV1 / 51kD / NUBM NDUV1_HUMAN NADH dehydrogenase [ubiquinone] flavoprotein 1, mitochondrial EC 1.6.5.3 EC 1.6.99.3 Pfam PF01512
7 NDUFS1 / 75kD / NUAM NDUS1_HUMAN NADH-ubiquinone oxidoreductase 75 kDa subunit, mitochondrial EC 1.6.5.3 EC 1.6.99.3 Pfam PF00384
8 ND1 / NU1M NU1M_HUMAN NADH-ubiquinone oxidoreductase chain 1 EC 1.6.5.3 Pfam PF00146
9 ND2 / NU2M NU2M_HUMAN NADH-ubiquinone oxidoreductase chain 2 EC 1.6.5.3 Pfam PF00361, Pfam PF06444
10 ND3 / NU3M NU3M_HUMAN NADH-ubiquinone oxidoreductase chain 3 EC 1.6.5.3 Pfam PF00507
11 ND4 / NU4M NU4M_HUMAN NADH-ubiquinone oxidoreductase chain 4 EC 1.6.5.3 Pfam PF01059, Pfam PF00361
12 ND4L / NULM NU4LM_HUMAN NADH-ubiquinone oxidoreductase chain 4L EC 1.6.5.3 Pfam PF00420
13 ND5 / NU5M NU5M_HUMAN NADH-ubiquinone oxidoreductase chain 5 EC 1.6.5.3 Pfam PF00361, Pfam PF06455, Pfam PF00662
14 ND6 / NU6M NU6M_HUMAN NADH-ubiquinone oxidoreductase chain 6 EC 1.6.5.3 Pfam PF00499
Core accessory subunits b
15 NDUFS6 / 13A NDUS6_HUMAN NADH dehydrogenase [ubiquinone] iron-sulfur protein 6, mitochondrial Pfam PF10276
16 NDUFA12 / B17.2 NDUAC_HUMAN NADH dehydrogenase [ubiquinone] 1 alpha subcomplex subunit 12 Pfam PF05071
17 NDUFS4 / AQDQ NDUS4_HUMAN NADH dehydrogenase [ubiquinone] iron-sulfur protein 4, mitochondrial Pfam PF04800
18 NDUFA9 / 39kDa NDUA9_HUMAN NADH dehydrogenase [ubiquinone] 1 alpha subcomplex subunit 9, mitochondrial Pfam PF01370
19 NDUFAB1 / ACPM ACPM_HUMAN Acyl carrier protein, mitochondrial Pfam PF00550
20 NDUFA2 / B8 NDUA2_HUMAN NADH dehydrogenase [ubiquinone] 1 alpha subcomplex subunit 2 Pfam PF05047
21 NDUFA1 / MFWE NDUA1_HUMAN NADH dehydrogenase [ubiquinone] 1 alpha subcomplex subunit 1 Pfam PF15879
22 NDUFB3 / B12 NDUB3_HUMAN NADH dehydrogenase [ubiquinone] 1 beta subcomplex subunit 3 Pfam PF08122
23 NDUFA5 / AB13 NDUA5_HUMAN NADH dehydrogenase [ubiquinone] 1 alpha subcomplex subunit 5 Pfam PF04716
24 NDUFA6 / B14 NDUA6_HUMAN NADH dehydrogenase [ubiquinone] 1 alpha subcomplex subunit 6 Pfam PF05347
25 NDUFA11 / B14.7 NDUAB_HUMAN NADH dehydrogenase [ubiquinone] 1 alpha subcomplex subunit 11 Pfam PF02466
26 NDUFB11 / ESSS NDUBB_HUMAN NADH dehydrogenase [ubiquinone] 1 beta subcomplex subunit 11, mitochondrial Pfam PF10183
27 NDUFS5 / PFFD NDUS5_HUMAN NADH dehydrogenase [ubiquinone] iron-sulfur protein 5 Pfam PF10200
28 NDUFB4 / B15 NDUB4_HUMAN NADH dehydrogenase [ubiquinone] 1 beta subcomplex subunit 4 Pfam PF07225
29 NDUFA13 /A13 NDUAD_HUMAN NADH dehydrogenase [ubiquinone] 1 alpha subcomplex subunit 13 Pfam PF06212
30 NDUFB7 / B18 NDUB7_HUMAN NADH dehydrogenase [ubiquinone] 1 beta subcomplex subunit 7 Pfam PF05676
31 NDUFA8 / PGIV NDUA8_HUMAN NADH dehydrogenase [ubiquinone] 1 alpha subcomplex subunit 8 Pfam PF06747
32 NDUFB9 / B22 NDUB9_HUMAN NADH dehydrogenase [ubiquinone] 1 beta subcomplex subunit 9 Pfam PF05347
33 NDUFB10 / PDSW NDUBA_HUMAN NADH dehydrogenase [ubiquinone] 1 beta subcomplex subunit 10 Pfam PF10249
34 NDUFB8 / ASHI NDUB8_HUMAN NADH dehydrogenase [ubiquinone] 1 beta subcomplex subunit 8, mitochondrial Pfam PF05821
35 NDUFC2 / B14.5B NDUC2_HUMAN NADH dehydrogenase [ubiquinone] 1 subunit C2 Pfam PF06374
36 NDUFB2 / AGGG NDUB2_HUMAN NADH dehydrogenase [ubiquinone] 1 beta subcomplex subunit 2, mitochondrial Pfam PF14813
37 NDUFA7 / B14.5A NDUA7_HUMAN NADH dehydrogenase [ubiquinone] 1 alpha subcomplex subunit 7 Pfam PF07347
38 NDUFA3 / B9 NDUA3_HUMAN NADH dehydrogenase [ubiquinone] 1 alpha subcomplex subunit 3 Pfam PF14987
39 NDUFA4 / MLRQ c NDUA4_HUMAN NADH dehydrogenase [ubiquinone] 1 alpha subcomplex subunit 4 Pfam PF06522
40 NDUFB5 / SGDH NDUB5_HUMAN NADH dehydrogenase [ubiquinone] 1 beta subcomplex subunit 5, mitochondrial Pfam PF09781
41 NDUFB1 / MNLL NDUB1_HUMAN NADH dehydrogenase [ubiquinone] 1 beta subcomplex subunit 1 Pfam PF08040
42 NDUFC1 / KFYI NDUC1_HUMAN NADH dehydrogenase [ubiquinone] 1 subunit C1, mitochondrial Pfam PF15088
43 NDUFA10 / 42kD NDUAA_HUMAN NADH dehydrogenase [ubiquinone] 1 alpha subcomplex subunit 10, mitochondrial Pfam PF01712
44 NDUFA4L2 NUA4L_HUMAN NADH dehydrogenase [ubiquinone] 1 alpha subcomplex subunit 4-like 2 Pfam PF15880
45 NDUFV3 NDUV3_HUMAN NADH dehydrogenase [ubiquinone] flavoprotein 3, 10kDa -
46 NDUFB6 NDUB6_HUMAN NADH dehydrogenase [ubiquinone] 1 beta subcomplex subunit 6 Pfam PF09782
Assembly factor proteins [31]
47 NDUFAF1 c CIA30_HUMAN NADH dehydrogenase [ubiquinone] 1 alpha subcomplex, assembly factor 1 Pfam PF08547
48 NDUFAF2 MIMIT_HUMAN NADH dehydrogenase [ubiquinone] 1 alpha subcomplex, assembly factor 2 Pfam PF05071
49 NDUFAF3 NDUF3_HUMAN NADH dehydrogenase [ubiquinone] 1 alpha subcomplex assembly factor 3 Pfam PF05071
50 NDUFAF4 NDUF4_HUMAN NADH dehydrogenase [ubiquinone] 1 alpha subcomplex, assembly factor 4 Pfam PF06784

  • a Found in all species except fungi
  • b May or may not be present in any species
  • c Found in fungal species such as Schizosaccharomyces pombe

Bullatacin (an acetogenin found in Asimina triloba fruit) is the most potent known inhibitor of NADH dehydrogenase (ubiquinone) (IC50=1.2 nM, stronger than rotenone). [34] The best-known inhibitor of complex I is rotenone (commonly used as an organic pesticide). Rotenone and rotenoids are isoflavonoids occurring in several genera of tropical plants such as Antonia (Loganiaceae), Derris and Lonchocarpus (Faboideae, Fabaceae). There have been reports of the indigenous people of French Guiana using rotenone-containing plants to fish - due to its ichthyotoxic effect - as early as the 17th century. [35] Rotenone binds to the ubiquinone binding site of complex I as well as piericidin A, another potent inhibitor with a close structural homologue to ubiquinone.

Acetogenins from Annonaceae are even more potent inhibitors of complex I. They cross-link to the ND2 subunit, which suggests that ND2 is essential for quinone-binding. [36] Rolliniastatin-2, an acetogenin, is the first complex I inhibitor found that does not share the same binding site as rotenone. [37]

Despite more than 50 years of study of complex I, no inhibitors blocking the electron flow inside the enzyme have been found. Hydrophobic inhibitors like rotenone or piericidin most likely disrupt the electron transfer between the terminal FeS cluster N2 and ubiquinone. It has been shown that long-term systemic inhibition of complex I by rotenone can induce selective degeneration of dopaminergic neurons. [38]

Complex I is also blocked by adenosine diphosphate ribose – a reversible competitive inhibitor of NADH oxidation – by binding to the enzyme at the nucleotide binding site. [39] Both hydrophilic NADH and hydrophobic ubiquinone analogs act at the beginning and the end of the internal electron-transport pathway, respectively.

The antidiabetic drug Metformin has been shown to induce a mild and transient inhibition of the mitochondrial respiratory chain complex I, and this inhibition appears to play a key role in its mechanism of action. [40]

Inhibition of complex I has been implicated in hepatotoxicity associated with a variety of drugs, for instance flutamide and nefazodone. [41]

The catalytic properties of eukaryotic complex I are not simple. Two catalytically and structurally distinct forms exist in any given preparation of the enzyme: one is the fully competent, so-called “active” A-form and the other is the catalytically silent, dormant, “deactive”, D-form. After exposure of idle enzyme to elevated, but physiological temperatures (>30 °C) in the absence of substrate, the enzyme converts to the D-form. This form is catalytically incompetent but can be activated by the slow reaction (k

4 min −1 ) of NADH oxidation with subsequent ubiquinone reduction. After one or several turnovers the enzyme becomes active and can catalyse physiological NADH:ubiquinone reaction at a much higher rate (k

10 4 min −1 ). In the presence of divalent cations (Mg 2+ , Ca 2+ ), or at alkaline pH the activation takes much longer.

