# Catalytic Reactions and Energy Conservation in the Cytochrome bc 1 and b 6 f Complexes of Energy-Transducing Membranes

Marcin Sarewicz, Sebastian Pintscher, Rafal Pietras, Arkadiusz Borek, Łukasz Bujnowicz, Guy Hanke, William Cramer, Giovanni Finazzi, Artur Osyczka

<figure><img src="image_1.png" /></figure>

## ► To cite this version:

Marcin Sarewicz, Sebastian Pintscher, Rafal Pietras, Arkadiusz Borek, Łukasz Bujnowicz, et al. Catalytic Reactions and Energy Conservation in the Cytochrome bc 1 and b 6 f Complexes of Energy-Transducing Membranes. Chemical Reviews, 2021, 121 (4), pp.2020-2108. 10.1021/acs.chemrev.0c00712. [hal-03122713](https://hal.science/hal-03122713)

HAL Id: hal-03122713

https://hal.science/hal-03122713

Submitted on 1 Feb 2021

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<figure><img src="image_2.png" /></figure>

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<table>
  <tbody>
    <tr>
      <td>4.1</td>
      <td>Basic redox properties of Quinones</td>
      <td>45</td>
    </tr>
    <tr>
      <td>4.1.1</td>
      <td>Stability and reactivity of semiquinone radical with molecular oxygen</td>
      <td>48</td>
    </tr>
    <tr>
      <td>4.1.2</td>
      <td>Stability constant of SQ<sup>-</sup></td>
      <td>49</td>
    </tr>
    <tr>
      <td>4.1.3</td>
      <td>Chloroplast photoactive vs non photoactive plastooquinone pools</td>
      <td>50</td>
    </tr>
    <tr>
      <td>4.2</td>
      <td>Cytochrome c and plastocyanin as electron acceptors</td>
      <td>51</td>
    </tr>
    <tr>
      <td>4.2.1</td>
      <td>Redox properties and Interaction of Cytochromes c with Cytochrome bc<sub>1</sub></td>
      <td>53</td>
    </tr>
    <tr>
      <td>4.2.2</td>
      <td>Redox properties and Interaction of Plastocyanin with Cytochrome b<sub>6</sub>f</td>
      <td>58</td>
    </tr>
    <tr>
      <td>4.3</td>
      <td>FNR/Fd interactions with cytochrome b<sub>6</sub>f</td>
      <td>61</td>
    </tr>
    <tr>
      <td>4.3.1</td>
      <td>Potential n side donors</td>
      <td>61</td>
    </tr>
    <tr>
      <td>4.3.2</td>
      <td>Direct electron donation from Fd to cytochrome b<sub>6</sub>f</td>
      <td>62</td>
    </tr>
    <tr>
      <td>4.3.3</td>
      <td>Direct electron donation from FNR to the cytochrome b<sub>6</sub>f</td>
      <td>65</td>
    </tr>
    <tr>
      <td>5</td>
      <td>Inhibitors</td>
      <td>66</td>
    </tr>
    <tr>
      <td>5.1</td>
      <td>Cytochrome bc<sub>1</sub> specific inhibitors</td>
      <td>66</td>
    </tr>
    <tr>
      <td>5.2</td>
      <td>Cytochrome b<sub>6</sub>f-specific inhibitors</td>
      <td>83</td>
    </tr>
    <tr>
      <td>6</td>
      <td>Thermodynamic background of cytochrome bc catalysis</td>
      <td>88</td>
    </tr>
    <tr>
      <td>7</td>
      <td>Mechanistic insights into the catalytic Q<sub>o</sub> / Q<sub>p</sub> site</td>
      <td>90</td>
    </tr>
    <tr>
      <td>7.1</td>
      <td>Overview of the structure of the Q<sub>o</sub> site</td>
      <td>90</td>
    </tr>
    <tr>
      <td>7.2</td>
      <td>Quinone binding to the Q<sub>o</sub> site</td>
      <td>91</td>
    </tr>
    <tr>
      <td>7.3</td>
      <td>Crucial amino acid residues involved in the binding and catalysis of QH<sub>2</sub> / PQH<sub>2</sub> oxidation at the Q<sub>o</sub>/Q<sub>p</sub> site</td>
      <td>94</td>
    </tr>
    <tr>
      <td>7.3.1</td>
      <td><sup>Rh</sup>H156 (<sup>Bt</sup>H161) in iron-sulfur protein and the <sup>Rh</sup>E295 (<sup>Bt</sup>E271) in cytochrome b95</td>
      <td></td>
    </tr>
    <tr>
      <td>7.3.2</td>
      <td>Cytb:<sup>Rh</sup>Y147 (<sup>Bt</sup>Y131) and Cytb:<sup>Rh</sup>Y302 (<sup>Bt</sup>Y278)</td>
      <td>97</td>
    </tr>
    <tr>
      <td>7.3.3</td>
      <td>Other residues involved in substrate binding to the Q<sub>o</sub> site</td>
      <td>99</td>
    </tr>
    <tr>
      <td>7.3.4</td>
      <td>Specific residues involved in PQ/PQH<sub>2</sub> binding to the Q<sub>p</sub> site of Cytochrome b<sub>6</sub>f99</td>
      <td></td>
    </tr>
    <tr>
      <td>7.4</td>
      <td>Catalytic and side reactions at the QH<sub>2</sub>-oxidation site of Cytochromes bc</td>
      <td>100</td>
    </tr>
    <tr>
      <td>7.4.1</td>
      <td>Forward, reverse and short-circuits reactions at the Q<sub>o</sub>/Q<sub>p</sub> site</td>
      <td>102</td>
    </tr>
    <tr>
      <td>7.4.2</td>
      <td>The possible physiological meaning of short-circuits</td>
      <td>105</td>
    </tr>
    <tr>
      <td>7.4.3</td>
      <td>Short-circuit suppression mechanisms</td>
      <td>106</td>
    </tr>
    <tr>
      <td>7.5</td>
      <td>Superoxide generation at the Q<sub>o</sub>/Q<sub>p</sub> site</td>
      <td>112</td>
    </tr>
    <tr>
      <td>7.6</td>
      <td>Semiquinone intermediate at the Q<sub>o</sub> / Q<sub>p</sub> site</td>
      <td>120</td>
    </tr>
    <tr>
      <td>8</td>
      <td>Mechanistic insights into the catalytic Q<sub>i</sub>/Q<sub>n</sub> site</td>
      <td>125</td>
    </tr>
    <tr>
      <td>8.1</td>
      <td>Overview of the structure of Q<sub>i</sub> and the proton paths</td>
      <td>125</td>
    </tr>
    <tr>
      <td>8.2</td>
      <td>Catalytic electron and proton transfers at the Q<sub>i</sub> site of Cytbc<sub>1</sub></td>
      <td>127</td>
    </tr>
    <tr>
      <td>8.3</td>
      <td>Fast- and slow-relaxing SQ<sub>i</sub> as dominant intermediates of forward and reverse reactions at the Q<sub>i</sub> site, respectively</td>
      <td>128</td>
    </tr>
    <tr>
      <td>8.4</td>
      <td>Charge polarization of SQ<sub>i</sub> facilitates electron and proton reactions at the Q<sub>i</sub> site</td>
      <td>130</td>
    </tr>
    <tr>
      <td>8.5</td>
      <td>Specific residues involved in PQ/PQH<sub>2</sub> binding to the Q<sub>n</sub> site of Cytb<sub>6</sub>f</td>
      <td>132</td>
    </tr>
    <tr>
      <td>9</td>
      <td>Inter-monomer electron transfer</td>
      <td>134</td>
    </tr>
    <tr>
      <td>10</td>
      <td>Higher level of organization and regulation (supercomplexes, state transition, kinase activation)</td>
      <td>139</td>
    </tr>
    <tr>
      <td>10.1</td>
      <td>A role for cytochrome b<sub>6</sub>f in the regulation of linear vs cyclic electron flow?</td>
      <td>139</td>
    </tr>
    <tr>
      <td>10.2</td>
      <td>The "elusive" Cytb<sub>6</sub>f-PSI CET supercomplex</td>
      <td>141</td>
    </tr>
    <tr>
      <td>10.3</td>
      <td>Role of cytochrome b<sub>6</sub>f in state transitions</td>
      <td>143</td>
    </tr>
    <tr>
      <td>11</td>
      <td>Comments on the nomenclature used for Cytbc<sub>1</sub> and Cytb<sub>6</sub>f</td>
      <td>146</td>
    </tr>
    <tr>
      <td>12</td>
      <td>References</td>
      <td>147</td>
    </tr>
  </tbody>
</table>
List of abbreviations

<table><tr><td>Δμ<sub>H</sub><sup>+</sup></td><td>The transmembrane electrochemical potential gradient</td></tr><tr><td>2Fe2S</td><td>Rieske 2Fe-2S cluster</td></tr><tr><td>ADP</td><td>Adenosine diphosphate</td></tr><tr><td>ATP</td><td>Adenosine triphosphate</td></tr><tr><td>b<sub>H</sub></td><td>The high-potential heme b of Cytbc<sub>1</sub></td></tr><tr><td>b<sub>H</sub>/b<sub>n</sub></td><td>The high-potential heme b of Cytbc<sub>1</sub> or heme b at n-side of Cytb<sub>6f</sub></td></tr><tr><td>b<sub>L</sub></td><td>The low-potential heme b of Cytbc<sub>1</sub></td></tr><tr><td>b<sub>L</sub>/b<sub>p</sub></td><td>The low-potential heme b of Cytbc<sub>1</sub> or heme b at p-side of Cytb<sub>6f</sub></td></tr><tr><td>b<sub>n</sub></td><td>Heme b at n-side of Cytb<sub>6f</sub></td></tr><tr><td>b<sub>p</sub></td><td>Heme b at p-side of Cytb<sub>6f</sub></td></tr><tr><td>CcO</td><td>Cytochrome c oxidase</td></tr><tr><td>CET</td><td>Cyclic Electron Transfer</td></tr><tr><td>CW</td><td>Continuous wave</td></tr><tr><td>Cyt-<i>bc</i></td><td>Enzyme belonging to cytochrome <i>bc</i><sub>1</sub> or cytochrome <i>b</i><sub>6f</sub></td></tr><tr><td>Cytb</td><td>Cytochrome <i>b</i></td></tr><tr><td>Cytb<sub>6</sub></td><td>Cytochrome <i>b</i><sub>6</sub></td></tr><tr><td>Cytb<sub>6f</sub></td><td>Cytochrome <i>b</i><sub>6f</sub> complex</td></tr><tr><td>Cytbc<sub>1</sub></td><td>Cytochrome <i>bc</i><sub>1</sub> complex</td></tr><tr><td>Cytc, Cytc<sub>2</sub>, Cytc<sub>6</sub></td><td>Cytochrome <i>c</i>, <i>c</i><sub>2</sub>, <i>c</i><sub>6</sub></td></tr><tr><td>Cytc<sub>1</sub></td><td>Cytochrome <i>c</i><sub>1</sub></td></tr><tr><td>Cytf</td><td>Cytochrome<sub>f</sub></td></tr><tr><td>DBMIB</td><td>Dibromothymoquinone</td></tr><tr><td>DMPO</td><td>5,5-Dimethyl-1-pyrroline N-oxide (spin trap)</td></tr><tr><td>ECS</td><td>Electro Chromic Band Shift</td></tr><tr><td>E<sub>m</sub></td><td>Redox midpoint potential</td></tr><tr><td>E<sub>m,x</sub></td><td>Redox midpoint potential at pH=x</td></tr><tr><td>EPR</td><td>Electron paramagnetic resonance</td></tr><tr><td>ES</td><td>Enzyme – substrate complex</td></tr><tr><td>ET</td><td>Electron transfer</td></tr><tr><td>ETC</td><td>Electron transport chain</td></tr><tr><td>Fd</td><td>Ferredoxin</td></tr><tr><td>heme <i>c</i><sub>1</sub></td><td>Type c heme in cytochrome <i>c</i><sub>1</sub></td></tr><tr><td>IMS</td><td>Intermembrane space</td></tr><tr><td>ISP</td><td>Iron-sulfur protein</td></tr><tr><td>ISP-HD</td><td>Mobile head domain of the iron-sulfur protein</td></tr><tr><td>K<sub>d</sub></td><td>Dissociation constant</td></tr><tr><td>MD</td><td>Molecular dynamics</td></tr><tr><td>MK</td><td>Menaquinone</td></tr></table>
<table>
  <tr>
    <td>NMR</td>
    <td>Nuclear magnetic resonance</td>
  </tr>
  <tr>
    <td>p, n side</td>
    <td>Electrochemically positive, negative sides of membrane</td>
  </tr>
  <tr>
    <td>PC</td>
    <td>Plastocyanin</td>
  </tr>
  <tr>
    <td>P<sub>f</sub></td>
    <td>An inhibitor which fixes ISP-HD at b-position</td>
  </tr>
  <tr>
    <td>P<sub>i</sub></td>
    <td>Phosphate</td>
  </tr>
  <tr>
    <td>P<sub>m</sub></td>
    <td>An inhibitor which does not fix ISP-HD</td>
  </tr>
  <tr>
    <td>pmf</td>
    <td>Transmembrane electrochemical potential gradient, Δμ<sub>H</sub><sup>+</sup></td>
  </tr>
  <tr>
    <td>PQ</td>
    <td>Plastoquinone</td>
  </tr>
  <tr>
    <td>PQ(H<sub>2</sub>)</td>
    <td>Plastoquinone or plastoquinol</td>
  </tr>
  <tr>
    <td>PQH<sub>2</sub></td>
    <td>Plastoquinol</td>
  </tr>
  <tr>
    <td>PSI</td>
    <td>Photosystem I</td>
  </tr>
  <tr>
    <td>PSII</td>
    <td>Photosystem II</td>
  </tr>
  <tr>
    <td>PSQ</td>
    <td>Plastosemiquinone</td>
  </tr>
  <tr>
    <td>PT</td>
    <td>Proton transfer</td>
  </tr>
  <tr>
    <td>Q</td>
    <td>Quinone (in general)</td>
  </tr>
  <tr>
    <td>Q(H<sub>2</sub>)</td>
    <td>Quinone or quinol</td>
  </tr>
  <tr>
    <td>QH<sub>2</sub></td>
    <td>Quinol</td>
  </tr>
  <tr>
    <td>Q<sub>i</sub></td>
    <td>Quinone reduction site of cytochrome bc<sub>1</sub> complex</td>
  </tr>
  <tr>
    <td>Q<sub>i</sub>/Q<sub>n</sub></td>
    <td>The quinone reduction site of Cytc<sub>c1</sub> or Cytb<sub>6f</sub></td>
  </tr>
  <tr>
    <td>Q<sub>n</sub></td>
    <td>Plastoquinone reduction site of cytochrome b<sub>6f</sub> complex</td>
  </tr>
  <tr>
    <td>Q<sub>o</sub></td>
    <td>quinol oxidation site of cytochrome bc<sub>1</sub> complex</td>
  </tr>
  <tr>
    <td>Q<sub>o</sub>/Q<sub>p</sub></td>
    <td>The quinol oxidation site of Cytc<sub>c1</sub> or Cytb<sub>6f</sub></td>
  </tr>
  <tr>
    <td>Q<sub>os</sub></td>
    <td>The strong-binding niche of Q<sub>o</sub> site</td>
  </tr>
  <tr>
    <td>Q<sub>ow</sub></td>
    <td>The weak-binding niche of Q<sub>o</sub> site</td>
  </tr>
  <tr>
    <td>Q<sub>p</sub></td>
    <td>Plastoquinol oxidation site of cytochrome b<sub>6f</sub> complex</td>
  </tr>
  <tr>
    <td>RC</td>
    <td>reaction center</td>
  </tr>
  <tr>
    <td>RhA<sub>xxx</sub>,<br/>BtA<sub>xxx</sub>,<br/>ScA<sub>xxx</sub>, A residue of code “A” at position number xxx from Rhodobacter capsulatus/sphaeroides, Bos taurus, Saccharomyces cerevisiae, Mastigocladus laminosus (Cytb<sub>6f</sub>), respectively</td>
    <td></td>
  </tr>
  <tr>
    <td>ROS</td>
    <td>reactive oxygen species</td>
  </tr>
  <tr>
    <td>SMP</td>
    <td>Submitochondrial particles</td>
  </tr>
  <tr>
    <td>SMP</td>
    <td>Sub-mitochondrial particles</td>
  </tr>
  <tr>
    <td>SQ</td>
    <td>General semiquinone</td>
  </tr>
  <tr>
    <td>SQ<sup>-</sup></td>
    <td>Semiquinone anion</td>
  </tr>
  <tr>
    <td>SQ-2Fe2S</td>
    <td>The semiquinone at the Q<sub>o</sub>/Q<sub>p</sub> site spin-coupled to the reduced 2Fe2S cluster</td>
  </tr>
  <tr>
    <td>SQH</td>
    <td>Protonated semiquinone</td>
  </tr>
  <tr>
    <td>SQ<sub>i</sub></td>
    <td>Semiquinone at the Q<sub>i</sub> site of Cytc<sub>c1</sub></td>
  </tr>
  <tr>
    <td>SQ<sub>o</sub></td>
    <td>Semiquinone at the Q<sub>o</sub> site of Cytc<sub>c1</sub></td>
  </tr>
  <tr>
    <td>TMH</td>
    <td>Transmembrane helix</td>
  </tr>
  <tr>
    <td>UQ</td>
    <td>Ubiquinone</td>
  </tr>
  <tr>
    <td>UQ(H<sub>2</sub>)</td>
    <td>Ubiquinone or ubiquinol</td>
  </tr>
  <tr>
    <td>UQH<sub>2</sub></td>
    <td>Ubiquinol</td>
  </tr>
</table>
<table><tr><td>UQH<sub>2</sub><sup>Qos</sup></td><td>Ubiquinol bound at the strong-binding niche</td></tr><tr><td>UQH<sub>2</sub><sup>Qow</sup></td><td>Ubiquinol bound at the weak binding niche</td></tr><tr><td>UQ<sup>Qos</sup></td><td>Ubiquinone bound at the strong-binding niche</td></tr><tr><td>UQ<sup>Qow</sup></td><td>Ubiquinone bound at the weak binding niche</td></tr><tr><td>USQ</td><td>Ubisemiquinone</td></tr></table>
Abstract

