7.4: Oxidative Phosphorylation - Biology

7.4: Oxidative Phosphorylation - Biology

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Skills to Develop

  • Describe how electrons move through the electron transport chain and what happens to their energy levels
  • Explain how a proton (H+) gradient is established and maintained by the electron transport chain

You have just read about two pathways in glucose catabolism—glycolysis and the citric acid cycle—that generate ATP. Most of the ATP generated during the aerobic catabolism of glucose, however, is not generated directly from these pathways. Rather, it is derived from a process that begins with moving electrons through a series of electron transporters that undergo redox reactions. This causes hydrogen ions to accumulate within the matrix space. Therefore, a concentration gradient forms in which hydrogen ions diffuse out of the matrix space by passing through ATP synthase. The current of hydrogen ions powers the catalytic action of ATP synthase, which phosphorylates ADP, producing ATP.

Electron Transport Chain

The electron transport chain (Figure (PageIndex{1})) is the last component of aerobic respiration and is the only part of glucose metabolism that uses atmospheric oxygen. Oxygen continuously diffuses into plants; in animals, it enters the body through the respiratory system. Electron transport is a series of redox reactions that resemble a relay race or bucket brigade in that electrons are passed rapidly from one component to the next, to the endpoint of the chain where the electrons reduce molecular oxygen, producing water. There are four complexes composed of proteins, labeled I through IV in Figure (PageIndex{1}), and the aggregation of these four complexes, together with associated mobile, accessory electron carriers, is called the electron transport chain. The electron transport chain is present in multiple copies in the inner mitochondrial membrane of eukaryotes and the plasma membrane of prokaryotes.

Complex I

To start, two electrons are carried to the first complex aboard NADH. This complex, labeled I, is composed of flavin mononucleotide (FMN) and an iron-sulfur (Fe-S)-containing protein. FMN, which is derived from vitamin B2, also called riboflavin, is one of several prosthetic groups or co-factors in the electron transport chain. A prosthetic group is a non-protein molecule required for the activity of a protein. Prosthetic groups are organic or inorganic, non-peptide molecules bound to a protein that facilitate its function; prosthetic groups include co-enzymes, which are the prosthetic groups of enzymes. The enzyme in complex I is NADH dehydrogenase and is a very large protein, containing 45 amino acid chains. Complex I can pump four hydrogen ions across the membrane from the matrix into the intermembrane space, and it is in this way that the hydrogen ion gradient is established and maintained between the two compartments separated by the inner mitochondrial membrane.

Q and Complex II

Complex II directly receives FADH2, which does not pass through complex I. The compound connecting the first and second complexes to the third is ubiquinone (Q). The Q molecule is lipid soluble and freely moves through the hydrophobic core of the membrane. Once it is reduced, (QH2), ubiquinone delivers its electrons to the next complex in the electron transport chain. Q receives the electrons derived from NADH from complex I and the electrons derived from FADH2 from complex II, including succinate dehydrogenase. This enzyme and FADH2 form a small complex that delivers electrons directly to the electron transport chain, bypassing the first complex. Since these electrons bypass and thus do not energize the proton pump in the first complex, fewer ATP molecules are made from the FADH2 electrons. The number of ATP molecules ultimately obtained is directly proportional to the number of protons pumped across the inner mitochondrial membrane.

Complex III

The third complex is composed of cytochrome b, another Fe-S protein, Rieske center (2Fe-2S center), and cytochrome c proteins; this complex is also called cytochrome oxidoreductase. Cytochrome proteins have a prosthetic group of heme. The heme molecule is similar to the heme in hemoglobin, but it carries electrons, not oxygen. As a result, the iron ion at its core is reduced and oxidized as it passes the electrons, fluctuating between different oxidation states: Fe++ (reduced) and Fe+++ (oxidized). The heme molecules in the cytochromes have slightly different characteristics due to the effects of the different proteins binding them, giving slightly different characteristics to each complex. Complex III pumps protons through the membrane and passes its electrons to cytochrome c for transport to the fourth complex of proteins and enzymes (cytochrome c is the acceptor of electrons from Q; however, whereas Q carries pairs of electrons, cytochrome c can accept only one at a time).

Complex IV

The fourth complex is composed of cytochrome proteins c, a, and a3. This complex contains two heme groups (one in each of the two cytochromes, a, and a3) and three copper ions (a pair of CuA and one CuB in cytochrome a3). The cytochromes hold an oxygen molecule very tightly between the iron and copper ions until the oxygen is completely reduced. The reduced oxygen then picks up two hydrogen ions from the surrounding medium to make water (H2O). The removal of the hydrogen ions from the system contributes to the ion gradient used in the process of chemiosmosis.


In chemiosmosis, the free energy from the series of redox reactions just described is used to pump hydrogen ions (protons) across the membrane. The uneven distribution of H+ ions across the membrane establishes both concentration and electrical gradients (thus, an electrochemical gradient), owing to the hydrogen ions’ positive charge and their aggregation on one side of the membrane.

If the membrane were open to diffusion by the hydrogen ions, the ions would tend to diffuse back across into the matrix, driven by their electrochemical gradient. Recall that many ions cannot diffuse through the nonpolar regions of phospholipid membranes without the aid of ion channels. Similarly, hydrogen ions in the matrix space can only pass through the inner mitochondrial membrane through an integral membrane protein called ATP synthase (Figure (PageIndex{2})). This complex protein acts as a tiny generator, turned by the force of the hydrogen ions diffusing through it, down their electrochemical gradient. The turning of parts of this molecular machine facilitates the addition of a phosphate to ADP, forming ATP, using the potential energy of the hydrogen ion gradient.

Art Connection

Dinitrophenol (DNP) is an uncoupler that makes the inner mitochondrial membrane leaky to protons. It was used until 1938 as a weight-loss drug. What effect would you expect DNP to have on the change in pH across the inner mitochondrial membrane? Why do you think this might be an effective weight-loss drug?

Chemiosmosis (Figure (PageIndex{3})) is used to generate 90 percent of the ATP made during aerobic glucose catabolism; it is also the method used in the light reactions of photosynthesis to harness the energy of sunlight in the process of photophosphorylation. Recall that the production of ATP using the process of chemiosmosis in mitochondria is called oxidative phosphorylation. The overall result of these reactions is the production of ATP from the energy of the electrons removed from hydrogen atoms. These atoms were originally part of a glucose molecule. At the end of the pathway, the electrons are used to reduce an oxygen molecule to oxygen ions. The extra electrons on the oxygen attract hydrogen ions (protons) from the surrounding medium, and water is formed.

Art Connection

Cyanide inhibits cytochrome c oxidase, a component of the electron transport chain. If cyanide poisoning occurs, would you expect the pH of the intermembrane space to increase or decrease? What effect would cyanide have on ATP synthesis?

ATP Yield

The number of ATP molecules generated from the catabolism of glucose varies. For example, the number of hydrogen ions that the electron transport chain complexes can pump through the membrane varies between species. Another source of variance stems from the shuttle of electrons across the membranes of the mitochondria. (The NADH generated from glycolysis cannot easily enter mitochondria.) Thus, electrons are picked up on the inside of mitochondria by either NAD+ or FAD+. As you have learned earlier, these FAD+ molecules can transport fewer ions; consequently, fewer ATP molecules are generated when FAD+ acts as a carrier. NAD+ is used as the electron transporter in the liver and FAD+ acts in the brain.