The high activation energy (270 kJ/mol) of the deactivation process indicates the occurrence of major conformational changes in the organisation of the complex I. However, until now, the only conformational difference observed between these two forms is the number of cysteine residues exposed at the surface of the enzyme. Treatment of the D-form of complex I with the sulfhydryl reagents N-Ethylmaleimide or DTNB irreversibly blocks critical cysteine residue(s), abolishing the ability of the enzyme to respond to activation, thus inactivating it irreversibly. The A-form of complex I is insensitive to sulfhydryl reagents. [42] [43]

It was found that these conformational changes may have a very important physiological significance. The deactive, but not the active form of complex I was susceptible to inhibition by nitrosothiols and peroxynitrite. [44] It is likely that transition from the active to the inactive form of complex I takes place during pathological conditions when the turnover of the enzyme is limited at physiological temperatures, such as during hypoxia, ischemia [45] [46] or when the tissue nitric oxide:oxygen ratio increases (i.e. metabolic hypoxia). [47]

Recent investigations suggest that complex I is a potent source of reactive oxygen species. [48] Complex I can produce superoxide (as well as hydrogen peroxide), through at least two different pathways. During forward electron transfer, only very small amounts of superoxide are produced (probably less than 0.1% of the overall electron flow). [48] [49] [50]

During reverse electron transfer, complex I might be the most important site of superoxide production within mitochondria, with around 3-4% of electrons being diverted to superoxide formation. [51] Reverse electron transfer, the process by which electrons from the reduced ubiquinol pool (supplied by succinate dehydrogenase, glycerol-3-phosphate dehydrogenase, electron-transferring flavoprotein or dihydroorotate dehydrogenase in mammalian mitochondria) pass through complex I to reduce NAD + to NADH, driven by the inner mitochondrial membrane potential electric potential. Although it is not precisely known under what pathological conditions reverse-electron transfer would occur in vivo, in vitro experiments indicate that this process can be a very potent source of superoxide when succinate concentrations are high and oxaloacetate or malate concentrations are low. [52] This can take place during tissue ischaemia, when oxygen delivery is blocked. [53]

Superoxide is a reactive oxygen species that contributes to cellular oxidative stress and is linked to neuromuscular diseases and aging. [54] NADH dehdyrogenase produces superoxide by transferring one electron from FMNH2 (or semireduced flavin) to oxygen (O2). The radical flavin leftover is unstable, and transfers the remaining electron to the iron-sulfur centers. It is the ratio of NADH to NAD + that determines the rate of superoxide formation. [55] [56]

Mutations in the subunits of complex I can cause mitochondrial diseases, including Leigh syndrome. Point mutations in various complex I subunits derived from mitochondrial DNA (mtDNA) can also result in Leber's Hereditary Optic Neuropathy. There is some evidence that complex I defects may play a role in the etiology of Parkinson's disease, perhaps because of reactive oxygen species (complex I can, like complex III, leak electrons to oxygen, forming highly toxic superoxide).

Although the exact etiology of Parkinson’s disease is unclear, it is likely that mitochondrial dysfunction, along with proteasome inhibition and environmental toxins, may play a large role. In fact, the inhibition of complex I has been shown to cause the production of peroxides and a decrease in proteasome activity, which may lead to Parkinson’s disease. [57] Additionally, Esteves et al. (2010) found that cell lines with Parkinson’s disease show increased proton leakage in complex I, which causes decreased maximum respiratory capacity. [58]

Brain ischemia/reperfusion injury is mediated via complex I impairment. [59] Recently it was found that oxygen deprivation leads to conditions in which mitochondrial complex I lose its natural cofactor, flavin mononucleotide (FMN) and become inactive. [60] [61] When oxygen is present the enzyme catalyzes a physiological reaction of NADH oxidation by ubiquinone, supplying electrons downstream of the respiratory chain (complexes III and IV). Ischemia leads to dramatic increase of succinate level. In the presence of succinate mitochondria catalyze reverse electron transfer so that fraction of electrons from succinate is directed upstream to FMN of complex I. Reverse electron transfer results in a reduction of complex I FMN, [62] increased generation of ROS, followed by a loss of the reduced cofactor (FMNH2) and impairement of mitochondria energy production. The FMN loss by complex I and I/R injury can be alleviated by the administration of FMN precursor, riboflavin. [61]

Recent studies have examined other roles of complex I activity in the brain. Andreazza et al. (2010) found that the level of complex I activity was significantly decreased in patients with bipolar disorder, but not in patients with depression or schizophrenia. They found that patients with bipolar disorder showed increased protein oxidation and nitration in their prefrontal cortex. These results suggest that future studies should target complex I for potential therapeutic studies for bipolar disorder. [63] Similarly, Moran et al. (2010) found that patients with severe complex I deficiency showed decreased oxygen consumption rates and slower growth rates. However, they found that mutations in different genes in complex I lead to different phenotypes, thereby explaining the variations of pathophysiological manifestations of complex I deficiency. [64]

Exposure to pesticides can also inhibit complex I and cause disease symptoms. For example, chronic exposure to low levels of dichlorvos, an organophosphate used as a pesticide, has been shown to cause liver dysfunction. This occurs because dichlorvos alters complex I and II activity levels, which leads to decreased mitochondrial electron transfer activities and decreased ATP synthesis. [65]

The following is a list of humans genes that encode components of complex I:

  • NADH dehydrogenase (ubiquinone) 1 alpha subcomplex
      – NADH dehydrogenase (ubiquinone) 1 alpha subcomplex, 1, 7.5kDa – NADH dehydrogenase (ubiquinone) 1 alpha subcomplex, 2, 8kDa
  • NDUFA3 – NADH dehydrogenase (ubiquinone) 1 alpha subcomplex, 3, 9kDa
  • NDUFA4 – NADH dehydrogenase (ubiquinone) 1 alpha subcomplex, 4, 9kDa
  • NDUFA4L – NADH dehydrogenase (ubiquinone) 1 alpha subcomplex, 4-like
  • NDUFA4L2 – NADH dehydrogenase (ubiquinone) 1 alpha subcomplex, 4-like 2 – NADH dehydrogenase (ubiquinone) 1 alpha subcomplex, 5, 13kDa – NADH dehydrogenase (ubiquinone) 1 alpha subcomplex, 6, 14kDa
  • NDUFA7 – NADH dehydrogenase (ubiquinone) 1 alpha subcomplex, 7, 14.5kDa – NADH dehydrogenase (ubiquinone) 1 alpha subcomplex, 8, 19kDa – NADH dehydrogenase (ubiquinone) 1 alpha subcomplex, 9, 39kDa – NADH dehydrogenase (ubiquinone) 1 alpha subcomplex, 10, 42kDa
  • NDUFA11 – NADH dehydrogenase (ubiquinone) 1 alpha subcomplex, 11, 14.7kDa – NADH dehydrogenase (ubiquinone) 1 alpha subcomplex, 12 – NADH dehydrogenase (ubiquinone) 1 alpha subcomplex, 13
  • NDUFAB1 – NADH dehydrogenase (ubiquinone) 1, alpha/beta subcomplex, 1, 8kDa – NADH dehydrogenase (ubiquinone) 1 alpha subcomplex, assembly factor 1
  • NDUFAF2 – NADH dehydrogenase (ubiquinone) 1 alpha subcomplex, assembly factor 2
  • NDUFAF3 – NADH dehydrogenase (ubiquinone) 1 alpha subcomplex, assembly factor 3
  • NDUFAF4 – NADH dehydrogenase (ubiquinone) 1 alpha subcomplex, assembly factor 4
    • – NADH dehydrogenase (ubiquinone) 1 beta subcomplex, 1, 7kDa – NADH dehydrogenase (ubiquinone) 1 beta subcomplex, 2, 8kDa
    • NDUFB3 – NADH dehydrogenase (ubiquinone) 1 beta subcomplex, 3, 12kDa
    • NDUFB4 – NADH dehydrogenase (ubiquinone) 1 beta subcomplex, 4, 15kDa
    • NDUFB5 – NADH dehydrogenase (ubiquinone) 1 beta subcomplex, 5, 16kDa – NADH dehydrogenase (ubiquinone) 1 beta subcomplex, 6, 17kDa – NADH dehydrogenase (ubiquinone) 1 beta subcomplex, 7, 18kDa – NADH dehydrogenase (ubiquinone) 1 beta subcomplex, 8, 19kDa – NADH dehydrogenase (ubiquinone) 1 beta subcomplex, 9, 22kDa – NADH dehydrogenase (ubiquinone) 1 beta subcomplex, 10, 22kDa – NADH dehydrogenase (ubiquinone) 1 beta subcomplex, 11, 17.3kDa
      – NADH dehydrogenase (ubiquinone) 1, subcomplex unknown, 1, 6kDa – NADH dehydrogenase (ubiquinone) 1, subcomplex unknown, 2, 14.5kDa
      – NADH dehydrogenase (ubiquinone) Fe-S protein 1, 75kDa (NADH-coenzyme Q reductase) – NADH dehydrogenase (ubiquinone) Fe-S protein 2, 49kDa (NADH-coenzyme Q reductase) – NADH dehydrogenase (ubiquinone) Fe-S protein 3, 30kDa (NADH-coenzyme Q reductase) – NADH dehydrogenase (ubiquinone) Fe-S protein 4, 18kDa (NADH-coenzyme Q reductase) – NADH dehydrogenase (ubiquinone) Fe-S protein 5, 15kDa (NADH-coenzyme Q reductase) – NADH dehydrogenase (ubiquinone) Fe-S protein 6, 13kDa (NADH-coenzyme Q reductase) – NADH dehydrogenase (ubiquinone) Fe-S protein 7, 20kDa (NADH-coenzyme Q reductase) – NADH dehydrogenase (ubiquinone) Fe-S protein 8, 23kDa (NADH-coenzyme Q reductase)
      – NADH dehydrogenase (ubiquinone) flavoprotein 1, 51kDa – NADH dehydrogenase (ubiquinone) flavoprotein 2, 24kDa – NADH dehydrogenase (ubiquinone) flavoprotein 3, 10kDa
      - mitochondrially encoded NADH dehydrogenase subunit 1 - mitochondrially encoded NADH dehydrogenase subunit 2 - mitochondrially encoded NADH dehydrogenase subunit 3 - mitochondrially encoded NADH dehydrogenase subunit 4 - mitochondrially encoded NADH dehydrogenase subunit 4L - mitochondrially encoded NADH dehydrogenase subunit 5 - mitochondrially encoded NADH dehydrogenase subunit 6
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    5. Wikström M (April 1984). "Two protons are pumped from the mitochondrial matrix per electron transferred between NADH and ubiquinone". FEBS Letters. 169 (2): 300–4. doi: 10.1016/0014-5793(84)80338-5 . PMID6325245.
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    As of this edit, this article uses content from "3.D.1 The H+ or Na+-translocating NADH Dehydrogenase (NDH) Family", which is licensed in a way that permits reuse under the Creative Commons Attribution-ShareAlike 3.0 Unported License, but not under the GFDL. All relevant terms must be followed.


    C1. ATP

    • Contributed by Henry Jakubowski
    • Professor (Chemistry) at College of St. Benedict/St. John's University

    Biological oxidation reactions serve two functions. Oxidation of organic molecules can produce new molecules with different properties. For example, increases in solubility is observed on hydroxylation of aromatic substrates by cytochrome P450. Likewise, amino acids can by oxidized to produce neurotransmitters. Most biological oxidation reactions occur, however, to produce energy to drive thermodynamically unfavored biological processes such as protein and nucleic acid synthesis, or motility. Chemical potential energy is not just released in biological oxidation reactions. Rather, it is transduced into a more useful form of chemical energy in the molecule ATP (adenosine triphosphate). This chapter will discuss the properties that make ATP so useful biologically, and how exergonic biological oxidation reactions are coupled to the synthesis of ATP.

    ATP contains two phosphoanhydride bonds (connecting the 3 phosphates together) and one phosphoester bond (connecting a phosphate to the ribose ring). The pKa's for the reactions

    are about 7.0, so the overall charges of ATP and ADP at physiological pH are -3.5 and -2.5, respectively. Each of the phosphorous atoms are highly electrophilic and can react with nucleophiles like the OH of water or an alcohol. As we discussed earlier, anhydrides are thermodynamically more reactive than esters which are more reactive than amides. The large negative (&DeltaG^o = -7.5, kcal/mol) for th e hydrolysis (a nucleophilic substitution reaction) of one of the phosphoanhydride bonds can be attributed to a relative destabilization of the reactants (ATP and water) and relative stabilization of the products (ADP = Pi). Specifically

    • The reactants can not be stabilized to the same extent as products by resonance due to competing resonance of the bridging anhydride O's.
    • The charge density on the reactants is greater than that of the products
    • Theoretical studies show that the products are more hydrated than the reactants.