This review focuses on key components of respiratory and photosynthetic energy-transduction
systems: the cytochrome *bc*₁ and *b*₆*f* (Cyt*bc*₁/*b*₆*f*), membranous multi-subunit homodimeric
complexes. These remarkable molecular machines catalyze electron transfer from membranous
quinones to water-soluble electron carriers (such as cytochromes *c* or plastocyanin), coupling electron
flow to proton translocation across the energy-transducing membrane and contributing to the
generation of a transmembrane electrochemical potential gradient which powers cellular metabolism
in the majority of living organisms. Cyts*bc*₁/*b*₆*f* share many similarities but also have significant
differences. While decades of research have provided extensive knowledge on these enzymes, several
important aspects of their molecular mechanisms remain to be elucidated. We summarize a broad
range of structural, mechanistic and physiological aspects required for function of Cyt*bc*₁/*b*₆*f*,
combining textbook fundamentals with new intriguing concepts that have emerged from more recent
studies. The discussion covers, but is not limited to: i) mechanisms of energy-conserving bifurcation
and energy-wasting superoxide generation at the quinol oxidation site, ii) the mechanism by which
semiquinone is stabilized at the quinone reduction site, iii) interactions with substrates and specific
inhibitors, iv) inter-monomer electron transfer and the role of a dimeric complex, and v) higher levels
of organization and regulation that involve Cyts*bc*₁/*b*₆*f*. In addressing these topics we point out
existing uncertainties and controversies which, it is suggested, will drive further research in this field.
# 1 Introduction

Cytochromes *bc* (Cytbc) constitute one of the broadest groups of energy-transducing enzymes, present in almost every living cell. They are key components of both respiratory and photosynthetic electron transport chains and their function is often indispensable for the operation of these chains.

In this review we focus on two important enzymes from that group: cytochrome *bc*₁ (Cytbc₁) – involved in respiration and bacterial photosynthesis and cytochrome *b*₆f (Cytb₆f) – studied in plant, algal and cyanobacterial photosynthesis. We summarize a broad range of structural, mechanistic and physiological aspects required for function of these enzymes, presenting both the textbook fundamentals based on the well-established ideas and the new intriguing concepts and attractive hypotheses emerging from more recent studies.

It is remarkable that the extensive knowledge gathered so far on Cytbc has left many key issues not satisfactorily understood and opened new intriguing issues for consideration. Among subjects still intensely debated are: the molecular mechanism of the catalytic electron bifurcation and the basis of high energetic efficiency of the enzyme, the mechanism of proton transfer to and from the catalytic sites, the role of dimer and the inter-monomer electron transfer, and the role of Cytbc in regulation of electron flow and in redox (ROS) signaling.

It should be noted that in some cases there are questions about mechanisms of Cytbc structure-function have been regarded as beyond doubt. Despite the fact that the catalytic Q cycle, the conceptual framework in which Peter Mitchell described principles of his chemiosmotic theory, is widely accepted for Cytsbc₁, its applicability to details of the function of Cytsb₆f remain subject to some questions. Although that the scheme showing cyclic electron transfer (CET) appears in every textbook of photosynthesis, the actual pathway through which electrons recycle back to the main electron transport chain remains controversial. It is clear that all these and many other questions related to this important group of enzymes await resolution in future studies.

Even though our intention was to comprehensively cover the knowledge on Cytbc, the breadth of the subject makes it impossible to include all issues which relate to it. The reader is referred to several excellent reviews on this subject that have appeared in the literature.¹⁻²⁶

# 2 Energy Conserving and Regulatory role of Intracellular Cytochromes-bc

Conversion of solar or chemical energy into a biologically useful form is one of the most fundamental processes of living organisms. As described by chemiosmotic theory ²⁷,²⁸, this process
involves generation of a transmembrane electrochemical potential gradient, $\Delta\mu_H^+$ (or proton motive force, pmf) by enzymes which use external sources of energy to couple electron transfer to proton translocation between cellular compartments separated by a membrane (termed energy-transducing membrane)$^{29}$. PMF forces the ATP-synthase to convert ADP and $P_i$ into ATP, and the energy is stored by maintaining the concentrations of ATP, ADP, and $P_i$ far from their equilibrium values, thus providing the universal source of energy that powers cellular metabolism.$^{29}$

The enzymes that generate pmf are typically organized in chains, such as the mitochondrial and photosynthetic electron transfer chains, where the individual membrane-embedded electron transport protein complexes are linked functionally by components having diffusional freedom to move between complexes: hydrophobic low-molecular weight electron carriers and water-soluble protein electron carriers, respectively, inside and outside the membrane$^{30,31}$. These carriers shuttle electrons between complexes which, overall, catalyze electron flow from a primary electron donor to a final electron acceptor, while the energy released during this process is used to transfer protons across the membrane.

There are two general mechanisms in electron transport chains by which protons are transferred across the energy-transducing membrane. In one mechanism, reduction of a quinone (coupled to proton uptake) occurs at one side of the membrane while quinol oxidation (coupled to proton release) takes place at the opposite side of the membrane (Mitchell's redox loop$^{27,32,33}$). In the other mechanism, a series of protonation/deprotonation reactions taking place within the specific proton channels of an electron transfer complex (e.g. complex I and IV of mitochondrial electron transport chain) involving successive pK changes of amino acids in the trans-membrane pathway, allow the pumping of protons between the two sides of the membrane without involvement of quinone molecules.

Cytochromes *bc* complexes (Cyt-*bc*) comprise a set of the ubiquitous complexes contributing to generation of pmf, through a mechanism involving quinone/quinol oxidation-reduction reactions. This superfamily of complexes includes cytochrome *bc*₁ (Cyt*bc*₁) (ubiquinol: cytochrome *c* oxidoreductase, EC 7.1.1.8) and cytochrome *b*₆f (Cyt*b*₆f) (plastohydroquinone:plastocyanin oxidoreductase, EC 7.1.1.6). Cyt*bc*₁ provides an electronic connection between quinone molecules and water-soluble electron carriers (such as cytochrome *c*, *c*₂ or *c*₆) catalyzing a net reaction of oxidation of quinol (ubiquinol in mitochondria and several prokaryotic cells or menaquinol some bacteria, see section 4.1) and reduction of water-soluble electron carrier (section 4.2.1). Cyt*b*₆f catalyzes an analogous reaction between membrane embedded plastoquinol molecules and the water-soluble plastocyanin (PC) of chloroplasts or Cyt*c*₆ (in cyanobacteria and some microalgae) (section 4.2.2). In all cases, the energy released in these electron transfer reactions powers the oxidation of
quinol and reduction of quinone, taking place at the distinct catalytic sites located on the opposite sides of the membrane, according to the first mechanism of H⁺ translocation described above.

Cytbc₁ participates in respiration in oxygen-utilizing cells and also electron transfer in numerous bacteria which utilize alternative terminal electron acceptors in addition to oxygen (Fig. 1). In mitochondria, Cytbc₁ (also referred to as mitochondrial complex III or respiratory complex III) is a confluence point for reducing equivalents from the various dehydrogenases (mitochondrial complexes I, II) (Fig 1 A). In animal mitochondria, with the sole exception of some parasites, it is essential for mitochondrial respiration, as there is no alternative route to oxidation of ubiquinol by molecular oxygen. This enzyme is not essential in bacteria, where alternative mechanisms to oxidize ubiquinol usually exist in parallel to Cytbc₁. On the other hand, Cytbc₁ is an essential enzyme, in some photosynthetic bacteria such as *Rhodobacter sphaeroides* or *R. capsulatus* for cyclic, non oxygenic, photosynthetic electron transfer in the absence of a terminal electron acceptor but it is not essential when these organisms are grown heterotrophically (Figure 1 B)³⁴.

Cytb₆f participates in oxygenic photosynthesis in cyanobacteria, and plant and algal chloroplasts. It is a crucial component of the linear electron transfer ('Z scheme' or LET) linking photosystem I (PSI) with photosystem II (PSII) (Figure 1 C). In addition it also participates in cyclic electron transfer (CET) (Fig. 1 C) (see sections 3.6.5 and 10.1).

Apart from the involvement in generating pmf, Cytbc are believed to be potential points of regulation of electron flow. This stems from the fact that their two quinone binding sites are directly connected with the two main redox pools (intra-membrane ubiquinone/plastoquinone and soluble cytochrome c/plastocyanin) of respiration/photosynthesis. Hence, by virtue of catalyzing opposite chemical reactions, these sites may influence one another (the product of one catalytic site becomes a substrate for the second site and *vice versa*). Another regulatory function of Cytb₆f is to trigger changes in the chloroplasts' light harvesting capacity in response to changes in the ambient light intensity by sensing variations in the redox state of the plastoquinone pool. These changes modulate the interaction of Cytb₆f with a kinase able to reversibly phosphorylate the light harvesting complexes, leading to their preferential association with one of the two photo centers (PSII and PSI) in a process called 'state transitions' (section 10.3)³⁵.
<figure><img src="image_3.png" /></figure>

<figure><img src="image_4.png" /></figure>

<figure><img src="image_5.png" /><figcaption>Figure 1. Overview of electron transfer paths involving Cyt-bc complexes: Cytbc₁ in mitochondrial respiratory chain (A) and in anoxygenic photosynthesis or respiration of purple bacteria (B); Cytb₆f in oxygenic photosynthesis in chloroplasts (C). Q and PQ denote ubiquinone and plastoquinone pool, respectively. PC, Fd and FNR denote plastocyanin, ferredoxin and ferredoxin/NADP⁺ oxidoreductase. LET depicts linear electron transfer in photosynthesis while CET shows cyclic electron transfer path which recycles the electrons from Fd pool back to the PQ(H₂) pool. Scheme in A considers mitochondria of both plants and animals (the diiron alternative oxidase is not present in animal mitochondria except for rare cases, such as in *Trypanosoma*).</figcaption></figure>

Moreover, although the activities of Cytbc₁ and Cytb₆f complexes are different, because of the above reactions, they are also inferred to be involved in mitochondrial and chloroplast redox signaling.³⁶

Finally under specific conditions, Cytbc can generate a limited amount of superoxide, as a side reaction of the catalytic cycle (section 7.5).³⁷

$$O_2 + e^- \rightarrow O_2^-; E_m = -140 \text{ mV}$$
It is noted that the specific rate of superoxide ($O_2^−$) production in purified active crystallizable Cytb<sub>6f</sub>, normalized to the rate of electron transport, is more than an order of magnitude greater than that of the yeast respiratory Cytbc<sub>1</sub><sup>38</sup>

**3 Overview of Structure and Function of Cytochromes bc**

Cytbc<sub>1</sub> and Cytb<sub>6f</sub> share several structural and mechanistic features. Both are homodimers with each monomer containing a set of similarly arranged spatially separated quinone binding sites and redox active cofactors required for completion of the catalytic cycle (fig. 2). The general features common to both complexes, are summarized in sections 3.1-3.4. Despite these similarities, several important differences exist between them. Therefore the two following sections of this chapter 3.5 and 3.6 discuss separately Cytbc<sub>1</sub>- and Cytb<sub>6f</sub>-specific properties.

**3.1 Catalytic Sites, Redox cofactors, and Cofactor Chains**

In each monomer of the Cyt-*bc* dimeric complex, the hemes and the iron-sulfur 2Fe-2S Rieske-type cluster (2Fe2S) assemble into two chains of cofactors designed to link the two catalytic quinone binding sites with diffusible pools of substrates (quinone and Cytc/PC). The two quinone binding sites catalyze opposing reactions (quinol oxidation / quinone reduction) as integral parts of the catalytic cycle (see section 3.2).

In Cytbc<sub>1</sub>, the quinol oxidation site (the Q<sub>o</sub> site) is linked with Cytc by the *c*-chain composed of 2Fe2S and heme *c*<sub>1</sub>. The *b*-chain, composed of heme *b*<sub>L</sub>/*b*<sub>p</sub> and heme *b*<sub>H</sub>/*b*<sub>n</sub>, connects the Q<sub>o</sub>/Q<sub>p</sub> site with the quinone reduction site (the Q<sub>i</sub>/Q<sub>n</sub> site) located at the opposite side of the membrane. Considering the relative values of the redox midpoint potentials of cofactors, the *c*-chain and the *b*-chain are commonly named the high- and low-potential chain, respectively. A characteristic feature of this arrangement is that the Q<sub>o</sub> site is located between the *c*- and *b*-chains, which has crucial consequences for the catalytic Q cycle mechanism (Figure 2).

In Cytb<sub>6f</sub>, the high- and low-potential chains and the quinone catalytic sites (the Q<sub>p</sub> and Q<sub>n</sub> for quinol oxidation and quinone reduction, respectively, see section 11 for explanation on the usage of the names of the catalytic sites in this review and on other nomenclatural issues) are arranged in the similar manner, but some variation in the redox cofactor exist: heme *f* (*E*<sub>m</sub> = +370 mV) is present in the high-potential chain in place of heme *c*<sub>1</sub> and the low-potential chain contains an additional and atypical heme *c*<sub>n</sub> (not present in Cytbc<sub>1</sub>).
<figure><img src="image_6.png" /><figcaption>Figure 2. Overview of structures and cofactor chains in dimeric Cyt-bc complexes. Top: Crystal structure representations of *M. laminosus* Cytb<sub>6f</sub> (left, PDB ID: 4H13), *R. capsulatus* Cytbc<sub>1</sub> (middle, PDB ID: 1ZRT) and mitochondrial Cytbc<sub>1</sub> (right, PDB ID: 1KYO). Bottom: Spatial arrangement of cofactors extracted from the structures above. The layout of cofactors in *R. capsulatus* and mitochondrial Cytbc<sub>1</sub> is the same. The simplified cartoon models depict symbols for cofactors that are used throughout the figures.</figcaption></figure>

## 3.2 Brief Overview of Catalytic Cycle

The mechanism by which Cytbc₁ links electron transfer and proton translocation reactions is the proton motive Q cycle.<sup>33,39</sup> It engages the high- and low-potential -chains (c- and b-chains, respectively) and the Qo, and the Qi sites. Protons are taken up at the Qi site, carried across the membrane by the quinol (QH₂), and released at the Qo site. The Qi site and the Qo site are located toward the electronegative and electropositive sides of the membrane (n and p sides, respectively). For that reason, the Qi site and Qo of Cytbc₁ site are alternatively named Qn and Qp, respectively (see section 11).

The Qo site, being positioned between the c- and b-chains, oxidizes QH₂ and passes its two electrons in a highly specific manner: one electron is delivered to the c-chain while the other electron is delivered to the b-chain. This process occurs in a so-called bifurcation reaction in which QH₂ is oxidized by 2Fe2S of the c-chain and by the heme bL of the b chain. The reduced 2Fe2S passes 1 electron to heme c₁, which subsequently reduces diffusible Cytc. The reduced heme bL passes 1 electron to heme bH. The electron is then used for quinone (Q) reduction at the Qi site. This way the Qo-site-mediated oxidation of QH₂ delivers one electron to the Qi site at the time, meaning that two cycles of the Qo site are needed to fully reduce a Q to QH₂ at the Qi site. Overall, the net oxidation of one QH₂ molecule and transfer of two electrons to two molecules of Cytc leads to the uptake of two
protons from the *n* side (mitochondrial matrix or bacterial cytoplasm) and the release of four protons on the *p* side (intermembrane space of mitochondria or bacterial periplasm). A step-wise scheme of the reaction taking place during the Q cycle in *Cytbc*₁ is shown in figure 3.