Another factor that affects the yield of ATP molecules generated from glucose is the fact that intermediate compounds in these pathways are used for other purposes. Glucose catabolism connects with the pathways that build or break down all other biochemical compounds in cells, and the result is somewhat messier than the ideal situations described thus far. For example, sugars other than glucose are fed into the glycolytic pathway for energy extraction. Moreover, the five-carbon sugars that form nucleic acids are made from intermediates in glycolysis. Certain nonessential amino acids can be made from intermediates of both glycolysis and the citric acid cycle. Lipids, such as cholesterol and triglycerides, are also made from intermediates in these pathways, and both amino acids and triglycerides are broken down for energy through these pathways. Overall, in living systems, these pathways of glucose catabolism extract about 34 percent of the energy contained in glucose.


The electron transport chain is the portion of aerobic respiration that uses free oxygen as the final electron acceptor of the electrons removed from the intermediate compounds in glucose catabolism. The electron transport chain is composed of four large, multiprotein complexes embedded in the inner mitochondrial membrane and two small diffusible electron carriers shuttling electrons between them. The electrons are passed through a series of redox reactions, with a small amount of free energy used at three points to transport hydrogen ions across a membrane. This process contributes to the gradient used in chemiosmosis. The electrons passing through the electron transport chain gradually lose energy, High-energy electrons donated to the chain by either NADH or FADH2 complete the chain, as low-energy electrons reduce oxygen molecules and form water. The level of free energy of the electrons drops from about 60 kcal/mol in NADH or 45 kcal/mol in FADH2 to about 0 kcal/mol in water. The end products of the electron transport chain are water and ATP. A number of intermediate compounds of the citric acid cycle can be diverted into the anabolism of other biochemical molecules, such as nonessential amino acids, sugars, and lipids. These same molecules can serve as energy sources for the glucose pathways.

Art Connections

[link] Dinitrophenol (DNP) is an uncoupler that makes the inner mitochondrial membrane leaky to protons. What effect would you expect DNP to have on the change in pH across the inner mitochondrial membrane? Why do you think this might be an effective weight-loss drug?

[link] After DNP poisoning, the electron transport chain can no longer form a proton gradient, and ATP synthase can no longer make ATP. DNP is an effective diet drug because it uncouples ATP synthesis; in other words, after taking it, a person obtains less energy out of the food he or she eats. Interestingly, one of the worst side effects of this drug is hyperthermia, or overheating of the body. Since ATP cannot be formed, the energy from electron transport is lost as heat.

[link] Cyanide inhibits cytochrome c oxidase, a component of the electron transport chain. If cyanide poisoning occurs, would you expect the pH of the intermembrane space to increase or decrease? What effect would cyanide have on ATP synthesis?

[link] After cyanide poisoning, the electron transport chain can no longer pump electrons into the intermembrane space. The pH of the intermembrane space would increase, the pH gradient would decrease, and ATP synthesis would stop.

ATP synthase
(also, F1F0 ATP synthase) membrane-embedded protein complex that adds a phosphate to ADP with energy from protons diffusing through it
prosthetic group
(also, prosthetic cofactor) molecule bound to a protein that facilitates the function of the protein
soluble electron transporter in the electron transport chain that connects the first or second complex to the third

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

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

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

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

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2,4-Dinitrophenol fails to stimulate the breakdown of the soluble high energy intermediate of oxidative phosphorylation isolated from Alcaligenes faecalis extracts.

Once the intermediate is broken down, by other reactions, the coupling enzyme is prevented from reassociation with the electron transport particles by 2,4-dinitrophenol.

Phosphorylating particles washed with 2,4-dinitrophenol lose coupling enzyme and with it the ability to form the energy-rich intermediate and to couple phosphorylation to electron transport.

Phosphorylating particles containing bound coupling enzyme are able to form the high energy intermediate even in the presence of 2,4-dinitrophenol. However, after this first cycle of formation, no more can be formed because 2,4-dinitrophenol prevents the reassociation of coupling enzyme with the electron transport particles.

2,4-Dinitrophenol stimulates respiration in the bacterial phosphorylating particles.

Table of Contents

Chapter 1 The Diagram Method: States
1.1 Introduction
1.2 Diagrams for Steady-State (and Equilibrium) Systems
1.3 Directional Diagrams and the Steady-State Populations of States
1.4 Simple Examples of the Use of Directional Diagrams
1.5 Transition Fluxes and Probabilities of States
Chapter 2 The Diagram Method: Cycles
2.1 Cycles and Cycle Fluxes
2.2 One-Way Cycle Fluxes Kinetics at the Cycle Level
2.3 Further Examples of Cycles and of Flux Diagrams
2.4 Calculation of State Probabilities from Cycle Fluxes
Chapter 3 Fluxes and Forces
3.1 Example: Membrane Transport of Two Ligands
3.2 Substrate-Product Rate Constant Relations
3.3 Example: Active Transport of Na+ and K +
3.4 Reciprocal Relations and Irreversible Thermodynamics
Chapter 4 Free Energy Levels of Macromolecular States
4.1 Free Energy Levels of the States
4.2 Single-Cycle Examples
4.3 Simple Multicycle Example with Two Forces
4.4 Enzyme-Substrate Modified by Ligand
Chapter 5 Muscle Contraction
5.1 General Principles
5.2 The Kinetic Formalism
5.3 Entropy Production and Directional Properties
5.4 Current Status of Muscle Models
5.5 Free Energy Transfer in Muscle Contraction
Chapter 6 Stochastics and Fluctuations at Cycle and State Levels
6.1 Stochastics of Cycle Completions
6.2 Some Further Stochastic Considerations in Muscle Contraction
6.3 State Stochastics: A Two-State System
6.4 State Stochastics: Arbitrary Diagram
Chapter 7 Interacting Subsystems and Multienzyme Complexes
7.1 Example: Two-Enzyme Complex
7.2 Example: Three-Enzyme Complex
7.3 Example: Two Interacting Enzymes
7.4 Oxidative Phosphorylation
Appendix 1 "Reduction" of a Diagram
Appendix 2 Diagram Solution for the Nix Flux Diagrams
Appendix 3 Charged Ligand and Membrane Potential
Appendix 4 Some Properties of Single-Cycle Diagrams
Appendix 5 Light Absorbing (and Emitting) Systems
Appendix 6 Basic and Gross Free Energy Levels in a Simple Special Case

Structure and Function of the Mitochondria

The mitochondria (Figure 8) are where the oxidative-phosphorylation reactions occur. The mitochondria are specialized, rod-shaped (oval-shaped) cellular compartments (organelles) with dimensions of approximately 2 µm by 0.5 µm.(Recall that the protein Ferritin has a diameter of about 80 Å, or 8 x 10 -3 µm.) Mitochondria are present in virtually every cell of the body. They contain the enzymes required for the citric-acid cycle (the last steps in the breakdown of glucose), oxidative phosphorylation, and the oxidation of fatty acids.

Figure 8

This is a schematic diagram showing the membranes of the mitochondrion. The purple shapes on the inner membrane represent proteins, which are described in the section below. An enlargement of the boxed portion of the inner membrane in this figure is shown in Figure 8, below.