    The (&DeltaG^o) for hydrolysis of ATP is dependent on the divalent ion concentration and pH, which affect the the stabilization and the magnitude of the charge states of the reactants and products.

    Jmol: Updated ATP Jmol14 (Java) | JSMol (HTML5) | ADP Jmol14 (Java) | JSMol (HTML5)

    Figure: STRUCTURE AND HYDROLYSIS OF ATP

    Carboxylic acid anhydrides are even more unstable to hydrolysis than ATP (-20 kcal/mol), followed by mixed anhydrides (-12 kcal/mol), and phosphoric acid anhydrides (-7.5 kcal/mol). These molecules are often termed "high energy" molecules, which is somewhat of a misnomer. They are high energy only in relation to the energy of their cleavage products, such that the reaction proceeds with a large negative (&DeltaG^o).

    Figure: HIGH ENERGY MOLECULES

    How can ATP be used to drive thermodynamically unfavored reaction? First consider how the hydrolysis of a carboxylic acid anhydride, which has a ( &Delta G^ o) = -12.5 kcal/mol can drive the synthesis of a carboxylic acid amide, with a ( &Delta G^o) of + 2-3 kcal/mol. The link below shows the net reaction, (anhydride + amine --> amide + carboxylic acid), which can be broken into two reactions: hydrolysis of the anhydride, and the synthesis of the amide.

    Figure: MECHANISM: COUPLED SYNTHESIS OF A CARBOXYLIC AMIDE

    Now consider the reaction of glucose + Pi to form glucose-6-P. In this reaction a phosphoester is formed, so the reaction would proceed with a positive &Delta G o = 3.3. Now if ATP was used to transfer the terminal (gamma) phosphate to glucose to form Glc-6-P, the reaction proceeds with a &Delta G o = -4 kcal/mol. This can be calculated since &DeltaG is a state function and is path independent. Adding the reactions and the &Delta G o 's for

    gives the resultant reaction and ( &Delta G^ o)

    In most biological reactions using ATP, the terminal P of ATP is transferred to a substrate using an enzyme called a kinase. Hence, hexokinase transfers the gamma phosphate from ATP to a hexose sugar. Protein kinase is an enzyme which transfers the gamma phosphate to a protein substrate.

    ATP is also used to drive peptide bond (amide) synthesis during protein synthesis. From an energetic point of view, anhydride cleavage can provide the energy for amide bond formation. Peptide bond synthesis is cells is accompanied by cleavage of both phosphoanhydride bonds in ATP in a complicated set of reactions that is catalyzed by ribosomes in the cells. (This topic is considered in depth in molecular biology courses). The figure below is a grossly simplified mechanism of how peptide bond formation can be coupled to ATP cleavage.

    Figure: MECHANISM: ATP-DEPENDENT PEPTIDE BOND SYNTHESIS

    Phosphorylation reactions using ATP are really nucleophilic substitution reactions which proceed through a pentavalent intermediate. The rest of the ATP molecule is then considered the leaving group, which could be theoretically ADP or AMP as well. If water is the nucleophile, the reaction is also a hydrolysis reaction. These reactions are also called phosphoryl transfer reactions.

    One last note. ATP exists in cells as just one member of a pool of adenine nucleotides which consists of not only ATP, but also ADP and AMP (along with Pi). These constituents are readily interconvertible. We actually break down an amount of ATP each day equal to about our body weight. Likewise we make about the same amount from the turnover products. When energy is needed, carbohydrates and lipids are oxidized and ATP is produced, which can then be immediately used for motility, biosynthesis, etc. It is very important to realize that although ATP is converted to ADP in a thermodynamically spontaneous process, the process is kinetically slow without an enzyme. Hence ATP is stable in solution. However, its biological half-life is not long since it is used very quickly as described above. This recapitulates a theme we have seen before. Many reactions (like oxidation with dioxygen, denaturation of proteins in nonpolar solvent, and now ATP hydrolysis) are thermodynamically favored but kinetically slow. This kinetic slowness is a necessary but of course insufficient condition, for life.


    ATP synthesis in the energy metabolism pathway: a new perspective for manipulating CdSe quantum dots biosynthesized in Saccharomyces cerevisiae

    1 Hubei Key Laboratory of Cell Homeostasis, 2 College of Life Sciences, Wuhan University, 3 Key Laboratory of Analytical Chemistry for Biology and Medicine (Ministry of Education), 4 College of Chemistry and Molecular Science, Wuhan University, Wuhan, People&rsquos Republic of China

    Abstract: Due to a growing trend in their biomedical application, biosynthesized nanomaterials are of great interest to researchers nowadays with their biocompatible, low-energy consumption, economic, and tunable characteristics. It is important to understand the mechanism of biosynthesis in order to achieve more efficient applications. Since there are only rare studies on the influences of cellular energy levels on biosynthesis, the influence of energy is often overlooked. Through determination of the intracellular ATP concentrations during the biosynthesis process, significant changes were observed. In addition, ATP synthesis deficiency caused great decreases in quantum dots (QDs) biosynthesis in the &Deltaatp1, &Deltaatp2, &Deltaatp14, and &Deltaatp17 strains. With inductively coupled plasma-atomic emission spectrometry and atomic absorption spectroscopy analyses, it was found that ATP affected the accumulation of the seleno-precursor and helped with the uptake of Cd and the formation of QDs. We successfully enhanced the fluorescence intensity 1.5 or 2 times through genetic modification to increase ATP or SeAM (the seleno analog of S-adenosylmethionine, the product that would accumulate when ATP is accrued). This work explains the mechanism for the correlation of the cellular energy level and QDs biosynthesis in living cells, demonstrates control of the biosynthesis using this mechanism, and thus provides a new manipulation strategy for the biosynthesis of other nanomaterials to widen their applications.

    Keywords: ATP, biosynthesis, Saccharomyces cerevisiae, QDs, CdSe

    Nanomaterials, 1 due to their unique properties, have been widely used in cell imaging, 2,3 diagnosis, 4,5 biosensing, 6,7 electro-optic device, 8 and so on, especially on medical imaging because of their (surface modified nanomaterials) high biocompatibility to cells. Thus, tremendous interest has arisen in the synthesis of well-dispersed and uniform-sized nanomaterials. 9,10 Nanoparticles biosynthesized by organisms, without involving the toxic solvents, strict conditions, or expensive processes that are needed in chemical methods, have become a popular subject. 11 Various nanoparticles are reported that are biosynthesized by bacteria, 12󈝺 fungi, 15,16 plants, 17,18 mammalian cells, 19 or even earthworms. 20 Considering the simple culture methods, the low cost of the equipment, and the easy way to get the gradients, the biosynthesis of nanoparticles using microorganisms is the most economic and common approach.

    There have been many investigations on the biosynthesis of nanomaterials, mainly concerning their applications. 21 In order to obtain nanoparticles with a defined expectation in the biosynthesis, whether through optimizing the culture conditions or adjusting the concentrations of primary substrates, former studies were devoted to manipulating the biosynthesis by extrinsic factors, and few were on the intrinsic characters of the microorganisms applied in the biosynthesis. Ayano et al 22 found that the optimal culture conditions to biosynthesize CdSe nanoparticles by Pseudomonas aeruginosa were when the cells were cultured at 25°C󈞔°C, 0.05󈝶 g L 𕒵 of NaCl concentration, and neutral pH. According to Gericke and Pinches, 23 the size and shape of gold nanoparticles biosynthesized by fungus could be manipulated by changing crucial growth parameters. They successfully obtained spherical, hexagonal, and triangular-shaped nanoparticles with different sizes through culturing under various pH and temperature conditions. Besides the studies of the adscititious substances and the culture conditions, there have also been some studies focused on the intrinsic metabolisms of the bioreactor used in the biosynthesis. For example, for Sb2O3 nanoparticles biosynthesized by yeast, reported by Jha et al, 24 it was suggested that the membrane-bound and cytosolic oxidoreductases and quinines might be the key influential factors of the process. Phenol oxidases in the Lentinus edodes were proved to be responsible for the biosynthesis of Au nanoparticles. 25 In general, current research has mostly focused on the culture conditions or the substances, while the role of energy has hardly been mentioned.

    In the previous work of our laboratory, we constructed a convenient way to biosynthesize CdSe quantum dots (QDs) with baker’s yeast Saccharomyces cerevisiae 26 and then promoted the QDs production by genetic modification. 27 Glutathione (L-γ-glutamylcysteinylglycine, GSH) was found to be a vital compound in the biosynthesis of CdSe QDs, the intracellular content of which showed synchronized increase with the fluorescence intensities when seleniumized cells were initially incubated with CdCl2. 27 Besides the substances that assisted with the biosynthesis of CdSe QDs, GO (gene ontology) analysis revealed that the proteins that encapsulated the CdSe QDs biosynthesized by yeast mostly functioned in cell energy metabolism (unpublished data) thus, this may indicate that energy has a noticeable role in the process. There may be a need for not only micromolecular polypeptides (such as GSH) 27 but also cell energy to ensure the ambient temperature and strict conditions for the stable biosynthesis of CdSe QDs with good dispersal and uniformity. With its favorable biocompatibility characteristic, the system was successfully applied to construct a cell beacon using Staphylococcus aureus as a bioreactor. 28

    Figuring out the actual role that energy plays in the biosynthesis would be conducive to further understanding of the mechanism. 26,27 The present study used the concentrations of intracellular ATP (the most commonly used direct energy resource) and fluorescence intensities as indicators to investigate the influences of energy on the biosynthesis of CdSe QDs in yeast. Intracellular Se or Cd concentrations were checked during the cell-seleniumized phase or the crystallization phase 26 with inductively coupled plasma-atomic emission spectrometry (ICP-AES) and atomic absorption spectroscopy (AAS) to confirm the influences on the absorption of the substrates and the crystallization of CdSe QDs. Finally, the fluorescence intensities were improved by genetic modification of the ATP metabolism pathway. In consequence, the biosynthesis can be controlled with defined expectation and it can be used as guidance for the biosynthesis of other similar nanoparticles with yeast or other microorganisms. The present work provides a new perspective for future research on the biosynthesis mechanisms of nanomaterials and makes it easier to convert the biosynthesized nanomaterials into applications.

    Saccharomyces cerevisiae BY4742 (MATα his3-Δ1 leu2-Δ0 lys2-Δ0 ura3-Δ0) (wild-type, WT) and the Δatp1, Δatp2, Δatp14, Δatp17, Δpfk1, and Δrgt2 mutant strains were obtained from the European Saccharomyces cerevisiae Archive for Functional Analysis (Bad Homburg, Germany) (all strain information is listed in Table S1). The PGAL1-ADK1 and PGAL1-SAM2 strains were constructed in this study (Table S2). Strains were cultured in 1 g L 𕒵 yeast extract and 2 g L 𕒵 peptone (YP), supplied with 2 g L 𕒵 glucose (YPGlu) or YP supplied with 2 g L 𕒵 galactose (YPGal) media. Unless otherwise indicated, the reagents used in the present work were obtained from the Sinopharm Chemical Reagent Co. (Shanghai, China).