In the case of *Cytb*<sub>6f</sub>, although details of the Q cycle have been studied less completely and flash-induced reduction of heme *b*<sub>n</sub> separate from that of heme *b*<sub>p</sub> not defined experimentally<sup>40</sup>, the general principles of the Q cycle and proton translocation mechanism are believed by most researchers in the field to be essentially the same as for *Cytbc*₁ (see perspective on this issue in section 3.6.4). This includes the bifurcation reaction taking place at the PQ oxidation site (Q<sub>p</sub> site) with electrons delivered into the high- and low-potential chains. One of the main differences is the presence of an additional cofactor within the low-potential chain. This is an atypical high-spin and relatively high-potential, heme *c*<sub>n</sub> covalently attached to the protein through a single thioether bond<sup>41,42</sup>. The planes of hemes *b*<sub>n</sub> and *c*<sub>n</sub> are oriented perpendicularly and share electrons<sup>43</sup> with each other with the latter being close to the PQ binding niche. Although a role of this heme is still unknown it was proposed that together with heme *b*<sub>n</sub> it forms a pair of cofactors that can deliver two electrons to PQ in order to reduce it to PQH<sub>2</sub> in a virtually one-step event. This idea, however, has not been experimentally validated yet. Because the heme *c*<sub>n</sub> is exposed to the stromal side of the thylakoids, this cofactor could provide an additional entry point for electrons at the level of the Q<sub>n</sub> site, possibly triggering a different Q cycle mechanism, or providing an oxidant for stromal electrons in cyclic electron flow (see below and discussion in section 3.6.5). Another difference between *Cytbc*₁ and *Cytb*<sub>6f</sub>, due to the variation in the composition of the high-potential chain, is the fact that electrons from the 2Fe2S are transferred to heme *f* which further reduces diffusible PC or *Cytc*<sub>6</sub> (instead of *Cytc* interacting with *Cytc*<sub>1</sub>).

<figure><img src="image_7.png" /></figure>
**Figure 3.** A simplified, static scheme, showing step-wise reactions taking place during the catalytic cycle of *Cytbc*₁. (A) Quinone (Q) and Quinol (QH₂) (*yellow hexagon and white hexagon with 2 dots*) bind to the Qᵢ and Q₀ sites, respectively, while oxidized *Cytc* (*oval with white rhombus*) binds to the *Cytc*₁ domain. (B) After binding, QH₂ is oxidized, 2 protons are released (*turquoise arrows*) to the *p* side, one electron is transferred (*red arrow*) to *Cytc* through the high-potential chain, while the second electron goes (*red arrow*), through the low-potential chain, to Q bound at the Qᵢ site. (C) Reduced *Cytc* (*oval with red rhombus*) and Q diffuse out of the respective sites (*black arrows*) while SQᵢ (*blue hexagon with dot*) stays stably bound at the Qᵢ site. (D) Second molecule of QH₂ and oxidized *Cytc* bind to the Q₀ site and *Cytc*₁, respectively. (E) Oxidation of QH₂ takes place as in case B, but the electron from the low potential chain reduces SQᵢ to QH₂, which is associated with uptake of two protons from the *n* side. (F) Q, QH₂ and reduced *Cytc* diffuse out of the Q₀, Qᵢ and *Cytc*₁, respectively, and the cycle starts again. Hemes are shown as rhombuses, the 2Fe2S is shown as yellow-white circles. Lipid bilayer is marked as green lines. For simplicity, only one monomer is shown and the motion of the head domain of iron-sulfur protein (ISP-HD) was not included. The scheme in general is believed to hold true also for *Cytb*₆f with some modifications at the Qₙ site associated with the presence of additional heme cₙ and possible modifications to the electron transfer sequence associated with the involvement of this enzyme in CET (see Fig.4).

The Q cycle originally proposed by Mitchell³² considered a mechanistic link between electron bifurcation at the Q₀ site and electron confurcation at the Qᵢ site. As mentioned above, the bifurcation at the Q₀ site results in one electron delivered to the Qᵢ site, implying that a completion of confurcation at the Qᵢ site requires the functional link with another redox-active enzyme/cofactor cooperating with *Cytbc*₁ and serving as an additional source of electrons for that site. The second electron to complete reduction of SQ at the Qᵢ site was initially proposed to originate from the activity of another dehydrogenase. Further studies revealed that *Cytbc*₁ does not need to be associated with any source of electrons but QH₂ to complete the cycle. Instead, *Cytbc*₁ must oxidize two QH₂ molecules at the Q₀ site to sequentially deliver two electrons to the Qᵢ site ³⁹. On the other hand the idea that the complex interacts with another dehydrogenase could be relevant to account for the possible role of *Cytb*₆f in CET. Indeed, it has been proposed that electrons donated by PSI to the NADPH⁺ / ferredoxin pool could be reinjected into the PQ(H₂) pool by electron transfer to PQ or putative plastosemiquinone (PSQ) at the Qₙ site. In such a case, the catalytic cycle of *Cytb*₆f under conditions favoring CET, could operate according to the original Mitchell Q cycle (Fig. 4).
<figure><img src="image_8.png" /></figure>

Figure 4: Two possible mechanisms of Q cycle in *Cytb*<sub>6f</sub>. (A) The “modified Q cycle”<sup>39</sup> which is similar to *Cytbc*<sub>1</sub> - two electrons are delivered to the Q<sub>n</sub> site from Q<sub>p</sub> site, which requires oxidation of 2 PQH<sub>2</sub> and reduction of 2 PC, per 1 reduced PQ at the Q<sub>n</sub> site. (B) Original Mitchell Q cycle<sup>32</sup> in which one electron delivered to the Q<sub>n</sub> site comes from the PQH<sub>2</sub> oxidation at the Q<sub>p</sub> site and the second electron comes from electron donors at the *n* side (Fd/FNR). According to the original Mitchell mechanism 1P PQH<sub>2</sub>, 1 PC, 1 PQ and 1 *n* side donor are needed to complete the cycle. Color code and symbols as in Fig 3. Copper in PC is marked as blue dot. For simplicity proton paths are omitted and reactions in one monomer are shown while the second monomer is grayed out.

The bifurcation reaction taking place at the Q<sub>o</sub>/Q<sub>p</sub> site is essential for the energetic efficiency of the catalytic cycle, and has long been considered a unique feature of *Cytbc*. However, more recent findings indicate that bifurcation reaction may exist in other systems, such as those utilizing flavins

44

**3.3 The H-shaped electron transfer system in the dimer**

*Cytbc*<sub>1</sub> complexes from mitochondria and bacterial cells have been crystallized and their structures solved to atomic resolution. The mitochondrial *Cytbc*<sub>1</sub> is a symmetrical, oligomeric homodimer, in which each monomer encompasses the three catalytic subunits (*Cytb*, *Cytc*<sub>1</sub> and ISP) surrounded by a periphery of non-redox, supernumerary subunits. The bacterial enzyme is also dimeric and the overall structure of the catalytic subunits as well as the position of the redox cofactors are highly similar to the mitochondrial enzyme (Fig 5). The major difference is a general lack of the supernumerary subunits in the bacterial enzyme.

One unusual aspect of the dimeric structure is that ISP spans the dimer. This ISP subunit is anchored in one monomer by a single transmembrane helix, while the peripheral, hydrophilic domain that contains 2Fe2S (ISP-HD) is located in the other monomer, where it forms part of the quinol
oxidation Q₀ site ²⁰,²¹,⁴⁵. A comparison of the structures of enzymes crystallized in the absence or presence of the Q₀ site inhibitors indicated that the ISP-HD occupies different positions with respect to the other parts of the complex. This suggested that movement of ISP-HD between positions proximal to Cytb and Cytc₁ is necessary to transfer an electron between the Q₀ site and heme c₁ in the c-chain. This hypothesis was confirmed by site-directed mutagenesis studies which demonstrated that such movement was essential for enzyme activity. However, the mechanistic rationale behind the movement is not clear (see section 3.4).

In view of the topographic arrangement of ISP, the functional module at the level of the *Cytbc*₁ monomer consists of the *Cytb* and *Cytc*₁ and ISP linked to them through its ISP-HD, while also interacting with *Cytb* and *Cytc*₁ of the other monomer through its hydrophobic anchor. The same structure feature applies to *Cytb*₆f (see below). ⁴⁶ The crystal structures of *Cytbc*₁ revealed that large distances separate the cofactors from different monomers, except for the two hemes *b*<sub>L</sub> that are positioned close enough to allow electron transfer between the monomers (Fig 5). The capacity of this electron transfer bridge to connect the monomers on a catalytically-relevant timescale was further proven in studies that exploited cross-inactivated forms of the enzyme ⁴⁷. The existence of the heme *b*<sub>L</sub>-*b*<sub>L</sub> electron transfer bridge implicates that cofactor chains in the dimer assemble into an H-shaped electron transfer system that connects the two Q₀ sites with the two Qᵢ sites. Given that the functional module of each of the monomers provides all necessary elements to support the catalytic Q cycle (the *c*- and *b*-chains, the Q₀ and Qᵢ sites), the mechanistic and physiological meaning of the H-shaped electron transfer system remains unclear but intriguing (see section 9 for details).

*Cytb*₆f is also a homodimer. The topographic arrangement of ISP that are intertwined with the dimer and the movement of ISP-HD are conserved, as is the distance between the two hemes *b*<sub>p</sub>. In view of the general similarity of cofactor architecture between *Cytbc*₁ and *Cytb*₆f it is reasonable to assume that the characteristic H-shaped electron transfer system is a shared feature of all *Cytbc*.
<figure><img src="image_9.png" /><figcaption>Figure 5. Crystal structure and spatial arrangement of electron-transfer cofactors in *Cytbc*₁ from *Rhodobacter capsulatus* (PDB ID: 1ZRT ⁴⁸). A) Ribbon structure showing all catalytic subunits in the dimer with the overlaid cofactors. B) Redox-active cofactors in the dimer with edge-to-edge (numbers) and iron-to-iron distances (numbers in brackets).</figcaption></figure>

**3.4 Large-scale movement of ISP-HD**

The large-scale movement of the extrinsic domain of the iron-sulfur protein (ISP-HD) is an inherent part of the catalytic cycle of *Cytbc*₁ and *Cytb*₆f. The movement alternately brings the 2Fe2S either to the position close to the catalytic Q₀/Qₚ site (referred to as the Q₀ position or the b-position) or close to heme c₁/f, (the c-position) separating in time and space the two electron transfers within the high potential chain: 2Fe2S at the b-position exchanges electrons exclusively with quinol bound at the site, while at the c-position it exchanges electrons exclusively with heme c₁/f. This movement was proven obligatory in many studies showing that impairment of the mobility of ISP-HD entails a severe limitation or complete loss of enzymatic activity⁴⁹⁻⁵⁵. This is commonly explained considering
the effect of changing in distance between cofactors on electron transfer rate:⁵⁶ the ISP-HD arrested at a specific position permanently (due to the presence of some inhibitors) or for prolonged period of time (due to specific mutations) cannot efficiently shuttle electrons between the Q₀/Qₚ site and heme c₁/f as specific distances between the cofactors in a frozen configuration are not optimal for rapid electron transfer along the entire cofactor chain. For example, in a configuration with the ISP-HD arrested at the Q₀ site, a large distance between the 2Fe2S and heme c₁ prevents rapid electron transfer between these two cofactors and consequently enzymatic activity is lost.

*First structural indications for ISP-HD movement.* The possibility that conformational changes might be associated with the catalytic reactions involving 2Fe2S was considered before the first crystallographic structures of *Cytbc* became available⁵⁷,⁵⁸. Although the movement itself was not mentioned, a mechanism evoking a “catalytic switch” of 2Fe2S was formulated on the basis of different affinities of MOA-stilbene and UHNQ inhibitors to the Q₀ site that depended on the redox state of 2Fe2S. The concept of ISP-HD movement during the catalytic cycle was born soon after crystallographic structures with resolved ISP became available⁵⁹⁻⁶¹. Early structures of chicken *Cytbc*₁ revealed two conformations of ISP-HD⁶¹, suggesting that it might switch between different positions during the catalytic cycle. This was further substantiated by the mammalian *Cytbc*₁ structures containing all 11 monomer subunits⁶⁰, which clearly showed that ISP-HD can adopt different conformations for which 2Fe2S was found at the b-position, at the c-position or somewhere in between. Further crystallographic and cryo-EM structures of *Cytbc* originating from many different organisms: bacteria ⁴²,⁴⁸,⁶²⁻⁶⁷, chlorophyta⁴¹, fungi⁶⁸⁻⁷², plants⁷³ to avian⁷⁴⁻⁷⁶ and mammalian proteins⁷⁷⁻⁷⁹, including humans⁸⁰, were all consistent with the concept that ISP-HD must undergo a constrained diffusion in order to facilitate electron transfer between the Q₀/Qₚ site and *Cytc*₁/*Cytf*. One recognized exception comes from recent analysis of the structure of *Cyt-bc* in supercomplex with cytochrome c oxidase from *Mycobacterium smegmatis*, which suggests that a globular domain of a diheme *Cytc* (equivalent of *Cytc*₁) might move instead of ISP-HD to support the catalytic cycle⁸¹,⁸²

*Effects of the Q₀/Qₚ site inhibitors.* It is recognized that two types of inhibitors exert opposite effects on the average position of ISP-HD seen in crystals⁸³. The first group of inhibitors includes compounds, such as stigmatellin, famoxadone, *n*HDBT, atovaquone and DBMIB in *Cytb*₆f, which fix ISP-HD at the b-position and thus are classified as *Pf* inhibitors (f- for fixed). The second group includes compounds that do not arrest the ISP-HD at a single b-position, but, quite oppositely, expel ISP-HD from the Q₀/Qₚ site forcing the domain to adopt positions remote from the Q₀/Qₚ site, including the c-position²¹,⁷⁴,⁷⁷,⁸⁴. As they do not prevent the motion of ISP-HD, they are classified as
$P_m$ inhibitors (m- for mobile). Typical inhibitors of this group are myxothiazol, MOA-stilbene and strobilurin derivatives (see section 5).

*Constrained diffusion or allosteric control.* There are several mechanistic aspects of the movement of ISP-HD which remain the subject of discussion<sup>20</sup>. One of the most important and still unsolved issues concerns a fundamental question of whether the motion is simply a stochastic, thermally-activated process or whether it is rather controlled by a particular state or states of the enzyme during the catalytic cycle<sup>21,85-90</sup>. The simplest answer one may consider is that the motion represents a stochastic process of constrained diffusion<sup>51,90-92</sup>, which is fast enough (~80 000 s<sup>-1</sup>)<sup>93</sup> not to limit the overall rate of catalysis<sup>94</sup>. The stochastic diffusion, in contrast to any mechanism that would rely on a specific element or elements of control imposed on the ISP-HD movement, has the clear advantage of not requiring energy expenditure for “information gain” needed to control the cycle. This seems important given that the primary function of the enzyme is to conserve energy.

However, the general difficulty with understanding the high energetic efficiency of bifurcation taking place at the $Q_o/Q_p$ site and associated with its efficient suppression of side reactions (see section 7.4) prompted scientists to look for structural factors responsible for controlled docking/release of the domain from the b-position during the catalysis. This seemed additionally justified by the specific effects of the $P_f$ and $P_m$ inhibitors on modulation of the position of ISP-HD and on interactions of ISP-HD with *Cytb*. None of the available structures succeeded in resolving the position of natural substrates, UQ/PQ or UQH₂/PQH₂, bound at the $Q_o/Q_p$ site²⁰ which left room for interpretation of any observed structural effects in the context of possible effects of the presence of Q or QH₂ at the catalytic site on the ISP-HD motion. Some additional clues have been drawn from MD simulations⁹⁵⁻⁹⁷.

*Structural changes in brief.* Prior to discussions on the mechanisms of the ISP-HD motion, it is necessary to briefly describe key structural elements of ISP. The ISP subunit can be divided into 3 domains⁹⁸: a) ISP-HD harbouring the redox-active [2Fe-2S] Rieske cluster (2Fe2S) b) the membranous anchor, consisting of a single hydrophobic helix (residues $Rh1-38, Bt1-61$), and c) the neck or hinge region (residues $Rh39-48$ or $Bt62-74$) containing highly conservative amino acid sequence⁹⁹ connecting the anchor with the ISP-HD (see Fig. 6). The two ISPs interwind the dimer in such a way that each of the ISP-HD, interacting with $Cytc_1$ or *Cytb* of one monomer, has its anchor associated with *Cytb* of the second monomer (see Fig 5). The movement of ISP-HD is associated with conformational change within the flexible neck region (described below), rotation of the domain by approximately 57 to 65° ⁹²,¹⁰⁰ and displacement of the domain by approximately 2 nm.
<figure><img src="image_10.png" /><figcaption>Figure 6. Structure of *Cytb* and two conformations of ISP in one monomer. (A) Structure based on PDB:1sqx with ISP-HD at the b-position (B) Structure based on PDB:1be3 with ISP-HD being at the c-position. The hydrophobic anchor of ISP (blue helix) is embedded in the lipid membrane (pale green square). In A and B the neck region connecting ISP-HD (green) with the anchor has two different conformations. *Cytb* is shown in gray.</figcaption></figure>

*Different positions of ISP-HD in crystal structures.* Comparing different crystal structures Berry et al.²⁰ proposed that the observed populations of ISP-HD fall into 4 main groups if one considers differences in distances and angles of the position of the 2Fe2S in the structure relative to the stigmatellin bound at the Q₀ site (Fig. 7). The first group encompasses a population of ISP-HD in “b-positions” or “fixed positions” with distances of up to ~0,15 nm and angles up to 7°. The second group gathers “famoxadone-positions” for which the distance is between ~0.15 and ~0.2 nm with the angle ~10°. The third group encompasses “low-affinity fixed positions” at distances ~0.4 to 0.45 nm and angles ~17 to ~20°. The last group considers ISP-HD at the “c-positions” for which angles fall between 55° – 60° and distances ~1.5 to ~2.0 nm. It is of note that the diagram by Berry et al.²⁰ includes a structure originally resolved by Iwata et al.⁶⁰ which was not classified to the group of c-positions and for which distance is ~2.3 nm and angle ~65°. For this structure, the distance between BtH161 (RhH156) of ISP and heme c₁ is short enough to allow the formation of a H-bond between this histidine N<sub>t</sub> atom and the propionate group of heme c₁ indicating the most probable conformation predisposed to fast electron exchange between the cluster and heme c₁.