The mitochondrial membranes are crucial for this organelle's role in oxidative phosphorylation. As shown in Figure 8, mitochondria have two membranes, an inner and an outer membrane. The outer membrane is permeable to most small molecules and ions, because it contains large protein channels called porins . The inner membrane is impermeable to most ions and polar molecules. The inner membrane is the site of oxidative phosphorylation . Although the membrane is mostly impermeable, it contains special H + (proton) channels and pumps that enable the coupling of the redox reaction involving NADH and O2 (Equations 9-10) to the phosphorylation reaction of ADP (Equation 8), as described below ("Oxidation-Reduction Reactions and Proton Pumping in Oxidative Phosphorylation"). (Recall the discussion of protein channels in the " Maintaining the Body's Chemistry: Dialysis in the Kidneys " Tutorial.)

As shown in Figure 8, inside the inner membrane is a space known as the matrix the space between the two membranes is known as the intermembrane space . The matrix side of the inner membrane has a negative electrical charge relative to the intermembrane space due to an H + gradient set up by the redox reaction (Equations 9 and 10). This charge difference is used to provide free energy (G) for the phosphorylation reaction (Equation 8).

4 Experimental Section

Cell Culture

Cell lines (293T, MCF7, HCT116 (p53 +/+ ), and HepG2) were cultured in high glucose (25 × 10 −3 m ) Dulbecco's modified Eagle's medium (Thermo Fisher Scientific) with 10% v/v fetal bovine serum (BI, Biological Industries), 4 × 10 −3 m L-glutamine, 1% v/v penicillin-streptomycin (Gibco), and 1 × 10 −3 m pyruvate (Gibco) and maintained at 37 °C in a humidified 5% CO2-containing atmosphere. Cell line authenticity was verified by short tandem repeat analysis.

Microarray Analysis

Transcriptomic analyses were obtained on a contract basis BGI (The Beijing Genomics Institute) Company, Shenzhen, China with data analyses performed according to publically available instructions ( ) .

RNA Inference and Transfections

Gene knockdown experiments were conducted by lentiviral-mediated transduction with short hairpin RNAs (shRNAs). Lentiviral particles were generated by transfection of 293T cells with PLKO.1 vectors containing specific shRNAs (Table S2, Supporting Information) along with pREV, pGag, pVSVG at the ratio of 2:2:2:1 in Opti-MEM medium (Gibco) for 48 h. Supernatants were filtered with 0.45 µm filter before cells infection, added to target cells for 24 h before selection with 5 µg mL −1 puromycin. Alternatively, transfections were performed with the indicated plasmids (Table S3, Supporting Information) using the Lipofectamine-2000 reagent (Invitrogen) according the manufacturer's instructions.

Glycolysis and Mitochondrial Stress Tests

Assays were performed using the Seahorse XFe96 analyzer (Agilent) according to the manufacturer's instructions. Cells were seeded at 1×10 4 cells per well in 96-well XF cell culture micro-plates for 24 h before performing glycolysis stress tests at 37 °C in XF base medium (2 × 10 −3 m glutamine, pH 7.4) with sequential additions of glucose (10 × 10 −3 m ), oligomycin (1 × 10 −6 m ), and 2-DG (5 × 10 −3 m ). Alternatively, mitochondria stress tests were performed in XF base medium (1 × 10 −3 m pyruvate, 2 × 10 −3 m glutamine, 10 × 10 −3 m glucose, pH 7.4) with sequential additions of oligomycin (1.0 × 10 −6 m ), FCCP (0.25 × 10 −6 m ), and Rot/AA (0.5 × 10 −6 m ). Data were analyzed by the Seahorse XF Glycolysis Stress Test and Mitochondria Stress Test Report Generator packages, respectively.

Subcellular Fractionation

For cytosol/nuclear fractions, cell suspensions were incubated with hypotonic buffer (25 × 10 −3 m Tris-HCl pH 7.4, 1 × 10 −3 m MgCl2, 5 × 10 −3 m KCl) on ice for 5 min before adding an equal volume of hypotonic buffer containing 1% NP-40 for another 5 min. Homogenates prepared by pipetting were then centrifuged at 5000 x g for 10 min at 4 °C, and the supernatant collected as the cytosol fraction. Pellets were rinsed twice with hypotonic buffer and re-suspended in nuclear resuspension buffer (20 × 10 −3 m 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid pH 7.9, 400 × 10 −3 m NaCl, 1 × 10 −3 m ethylenediaminetetraacetic acid, 1 × 10 −3 m ethylene glycol-bis(β-aminoethyl ether)-N,N,N′,N′-tetraacetic acid, 1 × 10 −3 m dithiothreitol, 1 × 10 −3 m phenylmethylsulfonyl fluoride). After incubation on ice for 30 min, the samples were centrifuged at 12 000 x g for 10 min at 4 °C and the supernatant collected as the nuclear fraction. Similarly, cytosolic/mitochondrial were performed with the mitochondria isolation kit (Sigma) but with stepwise centrifugation of homogenates at 600 x g for 10 min, and then at 11 000 x g for 10 min to derive supernatant (cytosol fraction) and pellet fractions, the latter re-suspended with mitochondria storage buffer as the mitochondrial fraction.

Western Blotting and Immunoprecipitation

Cell lysates were prepared with radio immunoprecipitation assay buffer containing protease inhibitors (Beyotime) and clarified by centrifugation at 10 000 x g for 15 min at 4 °C. Equal amounts as determined using the Bio-Rad RC/DC protein assay were electrophoresed by sodium dodecyl sulfate-polyacrylamide gel electrophoresis and transferred to nitrocellulose membranes. Membranes were blocked with 4% skim milk and incubated with primary antibodies overnight at 4 °C, decorated with horseradish peroxidase-conjugated secondary antibodies (Table S5, Supporting Information) with detection using chemiluminescence (Advansta). Alternatively, for immunoprecipitations, cell lysates prepared with IP buffer (0.5% NP-40, 20 × 10 −3 m Tris pH 7.4, 150 × 10 −3 m NaCl, 1.5 × 10 −3 m MgCl2 and protease inhibitor cocktail (Solarbio)) were incubated with primary antibodies adsorbed to protein A/G-sepharose (Invitrogen) beads for 4 h, washed five times with IP buffer. Antibody sources/dilutions are shown in Table S4 with uncropped immunoblot scans provided in the Supporting Information.

Nile Red Staining

Cells grown on glass coverslips were rinsed with warmed phosphate-buffered saline (PBS) and fixed in 4% formaldehyde before staining with a solution containing Nile Red (8 µg mL −1 ) and Hoechst 33342 (0.1 µg mL −1 ) for 10 min at room temperature. After mounting the coverslips, images were collected using a ZEISS LSM 700 confocal microscope. For quantitation, cells were harvested and cell suspensions were incubated with Nile Red solution, washed with PBS, and transferred to 96-well plate before measuring the absorbance at OD 530 nm.

Immunofluorescence Staining

Cells grown on glass coverslips were fixed in 4% formaldehyde for 15 min at room temperature before permeabilization (0.2% Triton X-100) for 10 min at room temperature. Samples were blocked (1% bovine serum albumin) for 60 min at room temperature and washed using PBS with 0.05% Tween-20 (PBST) before the addition of primary antibodies in PBST overnight at 4 °C. Samples were washed and bound antibodies decorated with appropriate fluorochrome-conjugated secondary antibodies diluted in PBST for 1 h at room temperature. Samples were counterstained with Hoechst 33342, mounted in anti-fade mounting medium, and images were collected using a ZEISS LSM 700 confocal microscope.