    The biosynthesis of CdSe QDs with yeast was performed according to the previous study, 26 with a slight adjustment, as follows: stationary phase cells were co-incubated with Na2SeO3 at a final concentration of 2.5 mM for 24 hours to get seleniumized (hereafter referred to as seleniumized cells), and then the cells were collected by centrifugation at 3,000 g for 1 min. The cell pellets were resuspended in an equal volume of fresh medium and then co-incubated with CdCl2 at a final concentration of 1 mM to synthesize CdSe QDs (the biosynthesis procedure was presented in Figure S1). After incubation for 24 hours, the fluorescence intensity was measured. The cells were cultivated under 30°C at 200 revolutions per minute (rpm) (cells’ growing curves are shown in Figure S2).

    To avoid influences of the cell biomass, all the cultures were adjusted to the same cell density (optical density [OD]600០.0) before co-incubating with Na2SeO3.

    Intracellular CdSe QD fluorescence intensity measurement

    Cell samples were collected after biosynthesis for fluorescence measurement (with the OD600 of 6.0) and washed with 1 mL of 1× phosphate-buffered saline (PBS, pH 7.5) and then resuspended with 1 mL of 1× PBS (pH 7.5) before fluorescence intensity determination. The intracellular CdSe QDs fluorescence intensity was calculated by subtraction of the cellular autofluorescence (seleniumized cells) from the total fluorescence. All fluorescence spectra of the cells synthesizing QDs and seleniumized cells were acquired with a Cytation3 Multi-mode Reader from BioTek (Winooski, VT, USA), with an excitation wavelength of 400 nm and the detection emission spectrum of 450 to 700 nm.

    Determination of intracellular ATP levels

    Samples of 1吆 7 cells (OD600ɣ.0) were harvested and washed with 200 μL sterilized ice-cold deionized water three times. Then, cell pellets were resuspended in 500 μL of ATP determination lysis solution and transferred to 2 mL snap-cap tubes with 0.5 g glass beads (425� μm in diameter Sigma, St Louis, MO, USA). Cells were ruptured with a Mini-Bead beater-16 (BioSpec, Bartlesville, OK, USA) for 1 min on and 1 min pause on ice to dissipate heat, and repeated for two times. Then, the tubes were centrifuged at 16,000 g, at 4°C, for 5 min, and the supernatants were collected in new centrifuge tubes. Samples were then processed and intracellular ATP concentrations were determined as described in the protocol of the ATP Assay Kit (S0026 Beyotime, Shanghai, China) with a Cytation3 Multi-mode Reader from BioTek.

    Determination of intracellular GSH levels

    Samples of 1吆 7 cells (OD600ɣ.0) were centrifuged and harvested, and then the pellets were washed with 200 μL of sterilized ice-cold 1× PBS (pH 7.5) two times. The cell pellets were resuspended with 160 μL of sterilized ice-cold deionized water, followed by the addition of 160 μL of sterilized ice-cold 20 mM HCl and 80 μL of sterilized ice-cold 5% (wt/vol) 5-sulfosalicylic acid. The suspensions were transferred to 2 mL snap-cap tubes with 0.5 g acid-washed glass beads, and the cells were ruptured with a Mini-Bead beater-16 (BioSpec) for 1 min on and 2 min pause on ice, and repeated once. Then, the tubes were centrifuged at 13,000 g, at 4°C, for 15 min, and the supernatants were collected in new centrifuge tubes. Samples were then processed and the intracellular GSH concentrations were measured as described in the protocol of the Total Glutathione Assay Kit (S0052 Beyotime) by a Cytation3 Multi-mode Reader from BioTek.

    Cell suspensions (2 mL) were centrifuged at 5,000 g for 3 min, and then the supernatants were collected in centrifuge tubes before filtering with a 0.22-μm filter unit (Merck Millipore, Billerica, MA, USA). HNO3 and water (Milli-Q, 18.2 MO Merck Millipore) were added to the solutions to get a final concentration of 1% (vol/vol), and the samples were stored at 4°C until needed. The cell pellets were collected and washed with ice-cold 1× PBS before resuspension with 500 μL of water (Milli-Q, 18.2 MO), and then transferred to polytetrafluoroethylene reactors. After 2 mL of HNO3 had been added to the samples, they were placed on a heating plate at 120°C for 2 hours and then at 90°C until the residues were around 100 μL. HNO3 (1%, vol/vol) was added to make a total volume of 1 mL and diluted to 10 mL with water (Milli-Q, 18.2 MO) prior to determination. The Se concentrations in the supernatants or inside the cells were determined with ICP-AES (IRIS II Thermo Fisher Scientific, Fair Lawn, NJ, USA). The Cd concentrations were characterized using AAS with a ContrAA 700 high-resolution continuous light source atomic absorption spectrophotometer (Analytik Jena, Jena, Germany).

    Seleno-compounds separation and characterization

    A wet weight of 0.5 g seleniumized cells was collected and washed with 1× PBS (pH 7.5), and the cell pellet was then resuspended with 1× PBS (pH 7.5). Cells were disrupted with a Mini-Bead beater-16 (BioSpec) for 1 min on and 2 min pause on ice, twice. Samples were centrifuged to get the lysate, the protein concentrations of the samples were determined by the Bradford method, 29 and then adjusted to the same value in all samples. Trypsin (100 mg) (Amersco, Solon, OH, USA) and TCS buffer (10×, 0.1 M Tris–Cl, pH 7.5, 10 mM CaCl2, 0.5% SDS) were added to the samples and then they were incubated at 37°C for 20 h. After centrifugation, the supernatants were collected and filtered with 0.22-μm filter units. Then, the samples were subjected to step-by-step ultrafiltration with 100 KD, 50 KD, 10 KD, and 3 KD Millipore ultrafiltration tubes at 4,500 rpm for 15 min. Then, the prepared samples were used for high-performance liquid chromatography (HPLC) with ICP mass spectrometry (MS) (HPLC–ICP-MS) measurement, which was performed on an Agilent 7500 ICP-MS system (Agilent, Santa Clara, CA, USA), coupled with a HPLC system (Shimadzu, Kyoto, Japan) equipped with a CAPCELL PAK C18 column (Shiseido, Tokyo, Japan).

    Improving the biosynthesis of CdSe QDs

    The promoter of gene ADK1 or SAM2 was replaced with a stronger promoter, PGAL1, through homologous recombination (details of the construction process are in Supplementary materials) to get overexpression. The modified strains were used to biosynthesize CdSe QDs cultured in YPGal medium (with galactose as an inducer of the overexpression).

    All statistical analyses were performed using Origin 2016. The schematic diagram was made with Photoshop CC. Graph data are presented as means and SD. Two-sample t-tests were used on the data where appropriate.

    Intracellular ATP concentrations during yeast QDs biosynthesis

    To confirm the influence of cellular ATP levels on the CdSe QDs biosynthesis in S. cerevisiae, cells of different processing stages of CdSe QDs biosynthesis were collected to determine the concentrations of intracellular ATP. The intracellular ATP concentrations of the WT cells were measured during a 12-hour incubation with Na2SeO3 at a final concentration of 2.5 mM a sharp peak appeared at 2 hours after the introduction of Na2SeO3 into the culture (Figure 1A). Starting from the exponential phase, the ATP concentrations of the sample cells all sharply increased right after either Na2SeO3 or CdCl2 was added to the culture, and then rapidly reduced. The ATP concentrations of the control group remained stable, with a slight fluctuation at 24- to 48-hour cultivation and after the cells were transformed to fresh medium at 48 hours, the concentrations decreased at first and then increased substantially. Clearly, the intracellular ATP concentrations had obvious changes during the process of QDs biosynthesis in the sample cells, while the control group had no such changes (Figure 1B).

    Figure 1 Intracellular ATP concentrations during CdSe QDs biosynthesis.
    Notes: (A) ATP concentrations during the 12-hour incubation with Na2SeO3 (B) ATP concentrations at different collection times during the whole process, from the cells in exponential phase to 24 hours after seleniumized cells were co-incubated with CdCl2. Vertical lines divide the process into three phases. The cell’s growing phase is from 0 to 24 hours Na2SeO3 was added at 24 hours. At 48 hours, cell pellets were transferred to fresh medium and CdCl2 was added to the culture.
    Abbreviations: OD, optical density QDs, quantum dots.

    CdSe QDs biosynthesis under low ATP synthesis conditions

    To inspect the relation between ATP synthesis and CdSe QDs biosynthesis, several ATP-synthesis-deficient strains were introduced. S. cerevisiae strain Δatp1 displays a complete loss of ATP synthase activity. 30 The determination of the intracellular ATP concentration, with half the value of WT cells, proved that the Δatp1 strain was deficient in ATP synthesis (Figure 2). The fluorescence intensity of Δatp1 was just a quarter of that in the WT cells (Figure 2), which was consistent with the ATP concentration changes of the two strains. Other gene knock-out strains that were deficient in ATP synthesis, Δatp2, 31 Δatp14, 32 and Δatp17, 33 were also applied in the synthesis of QDs to further confirm the results, with an outcome concurrent with that for the Δatp1 strain, namely much lower intracellular ATP concentrations and QDs fluorescence intensities than for WT cells (Figure 2C and D). The fluorescence intensities of Δatp2, Δatp14, and Δatp17 cells were 1/2, 2/5, and 1/4 (Figure 2C) of the WT cells, whereas the ATP concentrations were 1/2, 1/10, and 1/8 (Figure 2D) of the WT cells, respectively. The fluorescence intensities roughly conformed to the intracellular ATP concentrations.

    Figure 2 (A) Intracellular fluorescence intensities and (B) ATP concentrations of WT and Δatp1 cells, (C) intracellular fluorescence intensities and (D) ATP concentrations of some ATP-synthesis-deficient strains.
    Abbreviations: CPS, counts per second FL, fluorescence OD, optical density WT, wild-type.

    Total intracellular GSH of the Δatp1 strain and its QDs synthesis ability

    As to the specific reason for the influence of ATP on QDs biosynthesis, we suspected it was due to effects on intracellular GSH synthesis. In our previous study, we found that GSH was important for the QDs biosynthesis and the intracellular GSH content showed synchronized increase with the fluorescence intensity, 26,27 so the GSH contents were checked in this study too. Interestingly, when the total GSH concentrations were measured in the Δatp1 strain after seleniumized cells were treated with CdCl2, that tendency referred to above disappeared. Instead, the GSH level kept steady around a certain value (about 9 μM per 1 OD600 cells) with only a slight fluctuation, yet somehow was higher than that of WT cells (Figure 3A). At time 0, when the CdCl2 was added to the culture, GSH concentrations in Δatp1 and WT cells were 6.72 and 2.95 μM per 1 OD600 cells, respectively. Nevertheless, GSH concentrations in the WT cells presented a trend of gradual increase with culturing time for seleniumized cells incubated with CdCl2. The intracellular fluorescence intensities, however, had the same tendency of gradual increase in both WT and Δatp1 strains, but with a growing difference between them (Figure 3B).

    Figure 3 Intracellular GSH contents (A) and intracellular fluorescence intensities (B) of seleniumized WT and Δatp1 cells incubated with CdCl2 during the first 7 hours.
    Abbreviations: CPS, counts per second FL, fluorescence GSH, glutathione (l-γ-glutamylcysteinylglycine) OD, optical density WT, wild-type.