*Structure-function relationships for the neck region of ISP-HD.* While detailed comparisons of structures revealed several elements that might undergo specific conformational changes upon the motion of ISP-HD, the cause-effect relationships of these elements in the context of the mechanism are not fully understood. The most evident structural change concerns the neck region which, depending on the position of ISP-HD, adopts conformations differing in length²¹,⁵¹. When the ISP-
HD is at the c-position, the neck forms a small helical structure, whereas docking the domain to the Q₀ site coincides with the “melting” of this small helix, thus lengthening the connection between the anchor and the ISP-HD (Fig. 6). However, helix formation was seen upon transition of ISP-HD from the “low-affinity fixed position” (group 3) to b-position (group 1)²⁰.

The first evidence for involvement of the neck region in ISP-HD mobility can be found in studies showing that mutations in the conserved fragment of the Box 1 sequence of the ISP (ISP:RhL136) that destroy sensitivity of 2Fe2S to UQ at the Q₀ site, were mitigated by mutations in the hinge region (ISP:RhV44 or ISP:RhA46)¹⁰,¹⁰². Further mutational studies systematically investigated the effects of changing the length and amino acid composition of the neck region on movement of ISP-HD. Elongating the neck by insertion of 1, 2 or 3 alanine residues (+1Ala, +2Ala, or +3Ala mutants, respectively) was found to decrease overall electron transfer through the high-potential chain and the magnitude of this decrease depended on the length of the insertion. In +1Ala, the electron transfer to heme c was found to occur within the millisecond timescale, while in +2Ala and +3Ala – in the second timescale, which is orders of magnitude slower than in the native enzyme (micro to milliseconds). Apparently ISP-HD movement, which in the native enzyme is fast and not rate-limiting, becomes rate-limiting in the mutants⁵¹,⁹³,¹⁰³,¹⁰⁴.

Interestingly, +1Ala is still functional *in vivo*, while +2Ala and +3Ala are not, indicating that some level of functional tolerance to restrictions of the ISP-HD movement exists Detailed analysis of the kinetic data lead to the proposal that these mutants progressively restrict the movement in such a way that the ISP-HD gets arrested for a prolonged period of time at the b-position (for milliseconds in +1Ala or seconds in +2Ala or +3Ala)⁵⁰,⁵²,⁹⁰,¹⁰⁵⁻¹⁰⁷. Also, replacement of the neck-region residues with 6 prolines⁵⁰,¹⁰⁶ all but abolished helix formation and precluded the release of the domain from the b-position. Surprisingly, shortening of the neck region by 1 to 5 amino acids did not lead to loss of catalysis and the EPR spectra obtained for deletion mutants still reflected the ability of 2Fe2S to interact with the UQ and UQH₂ bound at the Q₀ site, albeit with stoichiometry of ISP to Cytb progressively decreasing to 0.2 in 5-residue-deletion mutant⁵⁰.

The conclusion reached from analysis of kinetic data that the alanine insertions arrest the ISP-HD at the b-position was confirmed by EPR studies with oriented membranes⁸⁸,¹⁰⁰,¹⁰⁸⁻¹¹¹ and pulse EPR with isolated Cytbc₁⁹⁰,¹⁰⁷,¹¹²,¹¹³. The latter studies allowed direct monitoring of changes in the average position of the ISP-HD with respect to heme bL by measuring the distance-dependent enhancement of reduced 2Fe2S relaxation by oxidized heme bL. The average distance between these two cofactors was found to increase in the following order +2Ala mutant, +1Ala mutant, WT¹⁰⁷. These measurements also revealed that the position of ISP-HD in +2Ala is no longer influenced by
$P_m$ inhibitors but remains sensitive to the addition of stigmatellin<sup>90</sup>. This suggests that conformational changes at the neck region can be overwhelmed by the energy of stigmatellin binding, with a large contribution coming from H-bond formation (typical value of energy required to break an H-bond is ~21 kJ/mol)<sup>20</sup>.

*Structure-function relationships for the interaction of ISP-HD with Cytb.* Given that the $P_f$ and $P_m$ inhibitors exert different effects on the position of ISP-HD the question arises as to how the $P_f$ inhibitors fix the domain at a position close to the $Q_o$ site. One of the most obvious structural effects of the presence of stigmatellin, *n*HDBT, atovaquone, crocacin D and ascochlorin (group-1 of $P_f$ inhibitors) at the $Q_o$ site is formation of the H-bond between $RhH156$ ($BtH161$ or $ScH181$) of the ISP subunit and the inhibitor. Formation of this bond is not only associated with docking of ISP-HD to the $Q_o$ site, but also stabilizes the reduced state of 2Fe2S by significantly raising its redox potential<sup>114-116</sup>. This suggests stabilization of the reduced cluster under conditions when it forms the H-bond with a molecule occupying the $Q_o$ site. Indeed, the observation that the reduced cluster is detected by EPR at the b-position in membranes indicates that interactions of the reduced 2Fe2S and the natural quinone at the $Q_o$ site are also stabilized. On the other hand ISP-HD appears to favor the c-position when the cluster is oxidized<sup>100</sup>. This difference in affinity to quinones at the $Q_o$ site between the reduced and oxidized cluster may be one of the elements contributing to the mechanism that diminishes the risk of energy-wasting short-circuits (see section 7.4.1). It should be noted however, that some inhibitors (famoxadone, fenamidione or JG-144 inhibitors, all belonging to the group 2 of $P_f$ inhibitors) hold the domain at the b-position without creating an H-bond to $RhH156/BtH161$<sup>117</sup>. This indicates that factors other than the H-bond between $RhH156/BtH161$ of ISP and the occupant of the $Q_o$ site can also stabilize binding of ISP-HD to Cytb. While formation of this bond would be expected to significantly strengthen the interaction, it is clearly not an absolute requirement for the domain fixation to Cytb or $Cytb_6$.

It seems reasonable to assume that some changes in conformation on the surface of Cytb or $Cytb_6$ contribute to the process of fixing to or release of the domain from the b-position<sup>20,21,78,85</sup>. The potential “holding and release lever”, if it exists, would be expected to localize to the part of the Cytb/b<sub>6</sub> which remains in close contact with the surface of ISP-HD. These regions encompass small helices that are connected to the transmembrane ones by loops: *cd1*, *cd2* and *ef* loop<sup>18,20,55,64,118</sup>. However, when structures containing $P_f$ and $P_m$ inhibitors are compared, there is no significant change in the position of helices in Cytb, except for the *cd1* helix, which seems to be slightly pushed inside Cytb by ISP-HD upon interaction of stigmatellin with 2Fe2S<sup>20</sup>. Furthermore, a comparison of the structures of intermediate positions (group 3) and the c-position (group 4) reveals no changes in the
*cd1* and *cd2* helices, suggesting that the motion of ISP-HD between “low-affinity fixed positions” (group 3) and the c-position (group 4) is a free diffusional process constrained by the neck region of the ISP acting as a peptide tether.

It is not clear if the shift of the *cd1* helix upon docking ISP-HD at the Q₀ site is an effect of the inhibitor itself, allowing the 2Fe2S cluster to “come closer” or if the H-bond between the inhibitor and $^{Rh}$H156 induces “pressure” of ISP-HD on the structure forcing *cd1* to move. Such a pressure could be an element of the “spring-loaded” mechanism which proposes that H-bond formation between ISP-HD and a substrate present in the Q₀ site stores some energy which is released after the reaction and facilitates the dissociation of ISP-HD from the b-position¹¹⁹.

The involvement of the *cd1* helix in binding of ISP-HD to the Q₀ site is also implicated by mutational studies in which $^{Rh}$G167, located at the end of the *cd1* and before the *cd2* helix, was replaced by proline. This significantly perturbed the docking of ISP-HD to *Cytb*, which was reflected in an observed shift in the equilibrium distribution of the ISP-HD toward the c-position ¹²⁰,¹²¹. As a result, the interaction of 2Fe2S with UQ or UQH₂ at the Q₀ site was broken and the enzymatic activity of the $^{Rh}$G167P mutant dramatically decreased, abolishing the functionality of the enzyme *in vivo*. Also superoxide generation by the Q₀ site increased in this mutant (see section 7.5 for details). Despite these effects, the mutant remained sensitive to stigmatellin. Interestingly, addition of the +1Ala or +2Ala mutation to the $^{Rh}$G167P mutant was found to mitigate the inhibitory effects of $^{Rh}$G167P mutation to various extent. A remarkable example was the double $^{Rh}$G167P/+2Ala mutant in which the equilibrium distribution of ISP-HD was similar to that of single mutant +1Ala mutant, partially restoring sensitivity of the reduced 2Fe2S to UQ at the Q₀ site. This brought back some level of functionality to the enzyme *in vivo* even though both single mutations were non-functional. Clearly, the opposite effects of $^{Rh}$G167P mutation (“pushing” ISP-HD away from the b-position) and the alanine insertions (arresting ISP-HD at the b-position) can partly compensate one another, establishing that dynamics of the docking/release of ISP-HD to/from cytochrome *Cytb* can be effectively coinfluenced by the specific interactions at the ISP-HD-*Cytb* interface and the conformational changes taking place within the neck region. A functional link between these two distinct structural domains was also documented in other mutational studies¹²².

Another eye-catching structural element which may somehow influence the ISP-HD movement is the region of the *ef* loop – (*Cytb*: $^{Rh}$286-292, $^{Bt}$262-268) a small peptide fragment connecting the transmembrane E helix, the conserved PEWY sequence and the transmembrane F helix²⁰,⁵⁵,⁶⁰. It is considered to be the main “obstacle” that forms a barrier to ISP-HD on its trajectory from the b- to c-position. The role of the *ef* loop was proposed following the finding that mutations
in this region can compensate the effect of +1Ala<sup>51,53</sup>. Further mutational studies revealed that bulky
side chains located in the middle part of the loop limit the ability of ISP-HD to move outside the Qo
site<sup>122</sup>. This leads to the proposal that the *ef* loop acts as a switch increasing or decreasing the rate of
electron transfer between the Qo site and heme c<sub>1</sub><sup>55,103</sup>, which together with the *cd1* helix constitutes
the mechanism of binding and release of ISP-HD depending on the redox state of hemes b<sup>78,86,88,89</sup>.
However, as demonstrated by mutational studies, amino acid side chains such as <sup>Rh</sup>L286, <sup>Rh</sup>I292 on
the *ef* loop in fact may serve to resist in transitions between *b*- and *c*-position<sup>55</sup>. The control
mechanisms steering this loop during the catalytic cycle is a matter of discussion. The observed effect
of modifications on the motion caused by mutations in the *ef* loop does not preclude a simple
stochastic model for the movement of ISP-HD in which the mobile loop regions undergo random
thermal fluctuations rather than specifically-controlled motion.

Not only the redox state of hemes *b*, but also specific events taking place at the Q<sub>i</sub> site have
been considered as potential factors that influence the mobility of ISP-HD. In view of topographical
arrangement of ISP-HD in the dimer, this would imply a highly ordered communication between
monomers which would allow transfer of information over ~ 4 nm across Cytb, possibly through the
membranous anchor of ISP that ends in proximity to the Q<sub>i</sub> site. Most of the proposals on this type of
allosteric influence are derived from analysis of the effects of antimycin bound at the Q<sub>i</sub> site.
However, as we discuss below, the interpretation of these effects needs to be treated with caution.

*Effect of the Q*<sub>*i*</sub>* site inhibitor, antimycin*. Evaluating the influence of antimycin on the basis of crystallographic structures containing antimycin bound at the Q<sub>*i*</sub> site is difficult as the majority of the antimycin-containing structures were obtained with stigmatellin bound at the Q<sub>o</sub> site, which overrides any possible structural effects of antimycin on the Q<sub>o</sub> site. To our knowledge, until now, there are two PDB crystallographic structures in the database, containing antimycin bound at the Q<sub>*i*</sub> and no other inhibitor at the Q<sub>o</sub> site<sup>117,123</sup>. As discussed by Berry and Huang (ibidem), antimycin does not seem to change the ground state of the structure but might change dynamics of the structure<sup>117</sup>. The dynamic change was considered to accommodate several biochemical studies suggesting that antimycin induces effects that might be associated with changes to the different mobility of the domain.

The first series of experiments was based on the susceptibility to proteolytic cleavage of the
neck region (between residues <sup>Rh</sup>46 and 47) as a sensor of conformational changes taking place in
this region. It was found that the cleavage was substantially limited by fixing of ISP-HD at b-position
by P<sub>f</sub> inhibitors, when compared to the enzyme with P<sub>m</sub> or without any bound inhibitors<sup>124</sup>. On the
other hand addition of antimycin increased proteolysis, suggesting that this inhibitor induced changes
that increased accessibility of this fragment for thermolysin.
The second series of experiments focused on various EPR methods to monitor the FeS.
Continuous wave (CW) EPR spectra of microscopically disordered samples containing antimycin- or
HQNO-supplemented membranes did not show any significant changes in the spectra when compared
to the non-inhibited samples¹¹¹. However, analysis of the samples containing ordered layers of
membranes revealed that after addition of antimycin, the angular-dependent amplitudes of gₓ and gᵧ
transitions of the 2Fe2S cluster are different from those detected in samples not containing
antimycin⁸⁸,¹¹¹. Pulse EPR measurements with isolated Cytc*bc*₁ showed that the presence of antimycin
at the Qᵢ site caused an increase in distance between the reduced 2Fe2S and oxidized heme b*L*⁹⁰ and
an increase in the efficiency of spin-lattice relaxation of the 2Fe2S cluster *via* two-phonon Raman
process. All recognized changes associated with the presence of antimycin at the Qᵢ site consistently
indicate that this inhibitor induces some structural changes that modify the average position of ISP-
HD. This effect was interpreted by some authors as evidence for allosteric inter- and/or intra-
monomer interactions between the Qᵢ/Q*n* and Q₀/Qp site that lead to specific sequences of a "duty
cycle" of Cytc*bc*₁/b*6f* during the catalysis⁸⁶⁻⁸⁸,¹²⁵,¹²⁶.

In our view, the observed effects of antimycin on the ISP-HD are associated with a decrease
in the rigidity of the Cytc*bc*₁ structure which decreases the energetic barrier for moving from b- to c-
positions. This simple concept assumes that the population of ISP-HD domains is distributed over
different conformations and various inhibitors, including antimycin, just modify a barrier for this
movement⁹⁰. In fact, pulse EPR measurements revealed that the average position of ISP-HD can be
effectively modified by mutations and/or inhibitors and the observed shifts in this position can be
placed in the following order (starting from b-position and going further toward the c-position): +2Ala
mutant > stigmatellin > Cytb:RhG167P/ISP:+2Ala double mutant > +1Ala mutant >
Cytb:RhG167P/ISP:+1Ala double mutant > no-inhibitor WT > WT + myxothiazol ≈ Cytb:RhG167P ≈
WT + antimycin > WT + antimycin + myxotiazol⁴⁷,⁹⁰,¹²⁰. In all cases, addition of stigmatellin
 overrides any other effect, which suggest that the energy of H-bonding between the inhibitor and
 ISP*RhH156* exceeds the energy associated with changing the structure of the neck region. Although
 stigmatellin is able to overcome the effects of alanine insertion into the neck region, neither Pm
 inhibitors (myxotiazol) nor antimycin influence the ISP-HD position in +2Ala mutations⁹⁰. Gathering
 all these observations we can propose that the strength of factors that influence the ISP-HD mobility
can be put in the following order: Pf inhibitors which form H-bonds > conformation of cd1 helix
(RhG160P mutation) > increase in the length of the neck region > Pm and Qᵢ inhibitors (myxothiazol,
antimycin). The *ef* loop was not included in this list since no direct spectroscopic measurements of
distance distribution of ISP-HD were done for mutants in this particular region. However, comparing
the kinetic data and the effect of the redox potential of 2Fe2S, it may be suggested that the energetic
barrier of crossing the *ef* loop by ISP-HD is lower than the barrier imposed by +2Ala, comparable
with the effect of +1Ala insertion and higher than the barrier imposed by P<sub>m</sub>-type or Q<sub>i</sub> inhibitors<sup>53,55</sup>.

We note that most of the structural studies involving EPR spectroscopy have been performed under conditions for which 2Fe2S was reduced. Although no changes on the surface of the ISP-HD associated with the changes in the redox state of the cluster, were observed<sup>20</sup>, the domain with a reduced 2Fe2S appear to show a greater tendency to occupy the b-position compared to the domains with the oxidized 2Fe2S. This is in line with the observation that E<sub>m</sub> of 2Fe2S depends on the position of ISP-HD<sup>106</sup>.