CRISPR/Cas9-mediated DDIT3, TIGAR, and ATF4gene-editing vectors were constructed by annealing gRNA oligonucleotide pairs (Table S3, Supporting Information) and subcloning into lentiCRISPRv2 (one vector system) according to the Zhang laboratory protocol. Lentiviral particles produced as described above using a 1:2:2 mixture of plasmids (Pmd2.g, PSPAX2 and lentiCRISPRv2) were used to transduce target cells. After selection with 5 µg mL −1 puromycin, stably infected cells were plated in 96-well plates and single cell clones were screened by Western blot and gDNA sequencing to obtain DDIT3, TIGAR, and ATF4 knockout cells.


RNA extraction using the TRIzol Reagent (Invitrogen) was performed on cells grown in 12-well plates. cDNA was prepared from total RNA using the PrimeScriptTM RT reagent kit (TaKaRa) according to the manufacturer's instructions. Quantitative RT-PCR analyses were performed using the specified primers (Table S4, Supporting Information) with the One-Step PrimeScript RT-PCR kit (TaKaRa). Relative expression values were calculated using the comparative Ct method normalized against the beta-microglobulin housekeeping gene.

Metabolite Assays

Cellular ATP, ROS, extracellular lactate levels, mitochondrial ROS, and glucose uptake were measured using the ATP assay Kit (Biovision), ROS assay kit (Beyotime), Lactate production assay kit (Biovision), Mitochondrial ROS Detection Assay kit (Cayman Chemical), and 2-NBDG glucose uptake assay kit (Abcam), respectively, according to the manufacturer's instructions. Fru-2,6-BP levels in HepG2 DDIT3-WT and -KO cells treated with or without glutamine were determined by Quantitative Metabolomics based on LC-MS from BGI (The Beijing Genomics Institute) company, Shenzhen, China.

Luciferase Reporter Assays

Cells seeded in 24-well plates were co-transfected with the indicated pGL3-based reporter plasmids (Table S3, Supporting Information) along with Renilla luciferase. After 24 h, the results were assessed using the Dual-Luciferase Reporter Assay System (Promega) with firefly luciferase values corrected against Renilla measurements according to the manufacturer's instructions.

Chromatin Immunoprecipitation

ChIP assays were performed with the Millipore ChIP kit according to the manufacturer's instructions. Bound DNA fragments were subjected to semi-quantitative RT-PCR using the specified primers (Table S4, Supporting Information).

DDIT3 Knockout and Animal Experiments

A general chromosome-engineering technique was used to target the imprinted DDIT3 box gene cluster by deleting an intronic fragment between exons 3 and 4 using CRISPR-CAS9 (Figure 5A, upper). After derivation, DDIT3+/− C57/BL mice were maintained under SPF conditions under contract with Cyagen Biology Company. Following genotyping, backcrossed mice were divided into DDIT3+/+ and DDIT3−/− groups (Table S4, Supporting Information and Figure 5A, upper) with half of each group, then fed a standard diet or glutamine deprivation diet (Medicience Ltd.). Following mating, 3 weeks old progenies were weaned and these mice were maintained for a further 9 weeks on the specified diets.

Xenograft Model

BALB/c nude mice (5 weeks old, male) were obtained from Shanghai SLAC Laboratory Animal Co. Ltd. After 3 days acclimatization, mice were injected subcutaneously (s.c.) with 5.0 × 10 6 WT and DDIT3-KO HepG2 cells on alternate sides of each mouse. Mice were randomized into normal diet and glutamine-free diet groups and maintained for 4 weeks before the animals were humanely sacrificed and the tumors were excised and weighed. Studies were conducted with the approval from the Animal Research Ethics Committee of University of Science and Technology of China.


Mice were fasted overnight for 12 h, and fasting blood glucose levels were determined by glucometer using tail vein blood. For GTT, mice were then injected i.p. with 10 uL of glucose solution (200 mg mL −1 ) per gram of body weight and blood glucose was measured at 15, 30, 60, 90, and 120 min post injection. For ITT, fasted mice were injected i.p. with insulin at 0.75 U kg −1 body weight using a stock insulin solution of 0.1 U mL −1 in 0.9% NaCl. Blood glucose was measured at 15, 30, 45, and 60 min after insulin injection. Studies were conducted with approval from the Animal Research Ethics Committee of University of Science and Technology of China.

Statistical Analysis

Data were assumed to be normally distributed and continuous variables expressed as mean ± SD. All analyses were performed by two-tailed Student's t-test using GraphPad Prism 8 with significance defined as p ≤ 0.05. Reproducibility and the number of replicates used are defined in the corresponding figure legends.

Photobiomodulation and Oxidative Stress: 980 nm Diode Laser Light Regulates Mitochondrial Activity and Reactive Oxygen Species Production

Photobiomodulation with 808 nm laser light electively stimulates Complexes III and IV of the mitochondrial respiratory chain, while Complexes I and II are not affected. At the wavelength of 1064 nm, Complexes I, III, and IV are excited, while Complex II and some mitochondrial matrix enzymes seem to be not receptive to photons at that wavelength. Complex IV was also activated by 633 nm. The mechanism of action of wavelengths in the range 900–1000 nm on mitochondria is less understood or not described. Oxidative stress from reactive oxygen species (ROS) generated by mitochondrial activity is an inescapable consequence of aerobic metabolism. The antioxidant enzyme system for ROS scavenging can keep them under control. However, alterations in mitochondrial activity can cause an increment of ROS production. ROS and ATP can play a role in cell death, cell proliferation, and cell cycle arrest. In our work, bovine liver isolated mitochondria were irradiated for 60 sec, in continuous wave mode with 980 nm and powers from 0.1 to 1.4 W (0.1 W increment at every step) to generate energies from 6 to 84 J, fluences from 7.7 to 107.7 J/cm 2 , power densities from 0.13 to 1.79 W/cm 2 , and spot size 0.78 cm 2 . The control was equal to 0 W. The activity of the mitochondria’s complexes, Krebs cycle enzymes, ATP production, oxygen consumption, generation of ROS, and oxidative stress were detected. Lower powers (0.1–0.2 W) showed an inhibitory effect those that were intermediate (0.3–0.7 W) did not display an effect, and the higher powers (0.8–1.1 W) induced an increment of ATP synthesis. Increasing the power (1.2–1.4 W) recovered the ATP production to the control level. The interaction occurred on Complexes III and IV, as well as ATP production and oxygen consumption. Results showed that 0.1 W uncoupled the respiratory chain and induced higher oxidative stress and drastic inhibition of ATP production. Conversely, 0.8 W kept mitochondria coupled and induced an increase of ATP production by increments of Complex III and IV activities. An augmentation of oxidative stress was also observed, probably as a consequence of the increased oxygen consumption and mitochondrial isolation experimental conditions. No effect was observed using 0.5 W, and no effect was observed on the enzymes of the Krebs cycle.