    Se and Cd concentrations during co-incubation with yeast cells while synthesizing QDs (ICP-AES and AAS assays)

    Both the intracellular and extracellular concentrations of Se and Cd were measured during the QDs biosynthesis processes of WT and Δatp1 strains. Within error, nearly all Se or Cd lost from the medium was absorbed into the cells (Figure 4). Although compared with the amount of Na2SeO3 (final concentration of 2.5 mM) or CdCl2 (final concentration of 1 mM) added to the culture (which was the optimized amount for CdSe QDs biosynthesis, data not shown), WT and Δatp1 both just took a small amount inside the cells they did have the same trend of absorption of both compounds (Figure 4). The differences were evident between WT and Δatp1 strains. The Se uptake of WT was quite conspicuous at first and then slowed down and stabilized after incubation for 12 hours, reaching a final concentration of 0.48۪.029 mM, whereas the Se concentration inside Δatp1 cells was negligible with a concentration of 0.04۪.016 mM. The intracellular Se differences between Δatp1 and WT began to emerge 2 hours after the addition of Na2SeO3 and increased during the culturing over the following 24 hours (Figure 4A). A similar result was observed with the uptake of Cd, the increase tendency of the two strains was the same, only with a lower concentration in Δatp1 cells. The final concentration in Δatp1 cells was 0.10۪.004 mM, less than half of that in WT cells, which was 0.27۪.045 mM (Figure 4B).

    Figure 4 Se (A) and Cd (B) concentrations inside (in) and outside (out) wild-type (WT) and Δatp1 cells, and the recovery rate of total Se or Cd added to the culture. Se/Cd-WT/Δatp1-in/out stands for Se/Cd concentrations inside/outside the WT/Δatp1 cells.

    Major seleno-amino acid determination in Δatp1 and WT cells

    Seleniumized cells of WT and Δatp1 strains were collected, and their total proteins were extracted and digested to oligopeptides with trypsin to measure the main reduced Se species inside. On the basis of the previous study, 26 organoSe compounds selenocystine [(Se-Cys)2], Se-methylseleno-L-cysteine (SeMC), and d,L-selenomethionine (Se-Met) were checked. HPLC–ICP-MS showed that the major seleno-amino acid (Se-Cys)2 signal in the WT sample was obviously stronger than that in the Δatp1 sample (Figure 5). The peak height of (Se-Cys)2 in Δatp1 cells was half that of WT cells.

    Figure 5 HPLC-ICP-MS chromatogram of Se species standards (ST) and seleno-amino acids characterized from WT and Δatp1 cells after treatment with Na2SeO3. The Se species standards are: 1, L-selenocystine 2, Se-methylseleno-L-cysteine 3, d,L-selenomethionine.
    Abbreviations: HPLC-ICP-MS, high-performance liquid chromatography–inductively coupled plasma mass spectrometry WT, wild-type.

    CdSe QDs biosynthesis under the influences of oxidative phosphorylation inhibitor or uncoupler

    To investigate the effects of ATP on the absorption of Cd and QDs crystallization, the intracellular fluorescence intensities of cells under different concentrations of oxidative phosphorylation inhibitor (NaN3) or uncoupler (2,4-dinitrophenol, DNP), which were added to the culture during the incubation of seleniumized cells with CdCl2, were measured. The results in Figure 6A show that the intracellular fluorescence intensities decreased to less than half of the normal value along with the increase in the NaN3 concentrations from 75 to 200 mM, indicating the remarkable influence of the addition of NaN3 on QDs biosynthesis. When DNP was introduced into the culture, intracellular fluorescence intensities had a slight yet noticeable reduction (Figure 6B). At the same time, Cd concentrations of seleniumized cells incubated with CdCl2 were checked in the presence of 150 mM NaN3 or 2 mM DNP, with concentrations that had visible impacts on CdSe QDs biosynthesis (Figure 6C). Differences were not noticeable in the fluorescence intensities.

    Figure 6 Intracellular fluorescence intensities of WT cells treated with different concentrations of NaN3 (A) or DNP (B), and Cd concentrations in the culture supernatants of cells samples treated with 150 mM NaN3 and 2 mM DNP (C) and when co-incubated with CdCl2.
    Abbreviations: DNP, 2,4-dinitrophenol FL, fluorescence WT, wild-type.

    QDs biosynthesis manipulation on the basis of ATP synthesis

    To investigate the use of energy influence in the manipulation of biosynthesis, genetic modification of the energy metabolism pathway was introduced. Overexpression of the ADK1 gene leads to an accumulation of ATP levels inside the cells. When the promoter of ADK1 gene was replaced with a stronger promoter, the intracellular ATP concentration increased nearly two times (Figure 7A). Figure 7B shows that the intracellular fluorescence intensity was twice that in the WT cells.

    Figure 7 Intracellular ATP concentrations (A) and FL intensities (B) of WT and PGAL1-ADK1 strains.
    Notes: The results are expressed as mean ± standard deviation, nɥ. *Pɘ.05.
    Abbreviations: FL, fluorescence OD, optical density WT, wild-type.

    This study was approached to determine the specific influence of energy on CdSe QDs biosynthesis by yeast and took advantage of it to manipulate the biosynthesis process, for the reason that most of the proteins that encapsulated the QDs were functioned in cell energy metabolism (with the result of bioinformatics analysis, data not shown).

    According to Li et al, 27 the system in the present work uses “temporally–spatially coupling strategy” to biosynthesize CdSe QDs. Within this system, there are three phases: the cell-growing phase, the cell-seleniumized phase, and the CdSe QDs crystallization phase. Herein, the influence of ATP on the biosynthesis is carried out through the three crucial phases and mainly through the latter two phases.

    To confirm the importance of the influence of ATP on biosynthesis, the intracellular ATP concentrations were determined. The ATP levels had significant variations not only in the cell-seleniumized phase (Figure 1A) but also in the whole procedure (Figure 1B). In contrast, the intracellular ATP concentrations of the control group (Figure 1B) were consistent with the cell growing state. 34 To further confirm our suspicions, we used ATP-synthesis-deficient strains (Δatp1, Δatp2, Δatp14, and Δatp17) to biosynthesize CdSe QDs. The fluorescence intensities of the strains were as expected (Figure 2A and C). The fluorescence intensities and ATP concentrations of Δatp2, Δatp14, and Δatp17 (fermentation-related genes were not taken into consideration because cells were not undertaking fermentative metabolism to generate ATP in the cell-seleniumized phase, Figures S3 and S4) were not exactly accordant (Figure 2C and D), which may be due to other influential factors. In this case, we proposed that CdSe QDs biosynthesis by yeast cells is an energy-consuming process, besides having many other necessary conditions. 27 In the quasi-biosystem referred to by Gu et al, 35 they only need several substrates and co-factors to synthesize nanomaterials, but the temperature must reach at least 80°C󈟆°C, which was significantly higher than that for the biosynthesis. Hence, it may suggest that ATP synthesis inside the cells meets with the request of energy in the biosynthesis.

    As already reported, GSH synthesis is an energy-consuming process. 36 It was reasonable to hypothesize that the ATP effects may be due to its influence on the synthesis of GSH with its important role in our system. 26,27 Interestingly, when the GSH contents were determined in the Δatp1 strain, the tendency of GSH levels increasing with fluorescence intensities, referred to above, did not appear (Figure 3A). The content level was higher than that of WT cells during the testing period, which may be because the addition of CdCl2 in seleniumized cells stimulated an increase of GSH. 37 And because the addition of Na2SeO3 would trigger the upregulation of GSH, 27 with lower intracellular Se concentration, the GSH cannot be used effectively 38 and results in the higher GSH content in Δatp1 cells at time 0 than that of WT cells. These results (Figure 3A and B) attested that within the low ATP synthesis strains (such as Δatp1) mentioned, it was the utilization rather than synthesis of GSH that was impeded by the ATP levels.

    With the results above, it was shown that ATP had a significant influence on CdSe QDs biosynthesis by yeast. Determining where ATP had its effects was important and urgent for its not insignificant role in the biosynthesis. Since there were two critical phases in the biosynthesis process, the influences of ATP were investigated through these two phases. Figure 4 shows the intracellular and the extracellular concentrations of the Se or Cd measured during the cell-seleniumized phase or the crystallization phase. The shortage of ATP synthesis caused an absorption deficiency of both elements constructing CdSe QDs, resulting in poor fluorescence intensity. We suspected this could be linked to the way that SeO3 2− or Cd 2+ entered the cells, which might need ATP. 39,40

    Once inside the cells, Se (Ʋ) ions are reduced to Se (𕒶) ions, then the Cd 2+ interacts with the seleno-precursors and forms QDs crystals. The main seleno-amino acids emerging during the biosynthesis, as the previous study suggested, 26 were detected by HPLC–ICP-MS. 41 In Figure 5, it is shown that the major seleno-amino acid peak heights in Δatp1 and WT samples were consistent with the fluorescence intensities between them (Figure 2B). The result was proof that the accumulation of the seleno-precursor was an energy-consuming process. A similar conclusion was reached for the crystallization phase. For the reason that Δatp1 cells cannot normally accumulate seleno-precursors, the oxidative phosphorylation inhibitor and uncoupler were introduced into the system instead. Both the inhibitor and uncoupler could induce the decline of the intracellular ATP concentration, except that the uncoupler had no influence on oxidoreduction. 42,43 The introduction of the DNP (uncoupler) was to ensure that the reduction of the selenite was unaffected. It turned out that NaN3 had an evident impact on the crystallization (Figure 6A), whereas DNP had a lesser yet noticeable influence (Figure 6B). Under their influence, the uptake of Cd was nearly impervious (Figure 6C) at the concentration when the intracellular ATP concentration became lower (Figure S5) (glycolysis inhibitor had been taken into consideration because the influence on ATP generation was not obvious, Figure S6). As a consequence, without altering the Cd absorption, the importance of ATP in the QDs crystallization was confirmed.

    Thus, the specific influence of ATP was made clear. Figure 8 briefly describes the QDs biosynthesis process in yeast and shows the points of ATP action. It shows that ATP guarantees the uptake of the two inorganic compounds, helps with the accumulation of the seleno-precursors, and facilitates the utilization of the GSH accumulated during the crystallization phase. As the most direct energy resource in the organisms, ATP plays indispensable roles in many biological processes, 44,45 but its influence in the biosynthesis of nanomaterials has often been overlooked. With the present work, we have determined the difference between the intrinsic processes and the constructed ones, thus allowing better application of the connection to bilaterally control the biosynthesis process.

    Figure 8 Schematic illustration of the points of action of ATP in the CdSe QDs biosynthesis process in Saccharomyces cerevisiae.
    Abbreviations: GSH, glutathione (l-γ-glutamylcysteinylglycine) QDs, quantum dots.

    It was quite feasible that we could increase the production of the QDs through accumulation of intracellular ATP according to our findings. The overexpression of the ADK1 gene leads to the augmentation of ATP content. 46 Through standard genetic techniques, 47 the PGAL1-ADK1 strain was constructed (Figures S7 and S8). The phenotype was confirmed (Figure 7A), and the fluorescence intensity was consistent with the ATP levels. As it was reported, S-adenosylmethionine (SAM), one of the intermediate species of the methionine and cysteine metabolism pathway, needs methionine as a substrate and ATP for synthesis under the action of acyltransferase. 48 Its seleno analog, SeAM, the synthesis of which is also affected by ATP content, 38 is believed to be an important intermediate compound in the biosynthesis of CdSe QDs in yeast (unpublished data). An increase in ATP could lead to the accumulation of SAM, and under Se-rich culture, it would be SeAM. By direct overexpression of SAM2 gene to achieve the SAM accumulation, a PGAL1-SAM2 strain was successfully constructed (Figures S7 and S9A), and the intracellular SAM content was checked (Figure S9B). Figure 9 shows that the fluorescence intensity of the PGAL1-SAM2 strain was about 1.5 times of that in the WT strain, which was consistent with the results of PGAL1-ADK1 strain. In summary, it does not matter whether the ATP content was indirectly increased or an intermediate compound in the process was directly increased, as it would give the same result, namely, a prominent increase in QDs biosynthesis ability.