*Effect of movement on redox potential of the 2Fe2S cluster.* There are numerous studies showing that the *E*<sub>m</sub> of 2Fe2S depends on interaction between ISP-HD with the Q<sub>o</sub> site <sup>50,52,53,106,109,113,114,116</sup>. In general, the greater the tendency for ISP-HD to be at the b-position, the more positive the redox potential of 2Fe2S. For the mutants +1Ala, +2Ala and 6Pro, that were discussed above, the *E*<sub>m</sub> values of 2Fe2S were found to increase by the same order. This could suggest decreased in water accessibility, which changes pK of <sup>Rh</sup>H156 (<sup>Bt</sup>H161). However, the position of the ISP-HD is not alone in influencing the redox potential of 2Fe2S. Another important factor is the interaction of the domain with the occupant of the Q<sub>o</sub> site. The most prominent effect is formation of a H-bond with stigmatellin, which increases the midpoint potential by more than 200 mV. Other inhibitors that form a H bond with <sup>Rh</sup>H156 of the ISP-HD, such as tridecylstigmatellin or UHDBT, exert a weaker effect on the rise in the redox potential<sup>106</sup>. Also, the presence of Q bound at the Q<sub>o</sub> site appears to slightly contribute to a slight increase in the redox midpoint potential of 2Fe2S<sup>109</sup>. However, it should be noted that the effect of Q bound at the Q<sub>o</sub> site has been recognized in +nAla mutant but not in WT<sup>109</sup> (see table I in ref <sup>127</sup>). On the other hand, P<sub>m</sub> inhibitors such as myxotiazol or MOA-stilebene slightly decrease the *E*<sub>m</sub> value of the 2Fe2S<sup>128</sup>. Thus, it was proposed that H-bonding is the factor which controls the redox potential and stabilizes binding of substrate and enzyme<sup>128</sup>.
<figure><img src="image_11.png" /></figure>

Figure 7. Several crystal structures of *Cytbc*₁ monomers with different conformations of ISP-HD. The figure shows positions of 2Fe2S clusters (yellow and orange spheres) from the overlaid positions from bovine (PDB IDs: 1sqx, 1bgy, and 1be3), mouse (PDB ID 3cx5) and chicken (PDB ID: 1bcc) crystallographic structures of ISP. Blue, ochre and red ribbons represent ISP, *Cytb* and *Cytc*₁, respectively. Hemes are shown as red sticks. Green sticks show the stigmatellin from 1sxq structure bound at the Q₀ site (yellow oval).

**3.5 Cytochrome **bc**1 – specific structural and functional elements**

**3.5.1 Subunit composition of Cytochrome bc₁**

All Cyts *bc*₁ contain three protein subunits per monomer with redox prosthetic groups: the diheme *Cytb*, the monoheme *Cytc*₁ and the ISP (Rieske iron-sulfur protein). *Cytb* is built of 8 transmembrane helices and contains a relatively high-potential heme *b*<sub>H</sub> and a lower potential heme *b*<sub>L</sub>. *Cytc*₁ and the ISP are membrane-anchored and their water-soluble, globular domains contain prosthetic groups (high-potential heme *c*₁ and 2Fe2S, respectively). All these three subunits are necessary for the catalytic function of *Cytbc*₁. In some bacteria, such as *Paracoccus denitrificans*, *Rhodospirillum rubrum*, or *Rhodobacter* (R.) *capsulatus*, *Cytbc*₁ contains only these three subunits. Other bacteria, including *R. sphaeroides*, contain a fourth subunit of unknown function that lacks prosthetic groups. <sup>129</sup>

In addition to the catalytic subunits, Cytsbc1 of mitochondria contain as many as seven (in some yeast) or eight (in bovine and human) supernumerary subunits. The largest supernumerary subunits associate to the catalytic subunits from the mitochondrial matrix side. The functions of the supernumerary subunits are not known, expect for the established mitochondrial processing peptidase
activity of large subunits in plants. The supernumerary subunits are believed to contribute to the
increased stability of mitochondrial complexes, but are not required for the electron transfer and
proton translocation activities of the enzyme; the three-subunit bacterial enzymes have the same
electron transfer and proton translocation functionality as the mitochondrial enzymes.

**3.5.2 Structure and spectral properties of hemes bL, bH, c1 and 2Fe2S cluster.**

The redox-active components of Cytbc₁ in all organisms are evolutionary conserved within the catalytic core.¹ Remarkably, as shown by the crystal structures of Cytbc₁ from various organisms, the positions of the stationary cofactors, hemes *b* and *c*₁, are unchanged among them.²¹,⁴⁸,⁶⁰

The two *b*-type hemes, (heme *b*<sub>L</sub> and *b*<sub>H</sub>) of Cyt*b* subunit are embedded within the four-helix-bundle, assembling in what appears to be a common (evolutionary conserved) structural motif of many bioenergetic complexes.<sup>130-132</sup> In the Cyt*b* subunit, helices named A-D are involved in bundle formation (Fig. 8 A), with helices B and D providing axial ligands for the heme iron of both hemes (Fig. 8 A,B).<sup>133</sup> Both hemes *b* are axially coordinated by the two conserved histidines (referred to as a bis-His ligation pattern), via bonds between the heme iron and Nε atom of the respective imidazole ring (Fig. 8 B).<sup>48,132</sup>

The heme c₁, is a c-type heme attached to the globular head of Cytc₁ (Fig. 8 C). It is covalently bound to the apoprotein via thioether linkers formed by its two vinyl side chains and two cysteine residues⁴⁸,⁶⁰, belonging to a characteristic CXXCH motif.¹³⁴ The heme iron of heme c₁ is axially coordinated by Nε of the conserved histidine from this motif and Sδ of the conserved methionine (referred to as a His-Met ligation pattern).⁴⁸,¹³⁵ Interestingly, in some bacterial species, the ligation of heme c₁ is secured by an additional pair of cysteines, that form a disulfide bond to stabilize the distant loop of the globular part of Cytc₁.¹³⁵ However, the freedom retained for conformational rearrangement results in a partial weakening of the methionine ligation when the cytochrome is in its oxidized form, which is reflected in an ability to bind small exogenous ligands, such as cyanide.¹³⁶ Removing the disulfide by mutations was found to significantly perturb the conformational stability and further weaken the methionine ligation. As a result, the potential of the heme dropped dramatically (more than 300 mV) and the reduced cytochrome showed a capability to bind carbon monoxide, all of which imply change in the heme ligation pattern¹³⁵. In other Cytochromes c₁, such as mitochondrial Cytc₁, the disulfide is not present. Instead, the βXM structural motif (where β stands for the β-branched amino acid located one residue away from the methionine axial ligand) appears to restrict the conformational flexibility of the domain of the methionine axial ligand to the point that
the binding of external ligands is not observed.¹³⁶ Some bacterial Cytsc₁ have both the disulfide and the βXM motif.¹³⁵,¹³⁷

Similarly to the heme c₁, 2Fe2S is attached to ISP-HD and situated between the two loops of the protein backbone, that are hooked by a disulfide bridge formed between two cysteine residues.¹³⁸ The cluster is coordinated by the thiolate side chains of the two conserved cysteine residues, acting as ligands for one Fe ion, and Nδ atoms of the two conserved histidine residues, acting as ligands for second Fe ion (Fig. 8 D) ⁴⁸,¹³⁸. This type of [2Fe-2S] cluster coordination by two Cys and two His residues is almost exclusive to ISP of Cytbc and aromatic ring-hydroxylating monooxygenases and dioxygenases.¹³⁹,¹⁴⁰

<figure><img src="image_12.png" /></figure>

**Figure 8.** Structural features and binding motifs of metallic cofactors of Cytbc₁. A) Schematic representation of the transmembrane part of Cytb, showing helices involved in formation of the heme-bearing 4-helix bundle (A-D, pale orange rectangles). Hemes are shown as orange polyhedrons, positions of axial ligands of hemes are marked by black dots, *R. capsulatus* residue numbering is used. B) Structural model showing hemes *b* orientation within the transmembrane bundle and orientation of their axial ligands. Helices providing axial ligands are shown in pale orange cartoon, ligating histidines are colored orange. C) Structural model showing heme c₁ binding site in the globular domain of Cytc₁. Cysteines of the binding motif are colored light blue, axial ligands are colored pale green. D) Structural model showing the [2Fe-2S] cluster binding site in ISP. Coordinating residues are labeled in black, cysteines that form the disulfide bridge are colored and labeled in green. All representations are based on PDB 1ZRT entry.

Iron atoms in all the hemes of Cytbc₁ are hexa-coordinated and have strong axial ligands (with bis-His and His-Met ligation pattern for hemes *b* and heme c₁, respectively). Therefore, they all exist
in low spin states. In the reduced (ferrous) state, the hemes exhibit partially overlapping α/β absorbance bands in the visible region of the UV-VIS spectrum (Fig. 9 A), with α-band maxima at 560-562 and 552-553 nm for hemes b and c₁, respectively ¹⁴¹⁻¹⁴⁴. The spectral resolution of heme bH and heme bL by optical spectroscopy is difficult. In the bacterial Cytbc₁ a deconvolution of the spectra obtained upon redox titration implicates a dominant contribution from the heme bH at 562 nm and a significantly smaller contribution from heme bL, which has a split α-peak spectrum with maxima at 564-566 and ~558 nm.³⁹,¹⁴⁵,¹⁴⁶ On the other hand, the heme c₁, due to its much higher redox midpoint potential and α-band maximum at a lower wavelength, can be easily isolated spectrally from hemes b in the partially-reduced enzyme (Fig. 9 A, in orange).¹⁴¹ In their oxidized (ferric) state, hemes b and c₁ are paramagnetic (S = ½)¹⁴⁷, thus detectable by electron paramagnetic resonance (EPR) spectroscopy at low temperatures (<20 K). All three hemes fall into the HALS (Highly Axial/Anisotropic Low Spin) hemes category, with only their gz (often called gmax) transition being observable by EPR. ¹³² In general, the bis-His coordinated hemes b exhibit a HALS signal when the imidazole rings of the two His residues are nearly perpendicular to each other.¹⁴⁸ In the case of hemes bL and bH the angles between the two imidazole ring planes are ~84° and ~58°, respectively¹³², which translates into the two separate EPR transitions at g = 3.78 and g = 3.44, respectively (Fig. 9 B)¹⁴⁹,¹⁵⁰. Thus, EPR, unlike optical spectroscopy, offers a full spectral separation of the two hemes b. It is of note that the gz transition of heme bH is sensitive to Qᵢ site occupancy: when the site is occupied by antimycin A (a specific Qᵢ site inhibitor) or semiquinone radical (SQᵢ), the gz value is shifted to g = 3.47 or 3.42, respectively (Fig. 9 B)¹⁴⁹,¹⁵¹. In R. capsulatus Cytbc₁, the gz transition of heme c₁ occurs at g = 3.16.¹⁵² The exact g-values for gz transitions of each heme may differ among organisms. Some of them were included in Table 1.

<table><thead><tr><th rowspan="2">organism<br>(reference)</th><th colspan="3">gz value (concerns Cytbc<sub>1</sub>)</th></tr><tr><th>heme b<sub>L</sub></th><th>heme b<sub>H</sub></th><th>heme c<sub>1</sub></th></tr></thead><tbody><tr><td colspan="4"><em>Rhodobacter capsulatus</em></td></tr><tr><td>153</td><td>3.78</td><td>3.45</td><td>n.d.</td></tr><tr><td>151</td><td>3.78</td><td>3.44<br>(3.47 / antimycin)<br>(3.42 / SQ<sub>i</sub>)</td><td>n.d.</td></tr><tr><td>152, 150</td><td>3.78</td><td>3.44</td><td>3.16</td></tr><tr><td colspan="4"><em>Rhodobacter sphaeroides</em></td></tr><tr><td>153</td><td>3.78</td><td>3.49</td><td>3.36</td></tr><tr><td colspan="4"><em>Saccharomyces cerevisiae</em></td></tr><tr><td>141</td><td>3.76</td><td>3.60</td><td>3.49</td></tr><tr><td colspan="4"><em>Bos taurus</em></td></tr><tr><td>155</td><td>3.79</td><td>3.43</td><td>3.37</td></tr></tbody></table>
<table><caption>Table 1. g<sub>z</sub> values measured for hemes in selected Cytbc<sub>1</sub> variants.</caption><tr><td>153</td><td>3.78</td><td>3.45</td><td>3.35</td></tr><tr><td>154</td><td>3.78</td><td>3.44</td><td>n.d.</td></tr><tr><td colspan="4">(3.47 / antimycin)</td></tr><tr><td>Aquifex aeolicus</td><td></td><td></td><td></td></tr><tr><td>156</td><td>3.70</td><td>3.45</td><td>3.55</td></tr></table>

It is difficult to use optical spectroscopy in studies on the 2Fe2S, as it has no prominent spectrum<sup>157</sup> and its transitions are dominated by the spectra of hemes. The one-electron reduced form ([2Fe-2S]<sup>1+</sup>) is paramagnetic (due to its ground state with S = ½ resulting from the antiferromagnetic spin coupling between Fe<sup>3+</sup> (S = 5/2) and Fe<sup>2+</sup> (S = 2) ions) and can only be detected by EPR at temperatures <50K. The fully oxidized ([2Fe-2S]<sup>2+</sup>) cluster is in the EPR-silent (S = 0) ground state (the spins of the two Fe<sup>3+</sup> ions are spin-coupled).<sup>140,158</sup> The EPR spectrum of the reduced cluster ([2Fe-2S]<sup>1+</sup>) exhibits a unique rhombic symmetry (Fig. 9 C), with transitions at g<sub>z</sub> = 2.03, g<sub>y</sub> = 1.90 and g<sub>x</sub> between 1.78 and 1.76, resulting in the average g value of approximately 1.9.<sup>157</sup> In bacterial Cytbc<sub>1</sub>, the shape and the position of the g<sub>x</sub> transition was found to be sensitive to the occupant of the Q<sub>o</sub> site (whether it is occupied by substrate, or inhibitor or remains empty), the redox state of quinone (whether quinone or quinol are present) and the size of the quinone pool/potential number or quinone molecules at the Q<sub>o</sub> site (see section 7.2). For those reasons, EPR analysis of the cluster has proven to be a convenient spectroscopic tool for monitoring the status of the Q<sub>o</sub> site.<sup>50,159</sup>

Midpoint redox potentials (E<sub>m</sub>) of hemes b, heme c<sub>1</sub> and 2Fe2S may vary in different systems, depending on the quinone derivatives employed by the organism to serve as electron/proton carriers. In organisms using menaquinones, they are usually lower than in organisms that employ ubiquinones.<sup>160</sup> However, a general rule can be drawn, that the hemes b have much lower E<sub>m</sub>'s than 2Fe2S and heme c<sub>1</sub>. For this reason, the chain of cofactors built by hemes b is referred to as a low-potential chain, while that formed by the 2Fe2S and heme c<sub>1</sub> – high-potential chain. The large gap in potential between components of the high- and low-potential chains is crucial for electron bifurcation and efficiency of the catalytic cycle (see section 7.4). Among the hemes b, heme b<sub>L</sub> has lower E<sub>m</sub> (-290 to -50 mV) than heme b<sub>H</sub> (-160 to 100 mV).<sup>21,160</sup> This difference in E<sub>m</sub>'s of the hemes b is evolutionary conserved.<sup>149,160</sup> It is generally assumed to facilitate cross-membrane electron transfer and assist in overcoming the barrier of membrane potential.<sup>161,162</sup> However, such a function becomes less obvious in light of recent studies demonstrating that the difference in E<sub>m</sub>'s of the hemes b is not required to support the *in vivo* and *in vitro* functionality of the enzyme.<sup>149</sup> Interestingly, E<sub>m</sub> of heme b<sub>H</sub> depends on the occupant (and the redox state) of the Q<sub>i</sub> site. When the Q<sub>i</sub> site is occupied by Q, heme b<sub>H</sub> has a higher potential than when the site is occupied by SQ<sub>i</sub>. This results in a biphasic titration
of the heme.¹⁶³ However, when the Qᵢ site is occupied by an inhibitor, the bH titrates as a single species.¹⁶³,¹⁶⁴ The 2Fe2S and the heme c₁ usually have close values of Eₘ, both falling in range of 100 to 350 mV.²¹,¹⁶⁰ Some of the known values of Eₘ for each of cofactors in various organisms were included in Table 2.