1. Introduction

Oxidative stress from reactive oxygen species (ROS) generated by mitochondrial activity is an inescapable consequence of aerobic metabolism. The vital role of mitochondria in tissue energy metabolism is to convert the products of biotransformation to CO2 and water. For this to happen, enzymes of the electron transport chain (ETC), such as NADH-dehydrogenase (Complex I), succinate dehydrogenase (Complex II), cytochrome bc1 (Complex III), and cytochrome c oxidase (Complex IV), which pump protons from the matrix to the intermembrane space, are necessary, determining a proton gradient and the synthesis of adenosine triphosphate (ATP) by the enzyme Fo-F1 ATP synthase (Complex V) [1]. When the ETC becomes saturated with electrons, it can pass to O2 by Complexes I and III and generate superoxide anions [1]. The generation of superoxide anions can also be correlated to the univalent reduction of oxygen by mitochondrial semiquinones [2]. In this way, more than 90% of ATP produced and 85–90% of oxygen breathed by aerobic cellular tissues are derived from mitochondria [2, 3], and under physiological conditions, between 0.2 and 2% of this activity is converted into ROS [4]. Mitochondria are also equipped with an antioxidant enzymatic system (AES) for ROS scavenging, which keeps them under control without causing damage to the cell [5]. However, alterations in the transfer of electrons and/or AES can cause electrons to accumulate on the ETC complexes and enhance ROS production. From a mitochondrial point of view, ROS and ATP can regulate cell homeostasis and play a role in cell death, cell proliferation, and cell cycle arrest.

Karu and Afanasyeva and Passarella and Karu first described the ability of red and near-infrared (NIR) light to interact with mitochondria [6, 7]. Indeed, light-cell interactions in nonplant cells, such as nonphotosynthesising prokaryotic and protozoan cells and animal cells, have been described [8]. Basically, when a photon interacts with a specific photoacceptor, its energy is absorbed to generate high-energy electrons. The excited molecule can lose its energetic status in the form of heat or fluorescence emission, or the absorbed light energy can be transferred to a photosystem molecule as an excited electron or state. In this way, the photosystem converts the photon’s energy into chemical energy, thanks to the tricky process of electron transport and a proton gradient, ending with the conversion of ADP to ATP [9]. In plants, this process occurs in the chloroplast, whereas it occurs in photoacceptors in bacteria, and the conversion of ADP takes place in the inner part of the cell membrane. In other eukaryotic cells, electron transport occurs in the mitochondrial respiratory chain [7, 8]. Complex IV was shown to be activated in vitro by a red laser (632.8 nm, 15 mW, CW, dose

J/m 2 , 10 s) [10]. Particularly, according to metal-ligand systems and absorption spectra, such as 450, 620–680, and 760–895 nm, characteristically different peaks may be related to oxidized or reduced copper into cytochrome c oxidase [11]. In previous papers, we showed that 808 nm electively stimulates Complex IV and that Complex III was excited poorly, while Complexes I and II were not affected [12]. In addition, by increasing the wavelength to 1064 nm, the photon and mitochondrial complex interaction changes, and Complexes I, III, and IV are affected, while the extrinsic mitochondrial membrane Complex II and mitochondrial matrix enzymes seem to be not receptive to photons at this wavelength [13].

The mechanism of action of wavelengths 900–1000 nm is, however, less well understood, and, in particular, interactions with mitochondrial complexes have not been described. Conversely, Wang et al. [14] concluded that at the parameters tested, 980 nm affected temperature-gated calcium ion channels but not mitochondrial cytochrome c oxidase when compared to 810 nm. Water could then be a candidate as a chromophore for longer wavelengths of NIR, based on its absorption spectrum.

Starting with these premises, the purpose of this study was to evaluate the interaction between a 980 nm diode laser light and mitochondrial activity. The investigators hypothesized that according to previous in vitro literature on 1064 nm wavelength, the 980 nm wavelength could have a modulatory effect on respiratory chain activities. The specific aims of the study were to determine the effectiveness of a 980 nm diode laser, irradiated at the powers from 0 to 1.4 W (0.1 W increment at every step), for 60 sec in continuous wave mode (CW), spot size 0.78 cm 2 , on bovine liver isolated mitochondria. The activity of the mitochondria’s complexes, ATP production, oxygen consumption, and production of ROS were measured. The results were discussed and compared to our previous data and literature.

2. Materials and Methods

2.1. Laser Features, Parameters, and Method of Irradiation

The experimental design is shown in supplementary Figure 1. Mitochondria were isolated from bovine liver and irradiated as described, at room-air temperature, or partially immersed in water to prevent macroscopic thermal effects.

The laser device utilised in the study was the Wiser wireless diode laser by Doctor Smile–LAMBDA Spa (Vicenza, Italy). The 980 nm diode laser light was irradiated by the AB 2799 hand-piece (Doctor Smile-LAMBDA Spa, Vicenza, Italy). The AB 2799 hand-piece is a novel hand-piece with a flat-top beam profile, which was set following the manufacturer’s specifications, to allow delivery of homogenous irradiation over the surface area with the same irradiation spot area (0.78 cm 2 ) and power from contact, extending to many centimetres (

100 cm) of distance from the target by comparison, the standard hand-piece would deliver a Gaussian profile of irradiation and is accompanied by beam divergence over distance [12, 15]. To make sure that the laser delivery power was constant during the irradiation mode and suitable for our experimental setup, the PM160T-HP power meter (ThorLabs, Germany) was utilised according to Hanna et al. [15].

To control the thermal increase on the irradiated samples, a thermal camera FLIR ONE Pro-iOS (FLIR Systems, Inc. designs, Portland, USA.) (dynamic range: -20°C/+400°C resolution 0.1°C) was used during irradiation. The temperature measures were collected before and after irradiation, which was performed at room-air temperature (25°C) or with a tube sample partially immersed in 300 ml of water (25°C) (supplementary Figure 1). The temperature of the sample of mitochondria irradiated at the room-air temperature was also measured after the addition of reagents for biochemical evaluation (temperature of the reagents 25°C).

For our experimental purpose, the Wiser wireless diode laser was set to irradiate for 60 sec in continuous wave mode with power from 0.1 to 1.4 W (0.1 W increment at every step), which generated energies from 6 to 84 J, fluences from 7.7 to 107.7 J/cm 2 , and power densities from 0.13 to 1.79 W/cm 2 (please see Table 1 for a more descriptive representation of the parameters).

Irradiations were performed with both the sample and the hand-piece fixed to a stand at a distance of 3.5 mm. Irradiations performed with the laser device kept off were considered a control.

2.2. Reagents

Salt, substrates, and all other chemicals (of analytical grade) were purchased from Sigma–Aldrich (St. Louis, MO, USA). Protein Molecular Weight (MW) markers were from Bio-Rad (Hercules, CA, USA). Ultrapure water (Milli-Q Millipore, Billerica, MA, USA) was used throughout. Safety precautions were taken for chemical hazards in carrying out the experiments. Ampicillin (25 μg/ml) was used in all the solutions, and sterile experimental conditions were employed where appropriate.

2.3. Mitochondrial Enriched-Fraction Isolation

Bovine liver from 2 female and 2 male cattle of less than 1 year of age were acquired from the slaughterhouse of Ceva, Torino, Italy. The cattle were bred for human consumption, following the directives of the Italian Ministry of Agricultural, Food and Forestry Policies. Samples were taken and processed immediately after slaughter, following all safety rules. Since the animals were not bred nor sacrificed at the University of Genoa, it was not necessary to request any ethical committee approval.