    Figure 9 Intracellular fluorescence intensities of WT and PGAL1-SAM2 strains.
    Abbreviations: FL, fluorescence SAM, S-adenosylmethionine WT, wild-type.

    In summary, we have uncovered a vital role of ATP played in the biosynthesis of CdSe QDs by yeast without changing the QDs fluorescent properties (Figures S10 and S11). Through investigating the specific interaction processes, the biosynthesis could be promoted simply by genetic modification of the WT strain. With the present work, we can integrate substance flux and energy flux 49 in the biosynthesis process. To the best of our knowledge, the role of energy in the biosynthesis of nanoparticles in microorganisms has not been reported yet, and this is the first study that has focused on energy influence, filling the gap in the research about its mechanism. Other than complementation of the understanding of the biosynthesis process, what’s more important is that this could offer a new perspective on the manipulation of nanoparticles biosynthesis using microorganisms aiding further applications.

    This work was supported by the National Natural Science Foundation of China (21272182 and 31570090) and the National Basic Research Program of China (973 Program, number 2013CB933904). This project is partially supported by the Chinese 111 Project Grant B06018, the National Fund for Fostering Talents in Basic Sciences (J1103513), National Infrastructure of Natural Resources for Science and Technology Program of China (number NIMR-2015𔃆), and the Laboratory (Innovative) Research Fund of Wuhan University. We are grateful to XS Feng for ICP-AES and AAS measurement.

    The authors report no conflicts of interest in this work.

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    Table S1 List of strains
    Note: EUROSCRAF, European Saccharomyces cerevisiae Archive for Functional Analysis (Bad Homburg, Germany).

    Table S2 PCR primers used in this study
    Abbreviations: PCR, polymerase chain reaction SAM, S-adenosylmethionine.

    Figure S1 Schematic flow chart of the biosynthetic procedure of the CdSe QDs in Saccharomyces cerevisiae.
    Abbreviation: QDs, quantum dots.

    Figure S2 Growth curves of the strains in the present work cultured in YPGlu (A) or YPGal (B).
    Abbreviations: OD, optical density WT, wild-type YP, 1 g L 𕒵 yeast extract and 2 g L 𕒵 peptone YPGlu, YP supplied with 2 g L 𕒵 glucose YPGal, YP supplied with 2 g L 𕒵 galactose.

    Figure S3 Sugar remaining in the supernatants of different strains after cultivation for 24 hours right before Na2SeO3 was added into the culture.
    Abbreviations: A540, absorbance OD, optical density WT, wild-type.

    Figure S4 FL intensities (A) and intracellular ATP concentrations (B) of the WT, Δpfk1, and Δrgt2.
    Abbreviations: CPS, counts per second FL, fluorescence OD, optical density WT, wild-type.

    Figure S5 Intracellular ATP concentrations of WT samples and WT cells under the influences of 150 mM NaN3 or 2 mM DNP.
    Abbreviations: DNP, 2,4-dinitrophenol WT, wild-type.

    Figure S6 Intracellular ATP concentrations of the WT cells under the effect of 0.2% (wt/vol) 2-dexoglucose (2-DG) added with the introduction of CdCl2. −, blank control without 2-DG +, 0.2% 2-DG into the culture.
    Abbreviations: OD, optical density WT, wild-type.

    Figure S7 Schematic of the homologous recombination process (A) and diagnostic polymerase chain reaction strategy (B). Gene specific primers CKA and CKB match the upstream and downstream of the insert site on the chromosome F4 and R2 primers match the inner sequences of the insert DNA.

    Figure S8 Diagnostic PCR electrophoresis image for confirming the correct replacement of the promoter of the PGAL1-ADK1 strain, the primers used as referred, templates were: 1, transformant genome 2, WT genome 3, no template control.
    Abbreviation: WT, wild-type.

    Figure S9 Electrophoresis image of the diagnostic PCR products using the primers referred to in Figure S7 templates were: 1, transformant genome 2, WT genome 3, no template control (A), and intracellular SAM concentration of the WT and correct transformant cells (B).
    Abbreviations: PCR, polymerase chain reaction WT, wild-type.

    Figure S10 FL intensity of the same WT samples determined three times with a 24 hour interval.
    Abbreviations: CPS, counts per second FL, fluorescence WT, wild-type.

    Figure S11 Emission spectrums of the corresponding strains used in the present work cultured in YPGlu (A) or YPGal (B) medium.
    Abbreviations: CPS, counts per second FL, fluorescence WT, wild-type.


    Structure and mechanism of soybean ATP sulfurylase and the committed step in plant sulfur assimilation

    Enzymes of the sulfur assimilation pathway are potential targets for improving nutrient content and environmental stress responses in plants. The committed step in this pathway is catalyzed by ATP sulfurylase, which synthesizes adenosine 5'-phosphosulfate (APS) from sulfate and ATP. To better understand the molecular basis of this energetically unfavorable reaction, the x-ray crystal structure of ATP sulfurylase isoform 1 from soybean (Glycine max ATP sulfurylase) in complex with APS was determined. This structure revealed several highly conserved substrate-binding motifs in the active site and a distinct dimerization interface compared with other ATP sulfurylases but was similar to mammalian 3'-phosphoadenosine 5'-phosphosulfate synthetase. Steady-state kinetic analysis of 20 G. max ATP sulfurylase point mutants suggests a reaction mechanism in which nucleophilic attack by sulfate on the α-phosphate of ATP involves transition state stabilization by Arg-248, Asn-249, His-255, and Arg-349. The structure and kinetic analysis suggest that ATP sulfurylase overcomes the energetic barrier of APS synthesis by distorting nucleotide structure and identifies critical residues for catalysis. Mutations that alter sulfate assimilation in Arabidopsis were mapped to the structure, which provides a molecular basis for understanding their effects on the sulfur assimilation pathway.

    Keywords: Enzyme Mechanisms Enzyme Structure Plant Plant Biochemistry Protein Structure.


    Mechanisms of sodium transport in bacteria

    In some bacteria, an Na+ circuit is an important link between exergonic and endergonic membrane reactions. The physiological importance of Na+ ion cycling is described in detail for three different bacteria. Klebsiella pneumoniae fermenting citrate pumps Na+ outwards by oxaloacetate decarboxylase and uses the Na+ ion gradient thus established for citrate uptake. Another possible function of the Na+ gradient may be to drive the endergonic reduction of NAD+ with ubiquinol as electron donor. In Vibrio alginolyticus, an Na+ gradient is established by the NADH: ubiquinone oxidoreductase segment of the respiratory chain the Na+ gradient drives solute uptake, flagellar motion and possibly ATP synthesis. In Propionigenium modestum, ATP biosynthesis is entirely dependent on the Na+ ion gradient established upon decarboxylation of methylmalonyl-CoA. The three Na(+)-translocating enzymes, oxaloacetate decarboxylase of Klebsiella pneumoniae, NADH: ubiquinone oxidoreductase of Vibrio alginolyticus and ATPase (F1F0) of Propionigenium modestum have been isolated and studied with respect to structure and function. Oxaloacetate decarboxylase consists of a peripheral subunit (alpha), that catalyses the carboxyltransfer from oxaloacetate to enzyme-bound biotin. The subunits beta and gamma are firmly embedded in the membrane and catalyse the decarboxylation of the carboxybiotin enzyme, coupled to Na+ transport. A two-step mechanism has also been demonstrated for the respiratory Na+ pump. Semiquinone radicals are first formed with the electrons from NADH subsequently, these radicals dismutate in an Na(+)-dependent reaction to quinone and quinol. The ATPase of P. modestum is closely related in its structure to the F1F0 ATPase of E. coli, but uses Na+ as the coupling ion. A specific role of protons in the ATP synthesis mechanism is therefore excluded.


    Vitamin B12 and B12-Proteins

    B12-Derivatives in Electron Transfer Reactions

    Oxidation–reduction processes are of key importance for the chemistry and biology of B12. Under physiological conditions B12-derivatives may exist in three different oxidation states (Co(III), Co(II), or Co(I)), each possessing different coordination properties and reactivities: Axial coordination to the corrin-bound cobalt center depends primarily on the formal oxidation state of the cobalt ion. As a rule, the number of axial ligands decreases in parallel with the cobalt oxidation state: In the thermodynamically predominating forms of cobalt corrins, two axial ligands are bound to the diamagnetic Co(III)-center, one axial ligand is bound to the paramagnetic Co(II)-center and axial ligands are assumed to be absent at the diamagnetic Co(I)-center. Electron transfer reactions involving B12-derivatives, therefore, are accompanied by changes of the number of axial ligands and depend upon the nature of axial ligands. Axial coordination of the nucleotide base and of strongly coordinating ligands stabilizes the cobalt center against reduction and the reduction of alkyl-Co(III)-corrins typically occurs at potentials more negative than that of the Co(II)/Co(I)-redox-pair B12r/B12s.


    4.1 Energy and Metabolism

    Scientists use the term bioenergetics to describe the concept of energy flow (Figure 4.2) through living systems, such as cells. Cellular processes such as the building and breaking down of complex molecules occur through stepwise chemical reactions. Some of these chemical reactions are spontaneous and release energy, whereas others require energy to proceed. Just as living things must continually consume food to replenish their energy supplies, cells must continually obtain more energy to replenish that used by the many energy-requiring chemical reactions that constantly take place. Together, all of the chemical reactions that take place inside cells, including those that consume or generate energy, are referred to as the cell’s metabolism .

    Metabolic Pathways

    Consider the metabolism of sugar. This is a classic example of one of the many cellular processes that use and produce energy. Living things consume sugars as a major energy source, because sugar molecules have a great deal of energy stored within their bonds. For the most part, photosynthesizing organisms like plants produce these sugars. During photosynthesis, plants use energy (originally from sunlight) to convert carbon dioxide gas (CO2) into sugar molecules (like glucose: C6H12O6). They consume carbon dioxide and produce oxygen as a waste product. This reaction is summarized as:

    Because this process involves synthesizing an energy-storing molecule, it requires energy input to proceed. During the light reactions of photosynthesis, energy is provided by a molecule called adenosine triphosphate (ATP), which is the primary energy currency of all cells. Just as the dollar is used as currency to buy goods, cells use molecules of ATP as energy currency to perform immediate work. In contrast, energy-storage molecules such as glucose are consumed only to be broken down to use their energy. The reaction that harvests the energy of a sugar molecule in cells requiring oxygen to survive can be summarized by the reverse reaction to photosynthesis. In this reaction, oxygen is consumed and carbon dioxide is released as a waste product. The reaction is summarized as:

    Both of these reactions involve many steps.

    The processes of making and breaking down sugar molecules illustrate two examples of metabolic pathways. A metabolic pathway is a series of chemical reactions that takes a starting molecule and modifies it, step-by-step, through a series of metabolic intermediates, eventually yielding a final product. In the example of sugar metabolism, the first metabolic pathway synthesized sugar from smaller molecules, and the other pathway broke sugar down into smaller molecules. These two opposite processes—the first requiring energy and the second producing energy—are referred to as anabolic pathways (building polymers) and catabolic pathways (breaking down polymers into their monomers), respectively. Consequently, metabolism is composed of synthesis (anabolism) and degradation (catabolism) (Figure 4.3).