<figure><img src="image_13.png" /><figcaption>**Figure 9. Spectral features of Cytbc₁ redox cofactors.** A) Optical spectra of α/β-band region of hemes. Spectrum of the fully oxidized enzyme is shown in black, spectrum after reduction with sodium ascorbate is shown in orange (showing prominent contribution from heme c₁) and spectrum of the fully reduced enzyme (with dithionite) is shown in magenta. B) EPR spectra of gₓ transitions of hemes bL and bH in an air-oxidized sample (top), sample of the enzyme inhibited with antimycin A (middle) and a sample with generated SQᵢ (bottom). Numbers indicate g values. C) EPR spectrum of a one-electron reduced [2Fe-2S] cluster in isolated Cytbc₁.</figcaption></figure>

<table><thead><tr><th rowspan="2">Organism<br>(reference), [pH]</th><th rowspan="2">[2Fe2S]</th><th colspan="3">E<sub>m</sub> value [mV] (concerns Cytbc<sub>1</sub>)</th></tr><tr><th>heme c<sub>1</sub></th><th>heme b<sub>L</sub></th><th>heme b<sub>H</sub><sup>*</sup></th></tr></thead></table>
<table><caption>Table 2. E<sub>m</sub> values of redox cofactors in selected Cytbc<sub>1</sub> variants.</caption><tbody><tr><td colspan="4"><strong>Rhodobacter capsulatus</strong></td></tr><tr><td><sup>150</sup>, [7.0]</td><td>+295</td><td>-138</td><td>+43, +134</td></tr><tr><td><sup>163</sup>, [7.0]</td><td></td><td>-137</td><td>+35, +202</td></tr><tr><td><sup>104</sup>, [7.0]</td><td>+335</td><td></td><td></td></tr><tr><td><sup>135</sup>, [7.0]</td><td>+320</td><td></td><td></td></tr><tr><td><sup>51</sup>, [7.0]</td><td>+310</td><td></td><td></td></tr><tr><td colspan="4"><strong>Rhodobacter sphaeroides</strong></td></tr><tr><td><sup>143</sup>, [7.0]</td><td>+280/+325</td><td>+240</td><td>-100</td><td>+48</td></tr><tr><td><sup>170</sup>, [7.0]</td><td></td><td>+237</td><td>-87</td><td>+41</td></tr><tr><td colspan="4"><strong>Rhodospirillum rubrum</strong></td></tr><tr><td><sup>142</sup>, [7.4]</td><td></td><td>+320</td><td>-90</td><td>-30</td></tr><tr><td colspan="4"><strong>Paracoccus denitrificans</strong></td></tr><tr><td><sup>165</sup>, [n.d.]</td><td>+265</td><td>+190</td><td>-95</td><td>+30, +120</td></tr><tr><td colspan="4"><strong>Aquifex aeolicus</strong></td></tr><tr><td><sup>156</sup>, [7.0]</td><td>+210</td><td>+160</td><td>-190</td><td>-60</td></tr><tr><td colspan="4"><strong>Saccharomyces cerevisiae</strong></td></tr><tr><td><sup>166,167</sup>, [7.0]</td><td>+285</td><td></td><td>-60</td><td>+82</td></tr><tr><td><sup>168</sup>, [7.4]</td><td>+286</td><td>+270</td><td>-20</td><td>+62</td></tr><tr><td colspan="4"><strong>Bos taurus</strong></td></tr><tr><td><sup>21</sup>, [7.0]</td><td>+238</td><td>-83, -60</td><td>+26, +85</td></tr><tr><td><sup>169</sup>, [7.0]</td><td>+230</td><td>-30</td><td>+100</td></tr></tbody></table>

## 3.6 Cytochrome b<sub>6</sub>f–specific structural and functional elements

The discussion of structure-function of Cytb<sub>6</sub>f is based on a high resolution (2.50 Å) crystal structure (PDB ID: 4OGQ) of the complex. The complex consists of a 250 kDa hetero-oligomeric intra-membrane lipo-protein complex containing, per monomer, thirteen transmembrane α-helices and seven prosthetic groups, the latter consisting of five hemes, one chlorophyll a molecule and one β-carotene. In addition, each monomer contains 23 lipid binding sites, the function of which involves a new area of research into the structure-function of intra-membrane proteins, both for Cytb<sub>6</sub>f and the entire field of membrane proteins.

*Comparison of Cytochromes f and c<sub>1</sub>.* In contrast to the conserved nature of Cytb, for the Cytc<sub>1</sub> and Cytf, except for the single transmembrane helix (TMH) which binds the subunit to the membrane, the structures are different. The only conserved sequence segment is the CXXCH motif, responsible for covalent binding of the c-type heme. The p side heme binding domain has a completely different secondary structure, β sheet and α-helical, respectively in Cytf and Cytc<sub>1</sub><sup>171,172</sup>. In fact, all three of the p side electron transfer proteins which are part of Cytb<sub>6</sub>f, or interact with it, the ISP, Cytf, and PC,

<footnote>* Two values of E<sub>m</sub> for heme b<sub>H</sub> results from two-component fit to data obtained for Cytbc<sub>1</sub> without antimycin.</footnote>
which mediate electron transfer between the quinol hydrogen donor and the P700 PSI reaction center are predominantly in the β-conformation. Both Cytf and Cytc₁ have a relatively positive potential, + 370 mV and + 260 mV, respectively, sufficient to provide an oxidant for the ISP.

### 3.6.1 Subunit composition of Cytochrome *b*<sub>6</sub>*f*

Crystal structures have been obtained for the *Cytb*<sub>6</sub>*f* from the green alga, *C. reinhardtii*<sup>41</sup> and cyanobacteria<sup>42,46,65–67,173,174</sup>.

The crystal structures of the mitochondrial and bacterial *Cytbc*₁ contain 11 and 3 or 4 polypeptide subunits, respectively, compared to 8 in the *Cytb*<sub>6</sub>*f* from cyanobacteria<sup>42,174</sup> and the green alga, *C. reinhardtii*<sup>41</sup>. This subunit count refers to those which remain after the purification and crystallization procedures, and does not include relatively weakly bound peptides which may be important for function but are lost during purification and crystallization. Molecular weights (MW) and pI values for the 8 core subunits of *Cytb*<sub>6</sub>*f* are summarized in table **错误! Source du renvoi introuvable.**

<table><thead><tr><th>Subunit</th><th>MW (kDa)*</th><th>pI</th></tr></thead><tbody><tr><td>Cytf<sup>a</sup></td><td>30.9</td><td>4.7</td></tr><tr><td>Cytb<sub>6</sub><sup>a</sup></td><td>24.7</td><td>8.5</td></tr><tr><td>ISP<sup>a</sup></td><td>19.3</td><td>5.3</td></tr><tr><td>SubIV<sup>a</sup></td><td>17.5</td><td>7.8</td></tr><tr><td>Pet G<sup>a</sup></td><td>3.9</td><td>8.1</td></tr><tr><td>Pet L<sup>a</sup></td><td>3.2</td><td>9.9</td></tr><tr><td>Pet M<sup>a</sup></td><td>3.5</td><td>4.3</td></tr><tr><td>Pet N<sup>a</sup></td><td>3.3</td><td>9.5</td></tr><tr><td>FNR<sup>b</sup></td><td>35.4</td><td>6.3</td></tr><tr><td>PetO<sup>c</sup></td><td>15.1</td><td>9.6</td></tr><tr><td>PetP<sup>d</sup></td><td>7.4</td><td>8.2</td></tr></tbody></table>

<caption>Table 3: Parameters of Subunits of *Cytb*<sub>6</sub>*f*</caption>

Subunits containing redox prosthetic groups with a high degree of conservation between *Cytb*<sub>6</sub>*f* and *Cytbc*₁ are: (i) *Cytb* in the *Cytbc*₁, which is equivalent in many structure-function aspects to *Cytb*<sub>6</sub> and subunit IV in the *Cytb*<sub>6</sub>*f*<sup>175</sup>; and (ii) the high potential (*E*<sub>m,7</sub> = + 300 mV) 2Fe2S, often

<footnote><sup>a</sup>From cyanobacterium *Nostoc* sp. PCC 7120; <sup>b</sup> PetH; resolved on SDS-PAGE, spinach prep; MW, *A. Thaliana*; <sup>c</sup> PetO, *C. reinhardtii*; <sup>d</sup> PetP, *A. platens*</footnote>
called the “Rieske” protein because of the origin of its discovery ¹⁷⁶. The protein subunit containing the covalently bound c-type cytochrome, Cytf and Cytc₁, respectively, is not conserved between Cytb₆f and Cytbc₁ complexes.

*The Four Small Peripheral Subunits, Pet G, L, M, and N.* A unique lattice of four short (3.3 – 4.1 kDa), single TMH “hydrophobic sticks,” denoted as PetG, PetL, PetM, and PetN (Table 3), provides a hydrophobic lattice, or “picket fence,” around each monomer, a structure which is unique among all known integral membrane protein structures. Of these four subunits, PetM ¹⁷⁷, along with the ISP, are the only subunits among the thirteen in the complex that are nuclear-encoded. Specific functions of these four subunits are not defined, although PetL interacts sterically with the TMH of the ISP and may provide a constraint on the membrane-spanning orientation of the latter. The position of much of the lipid in the complex (Fig. 12) between the Pet picket fence and the conserved Cytb₆-subIV core of the complex suggests a “boundary lipid” function and that the “picket fence” may have been added to the core of the complex relatively late in the evolution of the complex.

**3.6.2 Structure and spectral properties of hemes b, f, cₙ and 2Fe2S cluster.**

High resolution structures of a C-terminal soluble domain of the 250 residue extrinsic C-terminal domain of Cytf ¹⁷¹,¹⁷², and of the N-terminal soluble domain of the ISP¹⁷⁸, were obtained prior to their determination in the intact multi-subunit complex. The crystallized complex is a symmetric (C2 symmetry) dimer that consists of eight transmembrane polypeptide subunits in plants, green algae, and cyanobacteria ⁴². The analogous Cytbc₁ consists of eleven subunits in the yeast, PDB 1KYO ⁶⁸ and bovine, PDB 1QCK, 1BE3 ⁶⁰ complex, and three in the complex from the photosynthetic bacterium, *Rhodobacter sphaeroides* ⁶³. In all cases, the dimeric structure of the complex is required for a physiologically significant electron transfer activity, 250 – 400 electrons transferred (Cytf/sec)⁻¹ ¹⁷⁹–¹⁸¹. This is possibly because of a defined inter-monomer quinol entry pathway.

A ribbon diagram of Cytb₆f (PDB 2ZT9, 4H44, 4OGQ) derived from crystal structure analysis of the cyanobacterium *Nostoc* sp. PCC 7120 ⁶⁵,⁶⁶,¹⁷⁴, is shown in 2 views differing by a 180° rotation in order to display more clearly the arrangement of the 8 subunits that span the membrane with 13 TMH per monomer containing 7 prosthetic groups, including 5 with redox function consisting of 4 hemes, one 2Fe2S, one chlorophyll *a* ¹⁸² and one β-carotene ¹⁸³ (fig 10 A, B and fig. 11). These features were confirmed in a lower resolution (3.6 Å) cryo-EM structure of the spinach Cytb₆f.⁷³
<figure><img src="image_14.png" /></figure>

Figure 10. The Dimeric cytochrome *b*<sub>6f</sub> complex. (A) Polypeptide subunits from the cyanobacterium *Nostoc* sp PCC 7120 (PDB ID 4OGQ) are shown in ribbon format- *Cytb*<sub>6</sub> (yellow), subIV (light brown), *Cytf* (orange), ISP (dark green), PetL (red), PetM (light green), PetG (blue), PetN (black). (B) Prosthetic groups of *Cytb*<sub>6f</sub>. Transmembrane hemes *b*<sub>p</sub>, *b*<sub>n</sub> (red/blue), and *c*<sub>n</sub> (black/blue) are shown as sticks. On the *p* side, heme *f* (green/blue) is shown as sticks, the [2Fe-2S] cluster as spheres (brown/yellow), and the *Chl-a* (green/blue) and *β*-car (yellow) as sticks. (C) Heme-heme distances within the transmembrane domain of the complex (PDB ID 4OGQ). Distances shown as black dashes. Heme edge-edge, and center-center (Fe-Fe) distances are shown, respectively, outside and inside parentheses (selected atoms shown as spheres). The dielectric constant, ε<sub>ij</sub>, between each pair of hemes, ε<sub>n1,p1</sub>, ε<sub>n1,p2</sub>, ε<sub>n2,p1</sub>, etc.<sup>184</sup> is shown. (D) Transmembrane heme ligation in *Cytb*<sub>6f</sub>. The central Fe atom of heme *b*<sub>p</sub> and *b*<sub>n</sub> is axially ligated, respectively, by residues His86/His187 and His100/His202 which bridge the B and D TMH of *Cytb*, Heme *c*<sub>n</sub> is covalently attached to the protein via Cys35. Heme *c*<sub>n</sub> is unique as it lacks an amino acid axial ligand, and the central Fe-atom is penta-coordinated. The sole ligand of
heme $c_n$, is provided by H₂O or OH⁻ on the heme $b_n$ side. Fe of heme $c_n$ separated by 4.0 Å from a propionate oxygen of heme $b_n$, which results in electronic coupling, a high spin g =12 EPR signal ⁴³,¹⁸⁵, and an oxidase-like reaction with nitric oxide ¹⁸⁶. Electrochemically positive, negative sides of membrane labeled p, n. (Reprinted/adapted by permission from Springer Nature, Cytochrome Complexes: Evolution, Structures, Energy Transduction, and Signaling; Structure-Function of the Cytochrome b6f Lipoprotein Complex by William A. Cramer, S. Saif Hasan, 2016).

<figure><img src="image_15.png" /><figcaption>Figure 11. High potential electron acceptor of Cyt-bc: Cytf of Cytb6f (A, B) and Cytc1 of Cytbc1 complex (C, D). Cytf</figcaption></figure>

and Cytc₁ are attached to the respective complexes and membrane through one TMH. Cytf binds c-type heme (heme *f*, red/blue sticks) in the β-sheet extrinsic domain. Cytf extrinsic domain of Cytc₁, with c-type heme (heme *c*₁, red/blue sticks) is mostly α-helical. **B** and **D** rotated 90° about horizontal axis relative to **A**, **C**, **E**, tridecyl-stigmatellin (TDS, blue sticks), bound in an 11 Å long channel of Cytb₆ (purple) terminating proximal to the 2Fe2S cluster. The Qp-portal in Cytb6f is marked as the black rectangle. ISP-HD is shown as orange ribbons. A chlorophyll molecule (Chl *a*), inserted between F and G helices of subunit IV (cyan), using its phytyl tail, functions as a gate for quinol/quinone traffic in the Qp-portal. Drawing by S. Saif Hasan

*Unique Chlorophyll a and β-Carotene.* The presence of a single chlorophyll *a*¹⁸²,¹⁸⁷,¹⁸⁸ and β-carotene ¹⁸³ in each monomer of *Cytb*₆f is enigmatic. The β-carotene is separated by 14 Å from the chlorophyll ⁴¹,⁴², too large to allow β-carotene to quench a chlorophyll excited triplet state ¹⁸⁹. (ii) The β-carotene protrudes by approximately 11 Å from the complex between the TMH of Pet G and Pet M.

*Transmembrane Hemes, b*n and $b_p$. The four TMH of the *Cytb*₆ are connected by loops at the *p* and *n* side membrane interfaces. SubIV (~17 kDa) has three TMH (E–G) that form a *p* side saddle around the four helix bundle of *Cytb*₆. Helix E of SubIV is located in proximity to the A- and B-helices of *Cytb*₆ while the F and G-TMH span the four helix bundle, close, respectively, to the C, D and B-TMH (Fig. 12). The C-terminus of the E-TMH is separated from the N-terminus of the F-TMH by a distance of ~40 Å, which is bridged by the *p* side *ef*-loop. This seven TMH assembly forms the
conserved core of *Cytb*<sub>6f</sub>, which is bounded in each monomer by the four subunit hydrophobic “picket fence”.

The arrangement of the two pairs of transmembrane *b*-hemes, *b*<sub>p</sub> and *b*<sub>n</sub>, with inter-heme edge-
edge and center-center (Fe-Fe) distances (in parentheses) is shown (Fig. 10), along with the
heterogeneity of the inter-heme dielectric constants <sup>184</sup>, and the complex of heme *b*<sub>n</sub> with a skeleton
of the covalently bound *c*-type heme *c*<sub>n</sub> <sup>43,185,186,190,191</sup>. The heme *b*<sub>n</sub>-*c*<sub>n</sub> complex, a unique feature of
the *Cytb*<sub>6f</sub> compared to the *Cytbc*<sub>1</sub>, in which the close proximity of the two hemes results in sharing
of the 3d electronic shells is displayed in a *g* = 12 EPR signal<sup>43,185</sup>.

*Hemes, b*<sub>*n*</sub> *and b*<sub>*p*</sub>, *Oxidation-Reduction Potentials*. Determination of the oxidation-reduction potentials of these hemes in *Cytbc*<sub>1</sub> are consistent in showing a separation of 100-150 mV between the *E*<sub>m</sub>'s of the two hemes, which are thus often labeled *b*<sub>H</sub> and *b*<sub>L</sub> (high and low potential hemes). For the two hemes in the *Cytb*<sub>6f</sub>, there is disagreement as to whether the *E*<sub>m</sub>'s of the two hemes are <sup>191</sup> or are not <sup>40,192</sup> separable in a redox titration of thylakoid membranes. A simulation shows that a midpoint potential difference of 50 mV or less of a one electron titration cannot be resolved. For the isolated complex, the midpoint potentials of the two hemes in the *Cytb*<sub>6f</sub> are separated by approximately 90 mV.