To obtain a mitochondrial enriched fraction, the bovine liver was washed in PBS and homogenized in a buffer containing 0.25 M sucrose, 0.15 M KCl, 10 mM Tris-HCl pH 7.4, and 1 mM EDTA. The homogenate was centrifuged at 800 × g for 10 min. The supernatant was filtered and centrifuged at 12000 × g for 15 min. Pellet was resuspended in another buffer containing 0.25 M sucrose, 75 mM mannitol, 10 mM Tris-HCl pH 7.4, and 1 mM EDTA. The final supernatant was centrifuged at 12000 × g for 15 min, and the mitochondrial pellet resuspended in the same buffer [16].

2.4. Oxygen Consumption Measurements

The oxygen consumption rate (OCR) was assayed in a thermostatically controlled oxygraph apparatus equipped with an amperometric electrode (Unisense–Microrespiration, Unisense A/S, Denmark) on mitochondrial enriched fraction treated or not with the 980 nm laser. For each sample, 50 μg of total protein was used. The samples were incubated in the respiration buffer composed of 120 mM KCl, 2 mM MgCl2, 1 mM KH2PO4, 50 mM Tris-HCl pH 7.4, and 25 μg/ml ampicillin. As respiratory substrates, 5 mM pyruvate+2.5 mM malate were added to the respiration buffer [12].

2.5. Evaluation of Mitochondrial ATP Synthesis

To evaluate the ATP production through the Fo-F1 ATP synthase (ATP synthase), mitochondrial enriched fraction treated or not with the 980 nm laser was dissolved in a solution containing 100 mM Tris-HCl pH 7.4, 100 mM KCl, 1 mM EGTA, 2.5 mM EDTA, 5 mM MgCl2, 0.2 mM di(adenosine-5

) penta-phosphate, 0.6 mM ouabain, ampicillin (25 μg/ml), 5 mM KH2PO4, and 5 mM pyruvate+2.5 mM malate, used as respiratory substrates. ATP synthesis started after the addition of 0.1 mM ADP and was monitored for 2 minutes, in a luminometer (Glomax 20/20, Promega) by the luciferin/luciferase chemiluminescent method. An ATP standard solution between 10 -9 and 10 -7 M was used for calibration [12].

2.6. Calculation of Mitochondrial Efficiency

To evaluate the energy production efficiency by the mitochondrial enriched-fraction, the ratio between the produced ATP and consumed atomic oxygen (P/O) was calculated. When the oxygen consumption is perfectly associated to the ATP synthesis, the P/O ratio is about 2.5 in the presence of pyruvate+malate as respiratory substrates [17]. Conversely, in the uncoupled status, this value decreases correlating to the grade of the oxidative phosphorylation inefficiency.

2.7. Assay of Respiratory Complex Activity

The activity of the four respiratory complexes was assayed on 50 μg of total mitochondrial enriched-fraction protein treated or not with the 980 nm laser [13].

Complex I (NADH-ubiquinone oxidoreductase) was assayed following the reduction of ferricyanide at 420 nm the assay solution was composed by 100 mM Tris-HCl pH 7.4, 0.6 mM NADH, 0.8 mM ferricyanide, 50 mM KCl, 5 mM MgCl2, 1 mM EGTA, and 50 μM antimycin A.

Complex II (succinic dehydrogenase) activity was measured following the reduction of ferricyanide at 420 nm the assay solution was composed by 100 mM Tris-HCl pH 7.4, 20 mM succinate, 0.8 mM ferricyanide, 50 mM KCl, 5 mM MgCl2, 1 mM EGTA, 20 μM rotenone, and 50 μM antimycin A.

Complex III (cytochrome c reductase) activity was assayed following the reduction of oxidized cytochrome c (Cyt c) at 550 nm the assay solution was composed by 100 mM Tris-HCl pH 7.4 and 0.03% oxidized cytochrome c. The assay started with the addition of 0.7 mM NADH.

Complex IV (cytochrome c oxidase) was assayed following the oxidation of ascorbate-reduced Cyt c at 550 nm the assay solution was composed by 100 mM Tris-HCl pH 7.4, 50 mM, and 0.03% reduced cytochrome c.

2.8. Isocitric Dehydrogenase and Malate Dehydrogenase Assay

Isocitric dehydrogenase (IDH EC: and malate dehydrogenase (MDH EC have been assayed as Krebs cycle markers. For each assay, 50 μg of total mitochondrial enriched-fraction protein treated or not with the 980 nm laser was used.

For IDH, the assay solution contained 100 mM Imidazole buffer pH 8, 3.5 mM MgCl2, 0.41 mM NADP, and 0.55 mM isocitric acid [18].

For MDH, the assay solution is formed by 100 mM Tris-HCl pH 7.5, 0.5 mM oxaloacetic acid, and 0.2 mM NADH [13].

2.9. Superoxide Anion Assay

The production superoxide anion was assayed as the difference between total and SOD-inhibitable cytochrome c reduction. 50 μg of total mitochondrial enriched-fraction protein treated or not with the 980 nm laser was added to 100 μM cytochrome c and 300 U SOD, if present, and the changes in absorbance of reduced cytochrome c was measured spectrophotometrically, at 550 nm [19].

2.10. Evaluation of Lipid Peroxidation

To assess lipid peroxidation in mitochondrial enriched-fraction protein treated or not with the 980 nm laser, the malondialdehyde (MDA) level was evaluated, using the thiobarbituric acid reactive substances (TBARS). The TBARS solution contained 15% trichloroacetic acid (TCA) in 0.25 N HCl and 26 mM 2-thiobarbituric acid. To evaluate the basal concentration of MDA, 600 μl of TBARS solution was added to 50 μg of total protein dissolved in 300 μl of Milli-Q water, after 10 minutes of laser exposure. The mix will be incubated for 40 min at 100°C, then centrifuged at 14000 rpm for 2 min, and the supernatant was analysed spectrophotometrically, at 532 nm [20].

2.11. Statistical Analysis

Statistical analyses were performed with GraphPad Prism software version 7 (GraphPad Software). All parameters were tested by one-way ANOVA followed by Bonferroni test. Data are expressed as

(SD) from 3 to 5 independent determinations performed in duplicate. In the figures, SD is shown as error bars. An error probability with

was selected as significant.

3. Results

3.1. Temperature Evaluation and Its Effect on the Results

As shown in supplementary Figure 2, when irradiated at the room-air temperature (blue line), the temperature of the irradiated mitochondrial samples progressively increased (from 25°C to 38°C) by increasing the power and energy irradiated (yellow line). However, when the reagents for the biochemical analysis were added, the final temperature recovered to the initial value of 25°C (green line). Conversely, the irradiation with the mitochondrial samples partially immersed in water allowed them to keep the temperature quite constant at 25°C (red line). Comparison of the results obtained after irradiation at the room-air temperature or in water did not show statistically significant differences.