    It is important to know that the chemical reactions of metabolic pathways do not take place on their own. Each reaction step is facilitated, or catalyzed, by a protein called an enzyme. Enzymes are important for catalyzing all types of biological reactions—those that require energy as well as those that release energy.

    Energy

    Thermodynamics refers to the study of energy and energy transfer involving physical matter. The matter relevant to a particular case of energy transfer is called a system, and everything outside of that matter is called the surroundings. For instance, when heating a pot of water on the stove, the system includes the stove, the pot, and the water. Energy is transferred within the system (between the stove, pot, and water). There are two types of systems: open and closed. In an open system, energy can be exchanged with its surroundings. The stovetop system is open because heat can be lost to the air. A closed system cannot exchange energy with its surroundings.

    Biological organisms are open systems. Energy is exchanged between them and their surroundings as they use energy from the sun to perform photosynthesis or consume energy-storing molecules and release energy to the environment by doing work and releasing heat. Like all things in the physical world, energy is subject to physical laws. The laws of thermodynamics govern the transfer of energy in and among all systems in the universe.

    In general, energy is defined as the ability to do work, or to create some kind of change. Energy exists in different forms. For example, electrical energy, light energy, and heat energy are all different types of energy. To appreciate the way energy flows into and out of biological systems, it is important to understand two of the physical laws that govern energy.

    Thermodynamics

    The first law of thermodynamics states that the total amount of energy in the universe is constant and conserved. In other words, there has always been, and always will be, exactly the same amount of energy in the universe. Energy exists in many different forms. According to the first law of thermodynamics, energy may be transferred from place to place or transformed into different forms, but it cannot be created or destroyed. The transfers and transformations of energy take place around us all the time. Light bulbs transform electrical energy into light and heat energy. Gas stoves transform chemical energy from natural gas into heat energy. Plants perform one of the most biologically useful energy transformations on earth: that of converting the energy of sunlight to chemical energy stored within organic molecules (Figure 4.2). Some examples of energy transformations are shown in Figure 4.4.

    The challenge for all living organisms is to obtain energy from their surroundings in forms that they can transfer or transform into usable energy to do work. Living cells have evolved to meet this challenge. Chemical energy stored within organic molecules such as sugars and fats is transferred and transformed through a series of cellular chemical reactions into energy within molecules of ATP. Energy in ATP molecules is easily accessible to do work. Examples of the types of work that cells need to do include building complex molecules, transporting materials, powering the motion of cilia or flagella, and contracting muscle fibers to create movement.

    A living cell’s primary tasks of obtaining, transforming, and using energy to do work may seem simple. However, the second law of thermodynamics explains why these tasks are harder than they appear. All energy transfers and transformations are never completely efficient. In every energy transfer, some amount of energy is lost in a form that is unusable. In most cases, this form is heat energy. Thermodynamically, heat energy is defined as the energy transferred from one system to another that is not work. For example, when a light bulb is turned on, some of the energy being converted from electrical energy into light energy is lost as heat energy. Likewise, some energy is lost as heat energy during cellular metabolic reactions.

    An important concept in physical systems is that of order and disorder. The more energy that is lost by a system to its surroundings, the less ordered and more random the system is. Scientists refer to the measure of randomness or disorder within a system as entropy. High entropy means high disorder and low energy. Molecules and chemical reactions have varying entropy as well. For example, entropy increases as molecules at a high concentration in one place diffuse and spread out. The second law of thermodynamics says that energy will always be lost as heat in energy transfers or transformations.

    Living things are highly ordered, requiring constant energy input to be maintained in a state of low entropy.

    Potential and Kinetic Energy

    When an object is in motion, there is energy associated with that object. Think of a wrecking ball. Even a slow-moving wrecking ball can do a great deal of damage to other objects. Energy associated with objects in motion is called kinetic energy (Figure 4.5). A speeding bullet, a walking person, and the rapid movement of molecules in the air (which produces heat) all have kinetic energy.

    Now what if that same motionless wrecking ball is lifted two stories above ground with a crane? If the suspended wrecking ball is unmoving, is there energy associated with it? The answer is yes. The energy that was required to lift the wrecking ball did not disappear, but is now stored in the wrecking ball by virtue of its position and the force of gravity acting on it. This type of energy is called potential energy (Figure 4.5). If the ball were to fall, the potential energy would be transformed into kinetic energy until all of the potential energy was exhausted when the ball rested on the ground. Wrecking balls also swing like a pendulum through the swing, there is a constant change of potential energy (highest at the top of the swing) to kinetic energy (highest at the bottom of the swing). Other examples of potential energy include the energy of water held behind a dam or a person about to skydive out of an airplane.

    Potential energy is not only associated with the location of matter, but also with the structure of matter. Even a spring on the ground has potential energy if it is compressed so does a rubber band that is pulled taut. On a molecular level, the bonds that hold the atoms of molecules together exist in a particular structure that has potential energy. Remember that anabolic cellular pathways require energy to synthesize complex molecules from simpler ones and catabolic pathways release energy when complex molecules are broken down. The fact that energy can be released by the breakdown of certain chemical bonds implies that those bonds have potential energy. In fact, there is potential energy stored within the bonds of all the food molecules we eat, which is eventually harnessed for use. This is because these bonds can release energy when broken. The type of potential energy that exists within chemical bonds, and is released when those bonds are broken, is called chemical energy. Chemical energy is responsible for providing living cells with energy from food. The release of energy occurs when the molecular bonds within food molecules are broken.

    Concepts in Action

    Visit the site and select “Pendulum” from the “Work and Energy” menu to see the shifting kinetic and potential energy of a pendulum in motion.

    Free and Activation Energy

    After learning that chemical reactions release energy when energy-storing bonds are broken, an important next question is the following: How is the energy associated with these chemical reactions quantified and expressed? How can the energy released from one reaction be compared to that of another reaction? A measurement of free energy is used to quantify these energy transfers. Recall that according to the second law of thermodynamics, all energy transfers involve the loss of some amount of energy in an unusable form such as heat. Free energy specifically refers to the energy associated with a chemical reaction that is available after the losses are accounted for. In other words, free energy is usable energy, or energy that is available to do work.

    If energy is released during a chemical reaction, then the change in free energy, signified as ∆G (delta G) will be a negative number. A negative change in free energy also means that the products of the reaction have less free energy than the reactants, because they release some free energy during the reaction. Reactions that have a negative change in free energy and consequently release free energy are called exergonic reactions . Think: exergonic means energy is exiting the system. These reactions are also referred to as spontaneous reactions, and their products have less stored energy than the reactants. An important distinction must be drawn between the term spontaneous and the idea of a chemical reaction occurring immediately. Contrary to the everyday use of the term, a spontaneous reaction is not one that suddenly or quickly occurs. The rusting of iron is an example of a spontaneous reaction that occurs slowly, little by little, over time.

    If a chemical reaction absorbs energy rather than releases energy on balance, then the ∆G for that reaction will be a positive value. In this case, the products have more free energy than the reactants. Thus, the products of these reactions can be thought of as energy-storing molecules. These chemical reactions are called endergonic reactions and they are non-spontaneous. An endergonic reaction will not take place on its own without the addition of free energy.

    Visual Connection

    Look at each of the processes shown and decide if it is endergonic or exergonic.

    There is another important concept that must be considered regarding endergonic and exergonic reactions. Exergonic reactions require a small amount of energy input to get going, before they can proceed with their energy-releasing steps. These reactions have a net release of energy, but still require some energy input in the beginning. This small amount of energy input necessary for all chemical reactions to occur is called the activation energy .

    Concepts in Action

    Watch an animation of the move from free energy to transition state of the reaction.

    Enzymes

    A substance that helps a chemical reaction to occur is called a catalyst, and the molecules that catalyze biochemical reactions are called enzymes . Most enzymes are proteins and perform the critical task of lowering the activation energies of chemical reactions inside the cell. Most of the reactions critical to a living cell happen too slowly at normal temperatures to be of any use to the cell. Without enzymes to speed up these reactions, life could not persist. Enzymes do this by binding to the reactant molecules and holding them in such a way as to make the chemical bond-breaking and -forming processes take place more easily. It is important to remember that enzymes do not change whether a reaction is exergonic (spontaneous) or endergonic. This is because they do not change the free energy of the reactants or products. They only reduce the activation energy required for the reaction to go forward (Figure 4.7). In addition, an enzyme itself is unchanged by the reaction it catalyzes. Once one reaction has been catalyzed, the enzyme is able to participate in other reactions.

    The chemical reactants to which an enzyme binds are called the enzyme’s substrates . There may be one or more substrates, depending on the particular chemical reaction. In some reactions, a single reactant substrate is broken down into multiple products. In others, two substrates may come together to create one larger molecule. Two reactants might also enter a reaction and both become modified, but they leave the reaction as two products. The location within the enzyme where the substrate binds is called the enzyme’s active site . The active site is where the “action” happens. Since enzymes are proteins, there is a unique combination of amino acid side chains within the active site. Each side chain is characterized by different properties. They can be large or small, weakly acidic or basic, hydrophilic or hydrophobic, positively or negatively charged, or neutral. The unique combination of side chains creates a very specific chemical environment within the active site. This specific environment is suited to bind to one specific chemical substrate (or substrates).

    Active sites are subject to influences of the local environment. Increasing the environmental temperature generally increases reaction rates, enzyme-catalyzed or otherwise. However, temperatures outside of an optimal range reduce the rate at which an enzyme catalyzes a reaction. Hot temperatures will eventually cause enzymes to denature, an irreversible change in the three-dimensional shape and therefore the function of the enzyme. Enzymes are also suited to function best within a certain pH and salt concentration range, and, as with temperature, extreme pH, and salt concentrations can cause enzymes to denature.

    For many years, scientists thought that enzyme-substrate binding took place in a simple “lock and key” fashion. This model asserted that the enzyme and substrate fit together perfectly in one instantaneous step. However, current research supports a model called induced fit (Figure 4.8). The induced-fit model expands on the lock-and-key model by describing a more dynamic binding between enzyme and substrate. As the enzyme and substrate come together, their interaction causes a mild shift in the enzyme’s structure that forms an ideal binding arrangement between enzyme and substrate.

    Concepts in Action

    When an enzyme binds its substrate, an enzyme-substrate complex is formed. This complex lowers the activation energy of the reaction and promotes its rapid progression in one of multiple possible ways. On a basic level, enzymes promote chemical reactions that involve more than one substrate by bringing the substrates together in an optimal orientation for reaction. Another way in which enzymes promote the reaction of their substrates is by creating an optimal environment within the active site for the reaction to occur. The chemical properties that emerge from the particular arrangement of amino acid R groups within an active site create the perfect environment for an enzyme’s specific substrates to react.

    The enzyme-substrate complex can also lower activation energy by compromising the bond structure so that it is easier to break. Finally, enzymes can also lower activation energies by taking part in the chemical reaction itself. In these cases, it is important to remember that the enzyme will always return to its original state by the completion of the reaction. One of the hallmark properties of enzymes is that they remain ultimately unchanged by the reactions they catalyze. After an enzyme has catalyzed a reaction, it releases its product(s) and can catalyze a new reaction.