*Reducibility of b Hemes; Heterogeneity of Internal Dielectric Constants.* Excitonically split circular dichroism (CD) spectra of the *Cytbc*<sub>1</sub><sup>193</sup> and the *Cytb*<sub>6f</sub><sup>184,194</sup> arise from interactions of the reduced hemes. Determination of the time course of heme reduction and the onset of the split CD spectra for the isolated complex shows that the heme pair that is preferentially reduced is the intra-monomer pair, *b*<sub>p</sub> and *b*<sub>n</sub>. This result contrasts with that which is expected. If heme *b*<sub>n</sub>, whatever its exact redox potential, has a more positive potential in isolated *Cytb*<sub>6f</sub> than heme *b*<sub>p</sub>, the two hemes *b*<sub>n</sub> should be preferentially reduced under equilibrium conditions. The straight-forward explanation is that the protein medium between hemes *b*<sub>p</sub> and *b*<sub>n</sub> is more polarizable, i. e., has a higher dielectric constant, than the medium between the two hemes *b*<sub>n</sub><sup>184</sup> implying that the dielectric constant in the complex is heterogeneous and anisotropic.
<figure><img src="image_16.png" /><figcaption>Figure 12. Transmembrane helices and distribution of lipid in Cytb<sub>6f</sub> complex (Figure drawn by S. Saif Hasan). Reprinted/adapted by permission from Springer Nature, Cytochrome Complexes: Evolution, Structures, Energy Transduction, and Signaling; Structure-Function of the Cytochrome b<sub>6f</sub> Lipoprotein Complex by William A. Cramer, S. Saif Hasan, 2016.</figcaption></figure>

The unique heme c<sub>n</sub> at the n side quinone (Qn) binding site of Cytb<sub>6f</sub><sup>173,191</sup>, located in close (4 Å) proximity to heme b<sub>n</sub>, constitutes a major structure difference between Cytbc<sub>1</sub> and Cytb<sub>6f</sub> complexes. Consequently, it would also imply a difference, compared to the mitochondrial and bacterial Cytbc<sub>1</sub>, in the details of the involvement of heme b<sub>n</sub> in the transmembrane electron transport pathway [*n.b.*], heme c<sub>n</sub> is designated heme c<sub>i</sub> in the *C. reinhardtii* crystal structure, which first emphasized its unique existence in the structure of Cytb<sub>6f</sub><sup>41</sup>.

Heme c<sub>n</sub> has no axial ligand on the heme surface facing the intermonomer cavity. As shown by EPR studies which document its open axial ligand position <sup>186</sup>, and co-crystallization of the Cytb<sub>6f</sub> complex with quinone analog inhibitors <sup>173</sup>, it has been inferred that heme c<sub>n</sub> serves as the n side quinone binding site (Qn site). The Qn site is located closer to the membrane-water interface in Cytb<sub>6f</sub> complex than in the respiratory Cytbc<sub>1</sub>, as noted in the original crystal structure <sup>41</sup>. This surface-proximal location of the Qn site implies that a short pathway may suffice for proton conduction to the n side bound quinone from the n side aqueous phase. The crystal structure of Cytb<sub>6f</sub> from the filamentous cyanobacterium *Nostoc* PCC 7120, revealed a unique anhydrous Asp20→Arg207
(Cytb₆) pathway for proton conduction from the *n* side aqueous phase to the Qn site for reduction and quinone protonation ⁶⁶.

*ISP Electron Transfer; a p Side Conformation/Mobility Problem.* One remaining structure-function problem is that the closest approach distance of the Cytf heme to 2Fe2S in Cytb₆f is 28 Å, which is much too large to support physiologically meaningful millisecond electron transfer rates ¹⁹⁵,¹⁹⁶. How, then does 2Fe2S of ISP transfer electrons to Cytf? A major gap in understanding of the *p* side electron transfer reactions for the Cytb₆f results from the absence of the structure data that would allow kinetically competent electron transfer from 2Fe-2S to the heme of Cytf. Given the documented conformation change determined from crystal structure data for the avian mitochondrial Cytbc₁ (see section 3.4 for details), electron transfer from 2Fe2S to the Cytf heme must involve a rotational-translational conformational change in the ISP ⁶⁶.

### 3.6.3 Lipid content and other subunits

*Lipids in the Cytb*<sub>*f*</sub>* Lipoprotein Complex.* (i)'Boundary lipids' define a domain structure. Because the conserved Cytb<sub>6</sub>-SubIV core of the complex is separated by a lipid layer from the "picket fence", it is implied, as noted above, that the "picket fence," was added to the core structure at a later stage in the evolution of the Cytb<sub>6f</sub>, perhaps to facilitate interaction with the reaction center complexes. As noted above, the β-carotene, which protrudes from the surface of the picket fence, may act as a "latch" to facilitate super-complex formation with the PSI reaction center complex <sup>197</sup>.

*Lipid Content* The dominant lipid species in plant Cytb*<sub>f</sub>* are monogalactosyl-diacylglycerol (MGDG), digalactosyl-diacylglycerol (DGDG), phosphatidyl-glycerol (PG) and sulfoquinovosyl diacylglycerol (SQDG). The galactolipids (MGDG, DGDG and SQDG) and phospholipids dilinolenoyl-, phosphatidyl glycerol (DLPG), 1,2-dioleoylphosphatidylglycerol (DOPG) and 1, 2 dioleoyl-sn-glycerol-3-phosphatidylcholine (DOPC) stabilize the Cytb*<sub>f</sub>* to a varying extent <sup>198</sup>.

*PGRL Components.* A 35.7 kDa (pK = 5.2) protein, "PGRL1," is present at unknown stoichiometry in the PSI super-complex *C. reinhardtii* <sup>197</sup>, and at a stoichiometry comparable to, although smaller (0.5:1), that of ISP in *Arabidopsis thaliana*, and has been inferred to be the ferredoxin-quinone reductase <sup>199</sup>. The absence of the protein results in a partial decrease of the chlorophyll fluorescence yield that results from reduction of the quinone pool. The redox function is believed to be derived from the six cysteine residues in the protein, perhaps in conjunction with iron that is present in the preparation. PGRL1 is also inferred to be able to bind to the Cytb<sub>6f</sub> <sup>199</sup>, although it has not been detected in mass spectroscopic analysis of isolated Cytb<sub>6f</sub> <sup>200</sup>. In addition, it is noted: (a) the fluorescence yield effects ascribed to PGRL1 are perhaps not quantitative indicators of an
obligatory function in the ETC, as they involve changes of a factor of 2-3 on a time scale of 10 seconds, approximately a factor of 1000 times slower than the rate-limiting step of electron transport in this region of the ETC; (b) the rate of P700 reduction, a standard kinetically competent assay for cyclic electron transport, in the presence and absence of the peptide has not been reported. (c) Details of the chemical nature and quantitative properties of the hexaCys-Fe redox moiety in PGRL1, proposed to serve as the redox group responsible for FQR activity, are not presently available. The significance of the PGRL1 components may need to be re-evaluated in light of the recent information on their very diminished stoichiometry relative to the known electron transport components<sup>201</sup>

**3.6.4 Q cycle in cytochrome $b_6f$: mechanism and controversies**

The observation of oxidant-induced reduction<sup>202</sup> in *Cytbc*₁ (see details in section 7), and measurement of a greater than unity ratio of coupled protons translocated to electrons transferred, $H^+/e > 1$, was important in the formulation of the ‘modified Q cycle’ mechanism<sup>39</sup>, whose general formulation and application to photosynthetic membranes<sup>27</sup> followed 15 years after the original inference of the chemiosmotic concept of membrane energization<sup>28</sup>. The demonstration of oxidant-induced reduction of *Cytb* heme in the mitochondrial *Cytbc*₁ and the requirement in this electron transfer reaction for the high potential (ca. +0.25 V) 2Fe2S was elucidated in studies by B. Trumpower<sup>203</sup>. An apparently anomalous reduction of the *Cytb* component was observed in a preparation of mitochondrial membrane protein ‘complex III’. Support for the Q cycle model of proton translocation in the mitochondrial *Cytbc*₁ was provided in studies by Wikstrom and Saraste<sup>204</sup>, and Rich<sup>205</sup>.

Support for the application of Q cycle model to *Cytbc*₁ in the purple photosynthetic bacteria has been well documented and discussed extensively in reviews ¹,²,¹³,³⁹,¹¹⁷,¹¹⁸,²⁰⁶⁻²⁰⁸. Studies on function and, lately also the structure of the related *Cytb*<sub>6f</sub> functioning in chloroplasts, green algae, and cyanobacteria, have been reviewed in the last decade ¹⁷,¹⁹,²²,¹⁸⁴,²⁰⁹⁻²¹⁶.

While there is a general consensus on the occurrence of the “modified Q cycle” in the case of *Cytbc*₁, (see section 3.2 for details) experimental evidences exist that are not completely in agreement with this hypothesis in the case of *Cytb*<sub>6f</sub>. In the following, we summarize this evidence, compare contrasting results, and summarize different interpretations of the results.

Arguments which imply that the PQ/PQH₂ cycle in *Cytb*<sub>6f</sub> is different from the “modified Q cycle” proposed for regeneration of UQH₂ through function of *Cytbc*₁ in the respiratory chain and photosynthetic bacteria are:

i) the presence of an additional heme c<sub>n</sub>, which could act as a PQ reductase. Based on its redox
properties and location within the complex, this heme could catalyze the injection of an electron into the quinone binding side using stromal reductants as a donor (i.e. the presence of ferredoxin: NADP+ reductase (FNR) bound to purified plant (spinach) Cytb6f, see below). The electronic coupling between hemes b_n and c_n, would imply that PQ reduction need not proceed through a semiquinone intermediate but perhaps cooperatively through hemes b_n and c_n, providing protection against the formation of superoxide and other reactive oxygen species. This mechanism is not compatible with the modified Q cycle mechanism, but, as introduced in section 3.2, it is actually consistent with the original mechanism of the Q cycle as proposed by Mitchell (see Mulkidjanian ¹⁹ for a discussion) (see also Fig. 4).

The possibility of direct injection of stromal electron in the *b* hemes could also account for another observation that is not entirely compatible with the modified Q cycle in Cytb6f:

ii) the 'slow' electrochromic phase attributed to oxidation of the PSQ in the Qp site and transfer of its electron across part of the low dielectric membrane to reduce the intra-membrane *b* hemes is also observed when the hemes were chemically reduced and therefore not able to function as an electron acceptor <sup>192</sup>. In principle, injecting 1 electron into the stromal site via the heme c_n would allow oxidation of one *b* heme, generating one PQH<sub>2</sub> and regenerating the Q cycle (the so called 'activated' Q cycle mechanism, <sup>19</sup>). Note, however, that another explanation has been proposed by Joliot and Joliot, who suggest that the slow electrochromic phase observed in reducing condition could reflect the transmembrane movement of a charged SQ, (the SQ cycle hypothesis, <sup>217</sup>, see however <sup>218</sup> for a different view). This mechanism may represent an adaptation to reducing conditions when no Q is available at the Qn site. Note, however, that some inhibitors of Cytb6f (NQNO, stigmatellin and MOA-stilbene) act on the *n* side at the side of heme c_n that faces the quinone-exchange cavity, as shown in crystal structure<sup>173</sup> and prevent oxidant-induced reduction in Cytb6f, in agreement with the modified Q cycle hypothesis.

iii) the initial slope of the 'slow' electrochromic phase shows a pronounced isotopic effect, being slowed four-fold by a H<sub>2</sub>O/D<sub>2</sub>O substitution, in contrast to the redox-reactions of Cytf and Cytb<sub>6</sub> that were only slightly affected <sup>219</sup>. These findings have led to the hypothesis that proton pumping across Cytb6f could be triggered by the oxidation of PQH<sub>2</sub> <sup>220</sup>, an idea that is not conceived in the frame of the 'modified' Q cycle, but that is supported by the analysis of Cytb6f mutants of *Chlamydomonas* <sup>221</sup>.

iv) one component of the 'driving force' for the Q cycle in mitochondria and the purple photosynthetic bacteria is the pronounced difference (> 100 mV) in the redox potentials of the two b-type hemes, (i.
e., $b_L$ and $b_H$) which span the membrane. However, it has not been possible, in redox titrations done in *situ*, i. e., in thylakoid membranes to define, through anaerobic redox titrations a difference in midpoint redox potentials of the two *b* hemes, $b_p$ and $b_n$ ⁴⁰,¹⁹²,²²², More recently, however, Alric and colleagues used a different approach for redox titrations and found $E_m$ differences between the two *b* hemes of isolated Cytb₆f from *Chlamydomonas* (-130 mV and -35 mV)¹⁹¹, which are closer to those predicted by the ‘modified’ Q cycle. Thus, an $E_m$ difference is seen for the isolated Cytb₆f complex, but this difference is not clear in titrations done in *situ*, in membrane preparations.

v) studies on the kinetics of heme *b* reduction show that an intramonomer pair (hemes $b_n$ and $b_p$) is reduced first, rather than the two hemes $b_n$ being preferentially reduced as implied by the Q cycle ¹⁸⁴. Moreover, reduction of the two *b* hemes is not observed with repetitive laser flashes ⁴⁰.

vi) The existence of CET in oxygenic photosynthesis could help to maintain a high $H^+/e^-$ ratio, as required for proper CO₂ assimilation, thereby making the occurrence of a Q cycle less stringent in photosynthesis than in respiration. One of the main purposes of the Q cycle is to couple electron and proton transfer generating a $H^+/e^-$ ratio = 2 for proton translocation across the complex. Thus, the transfer through the high potential chain of the Cytb₆f of the 4 electrons arising from water splitting and oxygen evolution can result in the translocation of 8 protons into the chloroplast lumen which, together with the 4 translocated protons released into the lumen with each O₂ molecule, would provide 12 of the 14 $H^+$ needed to drive a full rotation of the rotatory c-ring of the chloroplast ATP synthase and therefore the synthesis of the three ATP molecules needed to fix a CO₂ molecule. In plants, the additional 2 $H^+$ could be generated by CET, assuming that it runs at a prescribed rate of 25 % of the linear non-cyclic pathway. Applying the same reasoning to the cyanobacterium, *Spirulina platensis*, where the ATP synthase c-ring contains 15 subunits, implies that CET would have to run at 3/8 or 37.5 % of the rate of the linear pathway to generate the additional three $H^+$ needed to drive a complete rotation. However, no such percentage of CET is observed in most photosynthetic organisms, at least under steady state conditions (see section 10). This suggests that CET alone, is not sufficient to provide the ‘extra’ $H^+$ for ATP synthesis. Consistent with this conclusion, Kramer and colleagues measured the $H^+/e^-$ ratio illumination in intact tobacco leaves and found that it was constant under low to saturating illumination. Therefore, they inferred that this ratio was maintained by a continuously engaged, proton-pumping Q cycle at Cytb₆f ²²³

Although, the Q cycle mechanism has a readily conceived role in contributing to the transmembrane proton flux required to provide ATP levels commensurate with the need
for CO₂ assimilation, it may be suggested that the Q cycle mechanism applied to Cyt b₆f is not obligatory (see however Cape et al, ²²⁴ for a different conclusion). An alternative mechanism is a membrane Bohr effect, a mechanism which is prominent in the description of the mechanism of proton pumping in the classical bacteriorhodopsin system ²²⁵, and which has been applied to the problem of proton translocation in the mitochondrial cytochrome c oxidase²²⁶.

**3.6.5 An additional electron path related with cyclic electron transfer in photosynthesis**

Electrons generated at the PSI reducing side can be re-injected into its donor side via the CET pathway. Discovered by Arnon in the 50's ²²⁷, this process is now considered as a relevant mechanism to counterbalance over-reduction of the PSI acceptor side ²²⁸ and to inject 'extra' protons into the thylakoid lumen, to adjust the ATP/NADPH ratio for CO₂ assimilation. Although the ratio of ATP/NADPH generated by LET is still uncertain ²²⁹, it could be insufficient to fuel CO₂ accumulation in chloroplasts and its assimilation via the Calvin, Benson and Bassham cycle (see Allen 2002 ²³⁰ for a discussion). In *viridiplantae* (including green algae and higher plants) CET could be the main route to optimise this process, by bypassing NADPH production while permitting the formation of the pmf, and therefore ATP synthesis.

Two main pathways have been proposed for CET (see section 10 for further details). The first one involves the activity of a chloroplast NAD(P)H dehydrogenase (NDH) complex (review in Peltier et al. ²³¹). In plants, this enzyme has similar characteristics to the mitochondrial complex I, while mainly sharing features with bacterial complex one in algae ²³². The plant complex would mainly use Fd as an electron donor, while the algal counterpart (Nda2 complex) uses NADPH as a source of electrons in the green alga *Chlamydomonas reinhardtii* ²³¹.

The second CET pathway would correspond to the so called Ferredoxin-Quinone Reductase complex (FQR), the existence of which was proposed by Bendall and coworkers in the 90's ²³³, based on the effect of antimycin – a putative inhibitor of this complex – on photosynthetic electron flow. An alternate target of antimycin in the chloroplast has been proposed by Sugimoto and colleagues as PGR5, a thylakoid protein ²³⁴ previously identified as a component of the CET pathway by a genetic screening or the *Arabidopsis thaliana* ²³⁵. The FQR complex itself has been putatively identified as a membrane complex ¹⁹⁹, containing PGR5 as well as PGRL1, another protein previously invoked as an essential component of the CET machinery in plants ²³⁶. However, see the critique of these studies in section 3.6.4.
Alternatively, the FQR may correspond to Cytb<sub>6f</sub> itself, mediating electron flow from reduced Fd to the PQ(H<sub>2</sub>) pool through the additional c heme in the stromal pocket<sup>41,42</sup>. This process could take place thanks to the interaction between the cytochrome complex and the Ferredoxin-NADP Reductase (FNR) enzyme (see section 4.3).