3.2. 980 nm Laser Affects the OxPhos Activity

To evaluate whether the 980 nm laser exerted effects on the mitochondrial energy production, as already observed for the 1064 nm Nd:YAG laser [13], ATP synthesis through ATP synthase was evaluated applying a power of 0–1.4 W. As reported in Figure 1, the 980 nm laser induced a marked decrease in the ATP production when exposed to 0.1 and 0.2 W. Conversely, the power between 0.8 and 1.1 W determined an enhancement of energy synthesis, while the intermediate values (0.3–0.7 W) did not display effects. Based on these data, we chose to evaluate the energy metabolism of mitochondria exposed to the 980 nm laser, using the following power: 0, 0.1, 0.5, and 0.8 W.

and indicate a significant difference for and 0.0001, respectively, between the nontreated sample (0 W) and the irradiated samples.

Data reported in Figure 2 show that the oxygen consumption rate (OCR) (panel b) followed the same trend observed for ATP synthesis (panel a) when mitochondria were stimulated with pyruvate and malate as respiratory substrates. Moreover, evaluating the efficiency of the mitochondrial energy production, we observed that the lower power determined not only a reduction of activity but also the decrement of the P/O ratio, which was around 0.7, instead of the regular value of 2.5 [17]. This suggested that 0.1 W induced an uncoupling status between the ATP production and respiration. In contrast, higher power did not affect mitochondrial efficiency (panel c).

3.3. Modulation of OxPhos Activity Induced by the 980 nm Laser Depended on Changes in the Activity of Complexes III and IV

To understand the targets of the 980 nm laser on the electron transport chain, the activities of the four respiratory complexes were evaluated. As reported in Figure 3, only Complexes III and IV were modulated by the laser (panels c and d), while Complexes I and II did not show a difference between untreated and treated samples (panels a and b). In particular, the lower power (0.1 W) determined a marked reduction of the activities of Complexes III and IV. Conversely, the higher power (0.8 W) induced an increment of the complexes’ activity. No effects were observed on any complex with the 0.5 W treatment.

3.4. The 980 nm Laser Did Not Alter the Activity of Isocitric Dehydrogenase and Malate Dehydrogenase

To evaluate whether the modulation of OxPhos activity could depend on the alteration of the Krebs cycles, the upstream pathway of the electron transport chain and activity of isocitric dehydrogenase (IDH) and malate dehydrogenase (MDH) were evaluated. Figure 4 shows that the three considered powers (0.1, 0.5, and 0.8 W) did not exert effects on the activity of IDH (panel a) and MDH (panel b).

3.5. Modulation of OxPhos Activity by the 980 nm Laser Determined an Increment of Superoxide Production and Lipid Peroxidation

Mitochondria are considered one of the principal sources of oxidative stress production, which increases when the ATP production and oxygen consumption are uncoupled [21]. Therefore, to evaluate whether OxPhos modulation due to the laser treatment could be associated with an increment of the oxidative stress production and accumulation of oxidative damage, the superoxide level and the malondialdehyde concentration were evaluated.

Figure 5(a) shows that both 0.1 and 0.8 W determined an increment of the superoxide production, although more evident was the effect of the lower power. In contrast, the increment observed after the treatment with 0.8 W may be associated with the increment of OxPhos enhancement.

The same trend was observed by evaluating the MDA intracellular concentration (Figure 5(b)), as a marker of lipid peroxidation, suggesting that the increment of oxidative stress was associated with oxidative damage of the mitochondrial membrane. No oxidative stress and lipid peroxidation were observed when the sample was treated with 0.5 W.

4. Discussion

The interaction between light and cells is well-known in vegetable cells. Red and NIR light is, however, able to modulate nonplant cell energetic metabolism, and this happens because of a parallel and convergent evolution of both chloroplasts and mitochondria from ancestral bacteria [8], known as the theory of endosymbiont models. The relative medical subject is known as photobiomodulation and can lead to improvement of pathological conditions [22]. Our data describe for the first time the ability of 980 nm light to interact with the respiratory chain of bovine liver mitochondria. Under our experimental conditions, a diode laser light was used to irradiate cells for 60 seconds, and three different effects were observed on ATP production. Lower powers (0.1–0.2 W) showed a sizable, inhibitory effect intermediate powers (0.3–0.7 W) had no effect and the higher powers (0.8–1.1 W) induced a strong and stable increment of ATP synthesis, which reached a peak at 1.1 W. Increasing the power (1.2–1.4 W) recovered the ATP production to the control level.

Because of the isolated experimental conditions of the mitochondria, we do not know what the consequences of cell homeostasis would be. However, the data suggest a reconsideration of the parameters able to interact with cell photoacceptors. Essentially, the statement that higher energies and powers have undoubtedly damaging effects with respect to the lower, which are “curative” [23, 24], should be reassessed. Actually, our data pointed out that a hormetic behaviour can certainly occur, but into narrow windows of positive effect/no effect/negative effect more than a watershed upper at or lower of. Additionally, the effectiveness of 980 nm joined to previous works resulted in a laser light and mitochondrial interaction of red and NIR wavelengths [7, 12, 13, 25], pointing to photobiomodulation on a wider spectrum of wavelengths than photosynthesis. Indeed, all life forms need energy for existence. However, only photosynthetic organisms developed light-energy conversion, and by evolution, ranges of light intensities have been selected for better survival in the biosphere [26]. The animal cell did not choose sunlight as a source of energy for its metabolism, and most cell types have no interaction with light from this point of view, only cells interacting with light evolved the ability to use specific light stimuli, for instance, for vitamin D production and the vision. Interaction between mitochondria and 980 nm light prevalently occurs with Complex IV, as shown in our results, despite the fact that Complex III activity is also deeply stimulated. Complexes I and II were not energized. Because of similar experimental conditions with bovine isolated mitochondria from liver tissue, the data showed here and our previous works on mitochondrial complexes can be compared.

Complex IV was similarly stimulated moving from 808 to 980 and 1064 nm. Complex III was also excited by the same wavelengths, but the intensity progressively increased when compared to the other complexes. Complex I was conversely stimulated only by 1064 nm, while Complex II never changed. Besides, Complex IV was shown to be activated in vitro by 633 nm [10].

The respiratory chain is situated in the inner membrane of mitochondria and comprises four multimeric protein complexes: Complex I containing eight Fe-S clusters implicated the electrons’ transfer from reduced flavin mononucleotide (FMNH2) to ubiquinone Complex II has a heme b prosthetic group in its anchor domain, essential for the structural integrity and its function Complex III contains cytochrome b subunit and two heme moieties, a cytochrome c1 subunit with one heme, and a Rieske protein subunit (UQCRFS1) with a (2Fe-2S) cluster and Complex IV that modulates the final step in the ETC, by catalysing the reduction of O2 to water. It presents two heme moieties and two Cu centres, participating in the electron transfer process [27].

As previously discussed in our paper on the characterization between 1064 nm light and mitochondrial complexes, explaining the change as a mundane variation of light–“metal” uptake is an implausible choice. This model was not able to explain the unaffected behaviour of Complex I and, in the second analysis, since both iron and copper coefficients of absorption did not significantly increase or change, respectively, increasing wavelength up to 1064 nm [28, 29].

Conversely, water had a weak absorption in the visible spectrum but it increased by moving toward the 1064 nm. Additionally, despite lipids in the near-infrared wavelength region, there were two high peaks at 1210 and 1720 nm and a third lesser peak in the range of 900–1000 nm [30]. Thus, the light-water interaction at the nanoscale level could modify cell membrane features and the activities of the intramembrane complexes. In contrast, OxPhos machinery function depends on the integrity of the inner mitochondrial membrane, since the protons transported by the respiratory complexes should not be free to pass the membrane. Protons must be returned to the mitochondrial matrix only through the Fo moiety of the ATP synthase, to produce energy efficiently [31, 32]. Thus, the alteration of lipids constituting the inner mitochondrial membrane could cause a change in its shape and energy metabolism [33]. In particular, the laser could affect the structure of cardiolipin, a phospholipid expressed only in the active respiring membranes [34], playing a pivotal role in the structure of the respiratory complexes [35] and proton transport [36].