    It would seem ideal to have a scenario in which all of an organism's enzymes existed in abundant supply and functioned optimally under all cellular conditions, in all cells, at all times. However, a variety of mechanisms ensures that this does not happen. Cellular needs and conditions constantly vary from cell to cell, and change within individual cells over time. The required enzymes of stomach cells differ from those of fat storage cells, skin cells, blood cells, and nerve cells. Furthermore, a digestive organ cell works much harder to process and break down nutrients during the time that closely follows a meal compared with many hours after a meal. As these cellular demands and conditions vary, so must the amounts and functionality of different enzymes.

    Since the rates of biochemical reactions are controlled by activation energy, and enzymes lower and determine activation energies for chemical reactions, the relative amounts and functioning of the variety of enzymes within a cell ultimately determine which reactions will proceed and at what rates. This determination is tightly controlled in cells. In certain cellular environments, enzyme activity is partly controlled by environmental factors like pH, temperature, salt concentration, and, in some cases, cofactors or coenzymes.

    Enzymes can also be regulated in ways that either promote or reduce enzyme activity. There are many kinds of molecules that inhibit or promote enzyme function, and various mechanisms by which they do so. In some cases of enzyme inhibition, an inhibitor molecule is similar enough to a substrate that it can bind to the active site and simply block the substrate from binding. When this happens, the enzyme is inhibited through competitive inhibition , because an inhibitor molecule competes with the substrate for binding to the active site.

    On the other hand, in noncompetitive inhibition , an inhibitor molecule binds to the enzyme in a location other than the active site, called an allosteric site, but still manages to prevent substrate binding to the active site. Some inhibitor molecules bind to enzymes in a location where their binding induces a conformational change that reduces the enzyme activity as it no longer effectively catalyzes the conversion of the substrate to product. This type of inhibition is called allosteric inhibition (Figure 4.9). Most allosterically regulated enzymes are made up of more than one polypeptide, meaning that they have more than one protein subunit. When an allosteric inhibitor binds to a region on an enzyme, all active sites on the protein subunits are changed slightly such that they bind their substrates with less efficiency. There are allosteric activators as well as inhibitors. Allosteric activators bind to locations on an enzyme away from the active site, inducing a conformational change that increases the affinity of the enzyme’s active site(s) for its substrate(s) (Figure 4.9).

    Career Connection

    Pharmaceutical Drug Developer

    Enzymes are key components of metabolic pathways. Understanding how enzymes work and how they can be regulated are key principles behind the development of many of the pharmaceutical drugs on the market today. Biologists working in this field collaborate with other scientists to design drugs (Figure 4.10).

    Consider statins for example—statins is the name given to one class of drugs that can reduce cholesterol levels. These compounds are inhibitors of the enzyme HMG-CoA reductase, which is the enzyme that synthesizes cholesterol from lipids in the body. By inhibiting this enzyme, the level of cholesterol synthesized in the body can be reduced. Similarly, acetaminophen, popularly marketed under the brand name Tylenol, is an inhibitor of the enzyme cyclooxygenase. While it is used to provide relief from fever and inflammation (pain), its mechanism of action is still not completely understood.

    How are drugs discovered? One of the biggest challenges in drug discovery is identifying a drug target. A drug target is a molecule that is literally the target of the drug. In the case of statins, HMG-CoA reductase is the drug target. Drug targets are identified through painstaking research in the laboratory. Identifying the target alone is not enough scientists also need to know how the target acts inside the cell and which reactions go awry in the case of disease. Once the target and the pathway are identified, then the actual process of drug design begins. In this stage, chemists and biologists work together to design and synthesize molecules that can block or activate a particular reaction. However, this is only the beginning: If and when a drug prototype is successful in performing its function, then it is subjected to many tests from in vitro experiments to clinical trials before it can get approval from the U.S. Food and Drug Administration to be on the market.

    Many enzymes do not work optimally, or even at all, unless bound to other specific non-protein helper molecules. They may bond either temporarily through ionic or hydrogen bonds, or permanently through stronger covalent bonds. Binding to these molecules promotes optimal shape and function of their respective enzymes. Two examples of these types of helper molecules are cofactors and coenzymes. Cofactors are inorganic ions such as ions of iron and magnesium. Coenzymes are organic helper molecules, those with a basic atomic structure made up of carbon and hydrogen. Like enzymes, these molecules participate in reactions without being changed themselves and are ultimately recycled and reused. Vitamins are the source of coenzymes. Some vitamins are the precursors of coenzymes and others act directly as coenzymes. Vitamin C is a direct coenzyme for multiple enzymes that take part in building the important connective tissue, collagen. Therefore, enzyme function is, in part, regulated by the abundance of various cofactors and coenzymes, which may be supplied by an organism’s diet or, in some cases, produced by the organism.

    Feedback Inhibition in Metabolic Pathways

    Molecules can regulate enzyme function in many ways. The major question remains, however: What are these molecules and where do they come from? Some are cofactors and coenzymes, as you have learned. What other molecules in the cell provide enzymatic regulation such as allosteric modulation, and competitive and non-competitive inhibition? Perhaps the most relevant sources of regulatory molecules, with respect to enzymatic cellular metabolism, are the products of the cellular metabolic reactions themselves. In a most efficient and elegant way, cells have evolved to use the products of their own reactions for feedback inhibition of enzyme activity. Feedback inhibition involves the use of a reaction product to regulate its own further production (Figure 4.11). The cell responds to an abundance of the products by slowing down production during anabolic or catabolic reactions. Such reaction products may inhibit the enzymes that catalyzed their production through the mechanisms described above.

    The production of both amino acids and nucleotides is controlled through feedback inhibition. Additionally, ATP is an allosteric regulator of some of the enzymes involved in the catabolic breakdown of sugar, the process that creates ATP. In this way, when ATP is in abundant supply, the cell can prevent the production of ATP. On the other hand, ADP serves as a positive allosteric regulator (an allosteric activator) for some of the same enzymes that are inhibited by ATP. Thus, when relative levels of ADP are high compared to ATP, the cell is triggered to produce more ATP through sugar catabolism.


    ATP formation during photosynthesis

    Photosynthesis generates ATP by a mechanism that is similar in principle, if not in detail. The organelles responsible are different from mitochondria, but they also form membrane-bounded closed sacs ( thylakoids) often arranged in stacks (grana). Solar energy splits two molecules of H2O into molecular oxygen (O2), four protons (H + ), and four electrons.

    This is the source of oxygen evolution, clearly visible as bubbles from underwater plants in bright sunshine. The process involves a chlorophyll molecule, P680, that changes its redox potential from +820 millivolts (in which there is a tendency to accept electrons) to about −680 millivolts (in which there is a tendency to lose electrons) upon excitation with light and acquisition of electrons. The electrons are subsequently passed along a series of carriers (plastoquinone, cytochromes b and f, and plastocyanin), analogous to the mitochondrial respiratory chain. This process pumps protons across the membrane from the outside of the thylakoid membrane to the inside. Protons (H + ) do not move freely across the membrane although chloride ions (Cl - ) do, creating a pH gradient. An ATP synthetase enzyme similar to that of the mitochondria is present, but on the outside of the thylakoid membrane. Passage of protons (H + ) through it from inside to outside generates ATP.

    Hence, a gradient of protons (H + ) across the membrane is the high-energy intermediate for forming ATP in plant photosynthesis and in the respiration of all cells capable of passing reducing equivalents (hydrogen atoms or electrons) to electron acceptors.


    About the mechanism of coupled reaction/metabolism with ATP - Biology

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    Metabolism and Cellular Respiration

    What is metabolism?
    All living things must have an unceasing supply of energy and matter. The transformation of this energy and matter within the body is called metabolism. Metabolism includes two different types: catabolism and anabolism. Catabolism is destructive metabolism. Typically, in catabolism, larger organic molecules are broken down into smaller constituents. This usually occurs with the release of energy. Anabolism is constructive metabolism. Typically, in anabolism, small precursor molecules are assembled into larger organic molecules. This always requires the input of energy.

    Anabolism and catabolism Pathways
    Anabolism is the synthesis of complex molecules from precursors. This includes synthesis of proteins, carbohydrates, nucleic acids and lipids, usually from their building block monomers. Catabolism is the breakdown of complex molecules into smaller precursors from which they are synthesized. It is a reversed process of anabolism. When cells have excess resources such as food and extra energy, anabolism occurs to store unused nutrients for later use. When cells are deficient for food or energy, catabolism occurs to break down the stored nutrients for the body to use.

    Energetics of biological Reactions
    Biological energy is the capacity to run biochemical reactions to enable the cells to do their work. Free energy (G) relates temperature, enthalpy and entropy. Free energy is used to determine if the reaction is spontaneous at a specific temperature.

    Determining spontaneity of a process
    Free energy describes whether a reaction will occur spontaneously. The First Law of Thermodynamics states that energy is conserved: energy can neither be created nor destroyed. The Second Law of Thermodynamics states that the work produced from a given energy can never be 100% efficient. In metabolism, reactions which are spontaneous are favorable because these run automatically and release free energy. Every reaction has an activation energy, which describes an energy barrier that is overcome every time the reaction occurs. Most of the reactions in the cell require enzymes. Enzymes are proteins to speed up reactions by grabbing onto reactants to bring them closer together. Reactants which are closer together can reach activation energy more easily. Thus, enzymes lower activation energy and speed up the reaction.

    ATP
    ATP is the energy currency of all cells. Most of the reactions in the cell require ATP. ATP is energy rich. When the energy is used by a reaction, ATP breaks up into ADP and Pi. In order to use the energy again, ADP and Pi must be changed back into ATP. This requires energy. Non-spontaneous reactions requires energy, and this is often done by coupling this reaction with an ATP breaking down reaction, the combined free energy will be negative and therefore enables the overall reaction.

    Cellular Respiration
    Cellular respiration is a series of metabolic processes which all living cells use to produce energy in the form of ATP. In cellular respiration, the cell breaks down glucose to produce large amounts of energy in the form of ATP. Cellular respiration can take two paths: aerobic respiration or anaerobic respiration. Aerobic respiration occurs when oxygen is available, whereas anaerobic respiration occurs when oxygen is not available. The two paths of cellular respiration share the glycolysis step. Aerobic respiration has three steps: glycolysis, Krebs cycle, and oxidative phosphorylation. During glycolysis, glucose is broken down into pyruvate and produces 2 ATP. The Krebs cycle is also known as TCA cycle which contains a series of Redox reactions to convert pyruvate into CO2 and produce NADH and FADH2. During oxidative phosphorylation, NADH and FADH2 are used as substrate to generate a pH gradient on mitochondria membrane which is used to generate ATP via ATP synthase. Anaerobic respiration contains two steps: glycolysis and fermentation. Fermentation regenerates the reactants needed for glycolysis to run again. Fermentation converts pyruvate into ethanol or lactic acid, and in the process regenerates intermediates for glycolysis.

    Metabolism includes catabolism and anabolism. Anabolism is the synthesis of complex molecules from precursors, while catabolism is the breakdown of complex molecules into smaller precursors from which they are synthesized. All these pathways involve biochemical reactions. Free energy describes whether a reaction will occur spontaneously. In metabolism, reactions which are spontaneous are favorable because these run automatically and release free energy. Every reaction has an activation energy which can be lowered down by enzymes. Enzymes do this by bringing the reactants closer together. ATP is the energy currency of all cells. Most of the reactions in the cell require ATP. A non-spontaneous reaction can be coupled to ATP hydrolysis reaction to enable the overall reaction release free energy and therefore become favorable. ATP is generated by cellular respiration, which contains fermentation (anaerobic respiration) and the Krebs cycle (aerobic fermentation).


    Watch the video: G Protein Coupled ReceptorsGPCRs - Structure, Function, Mechanism of Action. Everything! (May 2022).