However, it is important to note that respiratory Complexes I, III, and IV, but not Complex II, are embedded in the inner mitochondrial membrane, as well as the Fo moiety of ATP synthase. This suggests that Complex II could be less prone to be affected by laser treatment. Moreover, it has been demonstrated that only Complexes I, III, and IV, as well as ATP synthase, are organized in a super complex, probably to increase electron transport and make the relative energy production more efficient [37–39]. However, the irradiation could interact not only with the cytochromes that constitute the respiratory complexes but also with ubiquinone or cytochrome c, the electron shuttles between Complexes I or II with III, and between III and IV, respectively.

Contextually, Pasternak et al. [40] demonstrated that 808 and 905 nm laser light may induce functional and structural modifications in cell membranes.

The H3O + and OH - ions are important species in biology, chemical physics, and electrochemistry, which carry acid-base reactions and the charge transfer process in aqueous solutions, by serving as intermediates for protonic transport [41]. Recently, authors studied the infrared spectra of pure water and mixtures to show the hidden IR vibrations of water’s ionic species IR spectrum affected it and was partially defined by the dynamics of ionic species, which were influenced. Interestingly, Walski et al. [42] suggested the polychromatic light (750–2000 nm) disturbs the energy of hydrogen bonds, increases water molecules dissociation, and consequently, alters biological membrane features. In particular, these alterations could affect particular lipids, such as cardiolipin mentioned above, which are involved in the ability of electron transport between the respiratory complexes and/or in the proton transport through them, altering OxPhos functionality.

Indeed, the paradigm of mechanical vibrations of molecules is one of the accredited models that explain nonionizing radiation effects on cells, by interaction with water, transmembrane proteins, and phospholipidic bilayers to produce changes in membrane fluidity, permeability, and protein activities [43–45].

As mentioned in the introduction, ROS are products of oxygen metabolism, as the natural consequence of mitochondrial respiration in all aerobic organisms. However, under physiological conditions, mitochondria keep ROS levels under control by a system of ROS-scavenging that prevents reactive species increment and, thus, cell damage [5]. In this way, mitochondria and low concentrations of ROS can play a pivotal role in cellular homeostasis through modulation of cellular signalling pathways.

Conversely, when there are high levels of ROS and the production/scavenging system becomes unbalanced, oxidative stress occurs [2], leading to indiscriminate damage to biological molecules and loss of cell functions and cell death.

Our data showed some irradiation can induce increments of ROS production. However, we must make a distinction between the lower and higher dose effect, in terms of the “amount of” and “modality with.” In particular, the increment of superoxide production observed after irradiation with 0.1 W probably depends on the uncoupled status, although the OxPhos activity was reduced in comparison to the control. This depends on the fact that mitochondria are considered one of the principal sources of oxidative stress production, which increases when ATP production and oxygen consumption are uncoupled [21]. Moreover, increased oxidative stress production could induce a vicious circle in which the alteration of OxPhos activity due to the laser treatment increases the oxidative stress that, in turn, negatively affects OxPhos activity [46]. This hypothesis was confirmed by the massive accumulation of MDA, which suggested that oxidative stress was not counterbalanced by sufficient antioxidant defences, despite mitochondria expressing both catalase and superoxide dismutase type 2 as antioxidant enzymes. In contrast, we worked with isolated mitochondria, which may have partly lost detoxifying enzymes and scavenger molecules during their preparation. Moreover, the monoelectron transport between the respiratory complexes determines an easier oxidative stress production, both in terms of reactive oxygen species and lipid peroxidation, whether the respiratory complexes and the relative shuttle are altered.

Conversely, the increment of superoxide production and MDA accumulation in mitochondria irradiated with 0.8 W could be associated simply with the increment of OxPhos activity, which, in perfect coupled conditions, produced oxidative stress from Complexes I and III [47, 48].

Chinopoulos and Adam-Vizi [49] described that in addition to the mitochondrial respiratory chain, the Krebs cycle should also have a role in mitochondrial ROS management. However, our data showed no effect of 980 nm laser light on IDH and MDH, as well as Complex II (called also succinic dehydrogenase, one of the eight enzymes involved in the Krebs cycle other than the second respiratory complex). This suggested that in our experimental conditions, ROS formation was due to the modulation of respiratory chain activity that occurs as a consequence of light-photoacceptor interaction (copper, iron, water) more than a generic increment of temperature, increments of temperature that were macroscopically considered, measured, and avoided in our experimental design.

Notably, oxidative stress could be induced in the cell by light-mitochondrial interaction through an alternative way than the respiratory chain and oxygen consumed. In this respect, in our previous works, we demonstrated that “808 nm and 980 nm infrared laser light directly affect the stored Ca 2+ homeostasis, independent of the mitochondrial respiratory chain activities” [50] mitochondria are one of the major reserves of sequestered calcium [51]. Additionally, we also showed a connection between the modulation of calcium homeostasis and both nitric oxide production and glutamate release in a unicellular organism [52] and the murine nerve terminal [53]. The reciprocal relationship among ROS formation elevated intracellular calcium concentration, and the/mitochondrial death is known [49]. Therefore, PBM besides both positive stimulation and support of cell metabolism could, if wrongly administered, affect mediators of cell injury, such as Ca 2+ , ROS, and reactive nitrogen species formation and latent glutamate-induced delayed calcium deregulation [8].

5. Conclusions

In conclusion, we showed for the first time the ability of the 980 nm diode laser light to interact with the mitochondria from bovine liver. The interaction behaved as a window effect and interested Complexes III and IV, as well as ATP production and oxygen consumption. Investigation of the effect of 0.1 W power irradiated for 60 sec highlighted that photobiomodulation can uncouple the respiratory chain, induce higher oxidative stress, and cause drastic inhibition of ATP production. Conversely, 0.8 W kept mitochondria coupled and induced increments of ATP production by increments of Complex III and IV activities an increment of oxidative stress was also observed, as a likely consequence of the increased oxygen consumption and mitochondrial isolation experimental conditions.

Data Availability

The data used to support the findings of this study are available from the corresponding author upon request.

Conflicts of Interest

The authors declare no conflicts of interest.


The authors would like to express special appreciation and thanks to Prof. Alberico Benedicenti, for his guidance to our work.

Supplementary Materials

Figure 1 supplementary: experimental design. Mitochondria were isolated from bovine liver. Mitochondria samples were irradiated at the room-air temperature or with the tube sample immersed in water. The samples were then processed for the biochemical analysis. Figure 2 supplementary: representations of sample temperature behaviour during the experiments. Before: temperature of the sample before irradiation. Room-air after: temperature of the sample after irradiation performed at room-air temperature. Room-air after+reagents: temperature of the sample after irradiation performed at a room-air temperature and the addition of reagents for biochemical evaluation. Water after: temperature of the sample after irradiation performed with the sample partially immersed in water. (Supplementary Materials)


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