# 4 Characteristics of substrates for cytochrome bc<sub>1</sub> and b<sub>6f</sub>

As a main function of Cytbc is the generation of proton motive force utilizing energy associated with electron transfer from low-potential electron donors (membranous pool of quinone derivatives) to water soluble c-type cytochrome (c, c<sub>2</sub>, c<sub>6</sub>) or plastocyanin, it is worth introducing basic information on the substrates that are used by these enzymes. A brief summary on structure and redox properties of quinones is especially important as proton translocation and electronic bifurcation is largely associated with quinone chemistry.

## 4.1 Basic redox properties of Quinones

A role of quinones in electron transfer between membranous respiratory complexes was first proposed by Crane et al. in 1957<sup>237,238</sup>. It was shown that extraction of relatively low-weight molecules – ubiquinones from the membrane completely abolished the process of electron transfer within the mitochondrial respiratory chain<sup>239</sup>. In biology, the most widespread quinones belong to a family of 1,4-benzoquinone derivatives. While in eukaryotic cells quinones are generally restricted to ubiquinones (in mitochondria) and plastoquinones (in chloroplasts), prokaryotic organisms can use several other quinone derivatives as membranous electron carriers. Besides ubiquinones and plastoquinones that are commonly found in bacteria or cyanobacteria, respectively, there is a relatively large group encompassing naphthoquinone derivatives – menaquinones<sup>240</sup>. In some bacteria there are numerous, less common quinones that are based on other chemical structures of the redox-active rings. These are not discussed in this review. Interested readers are referred to a comprehensive review on biological quinones<sup>241</sup>.

Despite differences in chemical structure of the redox-active quinone rings, a common feature shared between mena-ubi- and plastoquinones is a relatively long hydrocarbon chain consisting of several isoprenoid fragments. This imposes a high level of hydrophobicity on natural quinones. The number of isoprenoid fragments in the chains is usually given by a number put after the name of the respective quinone<sup>241</sup>. For example, UQ-10 means that the ubiquinone posses the tail built of 10 isoprene molecules. The extent of hydrophobicity of quinones is expressed by a large partition
coefficient (log₁₀P) for organic/water mixtures such as octanol and water²⁴². Quinol forms are generally slightly more hydrophilic than the respective quinone forms. In case of ubiquinone-1 or plastoquinone-1, log₁₀P exceeds 3 and further increase in the length of isoprenoid tail increases the hydrophobicity, forcing quinone molecules to be constrained to a lipid environment²⁴². The natural quinones engaged in electron transfer usually have the hydrophobic tail built of 7 – 10 isoprenoid moieties, depending on the organism and very rarely exceeds 10. In plastoquinones the number of isoprenoids is usually 9, while in mammalian mitochondria ubiquinones usually have 9 or 10. For menaquinones, the length of the chain most often varies between 7 and 8²⁴¹. Comparison of the chemical structure of menaquinone-7, ubiquinone-10 and plastoquinone-9 is presented in figure 13.

<figure><img src="image_17.png" /><figcaption>**Figure 13.** Chemical structures of biologically important quinones with a different number of isoprenoid molecules comprising the hydrophobic side chains. From top to bottom there are: plastoquinone-9 (PQ-9), ubiquinone-10 (UQ-10) and menaquinone-7 (MK-7). The Eₘ,₇ values correspond to the average redox midpoint potentials at pH 7 for the respective Q/QH₂ couples.</figcaption></figure>

Another common feature that makes quinones important members of nearly all enzymatic processes of electron exchange between the membranous enzymes is their ability to undergo reversible two-electron, two-proton reactions of oxidation or reduction without damaging the chemical structure of the redox-active ring²⁴³. This reaction can be generally described as:

$$Q + 2H^+ + 2e^- = QH_2 \quad (\text{eq. 1})$$

where Q stands for quinone (fully oxidized form) and QH₂ – for quinol or hydroquinone (fully reduced form)
The complete oxidation/reduction reaction of eq. 1 can be divided into two, one-electron steps involving a semiquinone (SQ) intermediate.

$$Q + e^{-} = SQ^{-} \quad (\text{eq. 2})$$

$$SQ^{-} + e^{-} + 2H^{+} = QH_{2} \quad (\text{eq. 3})$$

Equations 2 and 3 assume that reduction of quinone leads to the semiquinone anion, and further reduction of SQ⁻ is coupled to the protonation of the reduced quinone. This particular proton/electron sequence is supported by the fact, that pK of neutral semiquinone (SQH) is significantly lower than the first pK (pK₁) of the first proton dissociation of QH₂²⁴⁴.

The average redox midpoint potential (Eₘ) of Q/QH₂ couple is equal to the arithmetic mean of redox midpoint potentials of the Q/SQ⁻ and SQ⁻/QH₂ couples²⁴⁵.

$$E_{m} (Q/QH_{2}) = 0.5(E_{1} + E_{2}) \quad (\text{eq. 4})$$

where: E₁ and E₂ represent Eₘ values of Q/SQ⁻ and SQ⁻/QH₂ couples, respectively.

The average redox potential of Q/QH₂ couple depends on the type of the redox-active ring of a particular quinone molecule and it generally decreases upon substituting the 2,3 and 5,6 positions of the ring with a methyl group²⁴⁶. Conversely, when the ring is substituted with a group that has high electronegativity such as chlorine or bromine, the Eₘ of Q/QH₂ rises considerably²⁴⁶⁻²⁴⁸. For example, when one considers a simple case of 1,4-benzoquinone, its average redox midpoint potential at pH 7 (Eₘ,₇) is approximately +300 mV and decreases to +230 and +176 mV upon attachement of one or two methyl groups to the ring at position 2 and 2,3, respectively²⁴⁶. On the other hand, substituting the ring with chlorine at position 2 and 5 increases the Eₘ to approximately +540 mV. Attachment of methoxy groups to the ring also increases the measured Eₘ in relation to 1,4-bezoquinone. For example, the Eₘ of 2,6-dimethoxy-1,4-benzoquinone is approx. +340 mV, which is higher by about 200 mV than the Eₘ of 2,6-dimethylo-1,4-bezoquinone. The redox midpoint potentials of quinones belonging to the family of menaquinones are significantly lower than those of benzoquinone derivatives, for example the Eₘ,₇ of 2-methy-1,4-naphtoquinone (menadione) is only -5 mV, which is much lower than +230 mV of methyl-1,4-benzoquinone. Due to such a low redox midpoint potentials of menaquinones, they are considered as “ancient devices” for electron transport in living organisms recruited at early stages of evolution by some eubacteria and/or archaebacteria, at a time, when the atmosphere on Earth was anaerobic¹⁶⁰,²⁴¹.

This general rule of changes in the Eₘ values of quinones containing different groups substituted to the ring holds true for natural, biologically active quinones that are used by living organisms as substrates for Cytbc. It means, that the Eₘ of Q/QH₂ couples increases in the order:
menaquinone/menaquinol ($E_{m,7}$[MK/MKH₂] = -60 mV), ubiquinone/ubiquinol ($E_{m,7}$[UQ/UQH₂] = 70 mV) and plastoquinone/plastoquinol ($E_{m,7}$[PQ/PQH₂] = 110 mV) (see fig. 13)

As the redox reactions of quinones involve 3 electron and 3 protonation states, the theoretical number of possible states is 9. However, from all these possible states (electrons and protons), QH⁺ (protonated quinone), QH₂²⁺ (doubly-protonated quinone) and SQH₂⁺ (doubly-protonated semiquinone) are not possible since their expected pK values are less than 0²⁴⁴,²⁴⁹. The typical pK₁ and pK₂ values for the first deprotonation of QH₂, leading to QH⁻, and the second deprotonation, leading to Q²⁻ are separated by 2, thus pK₂ = pK₁+2. Interestingly, pK₁ and pK₂ are inversely correlated with the redox midpoint potential of Q/SQ⁻ couples and for mena- ubi- and plastoquinone pK₁ is in the range 9-11²⁴⁶. For semiquinone, the pK value also depends on the $E_m$ of Q/SQ⁻ couple with typical values falling in the range 4 - 5²⁴⁴.

### 4.1.1 Stability and reactivity of semiquinone radical with molecular oxygen

A potential danger associated with quinone redox reactions, not only those catalyzed by enzymes, is the possibility of reaction of SQ with molecular oxygen²⁴⁸,²⁵⁰⁻²⁵⁴

$$ SQ^{-} + O_{2} = Q + O_{2}^{-} \quad (\text{eq. 5}) $$

Although this reaction is reversible, occurrence of superoxide dismutation or superoxide scavenging by, for example Cytc or PC, will shift the equilibrium to the right side²⁴⁶. A superoxide release during the enzymatic catalysis by Cytc₆ or Cytb₆f is considered deleterious and leads to a decrease in energetic efficiency of the enzyme, as generation of superoxide decouples the oxidation of QH₂ at the Q₀/Qₚ site with Q reduction at the Qᵢ/Qₙ site. Thus, this side reaction decreases the number of protons translocated across the membrane during the catalytic cycle. However, in living cells some portion of O₂⁻ can be scavenged by electron donation from the radical to the oxidized Cytc²⁵⁵ or PC²⁵⁶ with a relatively large second order rate constant. Such scavenging of superoxide can be considered as a protection and partial remedy to energy-wasting side reactions, as the electrons that leak to oxygen can be subsequently used to generate pmf at the level of cytochrome c oxidase (CcO) (complex IV)²⁵⁷.

The second-order reaction rate constant for the reaction of SQ⁻ with O₂⁻ depends on the $E_m$ value of the Q/SQ⁻ couple and generally increases when the potential of the $E_m$ of the Q/SQ⁻ couple decreases. When $E_m$($Q$/SQ⁻) is more negative than -200 mV, the reaction of SQ⁻ with dioxygen becomes diffusion-limited²⁴⁶.

The mechanism of reduction of O₂⁻ to O₂⁻ by the SQ⁻ radical is generally viewed as electron transfer process that follows the Marcus theory of electron transfer²⁵². An alternative mechanism
proposes a chemical reaction of addition of oxygen to the SQ⁻ ring followed by release of the superoxide anion, which after protonation changes to the neutral, peroxyl radical²⁵³.

$$SQ^{-} + O_{2} \rightarrow SQ-OO^{-} + H^{+} \rightarrow Q + HOO* \quad (\text{eq. 6})$$

Valgimigli et al. showed evidence that reaction 6 is significantly slower in solutions in which a semiquinone can form H bonds with molecules of the solvent²⁵³. Therefore, it can be supposed that formation of a hydrogen bond to the SQ⁻ may be protective against superoxide formation in catalytic sites of enzymes (see section 7.5).

### 4.1.2 Stability constant of SQ⁻.

Under equilibrium conditions Q, QH₂ and SQ⁻ are linked by a reaction of comproportionation (or in reverse direction by disproportionation)²⁴⁶:

$$Q + QH_{2} = 2 SQ^{-} + 2H^{+} \quad (\text{eq. 7})$$

The equilibrium constant of this reaction defines the stability constant (Kₛ) of SQ⁻ which depends on the difference in Eₘ values of Q/SQ⁻ (E₁) and SQ⁻/QH₂ (E₂) couples²⁵⁸⁻²⁶⁰:

$$K_{s} = \frac{[SQ^{-}]^{2}[H^{+}]^{2}}{[Q][QH_{2}]} = 10^{(E_{m}(Q/SQ) - E_{m}(SQ/QH_{2})) / 59.1} = 10^{(E_{1} - E_{2}) / 59.1} \quad (\text{eq. 8})$$

The stability constant for SQ⁻ defines the equilibrium between concentrations of Q/SQ⁻/QH₂ in this triad. It means that in equimolar mixtures of Q and QH₂ some SQ⁻ is also present but its relative amount depends on the equilibrium constant Kₛ. For some chlorine substituted quinones Kₛ can be higher than 1²⁴⁷. However, for biologically important quinone derivatives, the Eₘ of the Q/SQ⁻ couple is more negative than SQ⁻/QH₂, thus the exponent in the equation 8 is always less than zero (and therefore Kₛ < 1). Any further increase in the split between the Eₘ values of these two couples (E₁ - E₂) leads to a decrease in the equilibrium concentration of SQ⁻ in solutions containing a mixture of Q and QH₂ forms. This, on one hand, increases the rate of reaction of SQ⁻ with O₂ but on the other hand, decreases the probability of superoxide generation as the concentration of SQ⁻ decreases.

Despite the fact that Kₛ for SQ⁻ is used in chemistry of quinones under equilibrium conditions, it is also widely used to describe the properties of semiquinones that are generated at catalytic sites of enzymes such as the Qo/Qp and Qᵢ/Qn site of Cytbc during catalysis²⁶¹⁻²⁶⁴.
The concentration of quinones within the membrane is usually expressed as the number of molecules (Q and QH₂) vs the number of CcO molecules. In mammal and plant mitochondria the number of quinones per CcO is around 6-8, while it is much larger in yeast mitochondria (~ 38 per CcO). In purple bacteria the estimated number of quinone molecules per reaction center (RC) is also quite large - in the order of 15 – 30. The number of quinones in humans depends significantly on the tissue and the age. The content of Q in mitochondria increases during the first 20 years and after that time gradually decreases to the level even lower than after birth (see discussion in ref.¹³).

A significant content of the quinone molecules in the membrane may suggest that the comproportionation reactions should be very efficient leading to generation of a significant amount of SQ, which after dismutation would lead to the process of electron self exchange:

$$Q^{(A)} + Q^{(B)}H_2 = Q^{(A)}H_2 + Q^{(A)} \quad (\text{eq. 9})$$

However, such a reaction is very unlikely in membranes as it requires at least two steps that involve reactions of protonation and deprotonation²⁴³,²⁶⁵, which is difficult in the lipid environment.

### 4.1.3 Chloroplast photoactive vs non photoactive plastoquinone pools.

The concepts discussed above apply to quinol or quinone (Q(H₂)) in bioenergetic membranes in ideal conditions. However, due to crowding of both the photosynthetic and respiratory membranes, differences exist between Q(H₂) located in different lipid environments. In the case of photosynthetic membranes (the thylakoids) the proportion of Q(H₂) that participates in electron transport (i.e. that feeds the Cytb6f complex) does not correspond to the entire population of PQ(H₂). This fraction (the photoactive PQ(H₂) pool) makes up a relatively small percentage of the entire PQ(H₂) pool, because 60-70% of the total PQ(H₂) is non-photoactive, being stored inside thylakoid-associated lipid droplets known as plastoglobules²⁶⁶⁻²⁶⁸. There is increasing evidence that the size of this photoactive PQ(H₂) pool is tightly regulated in response to environmental cues, for example changes in light intensity²⁶⁹,²⁷⁰. This homeostasis is necessary because the PQ(H₂) pool fulfills multiple functions *in vivo*. Firstly, it shuttles electrons within the crowded thylakoid membranes,²⁷¹ probably via percolation through the lipid environment between other membrane complexes²⁷²,²⁷³. Moreover, its redox state controls physiological processes in response to environmental changes, such as changes in the PSII and PSI antenna size via state transitions²⁷⁴), as well as gene expression and pigment biosynthesis²⁷⁵.
Recent results suggest that the amount of PQ(H₂) active in electron transport is mainly regulated through exchanges between the photoactive pool (in the thylakoids) and the non-photoactive pool (in the plastoglobules). Homeostasis of photoactive PQ(H₂) abundance seems to be under the control of ABC1 (Activity of Cytbc₁) atypical kinases <sup>276</sup>. Knocking out a member of this family (*pgr6*, <sup>277</sup>, where the ABC1K1 kinase is inactivated (*abc1k1*) prevents changes in the size of the photoactive PQ(H₂) pool, that would otherwise occur in response to high light, facilitating increased electron flow and replacing photo oxidized plastoquinones. Thereby, *abck1* mutants display decreased electron flow and photoacclimation responses in high light <sup>278</sup>. ABC1K1 acts in tandem with ABC1K3, another member of the ABC1 family, and the push pull relationship between these two kinases provides a fine tuning mechanism to control the size of the PQ(H₂) pool in response to high light <sup>279</sup>.

This tight control of the size of the photoactive PQ(H₂) pool *in vivo* likely explains the phenotype of the *menD1* mutant of *Chlamydomoans reinhardtii*. This mutant is deficient in MenD, the enzyme catalyzing the first step of phylloquinone biosynthesis <sup>280</sup>. Therefore, the phylloquinones playing the role of electron acceptors within PSI are replaced by PQ in this mutant, making PSI less active. However, the *menD1* mutant has another phenotype - its photoactive PQ(H₂) pool is decreased by 20-30%. This can be explained by assuming that the size of the photoactive PQ(H₂) pool is constant between the WT and the mutant. Therefore, the overall number of diffusing PQ(H₂) molecules would decrease from around 6 per *Cytb6f* in the WT to around 4 per *Cytb6f* complex in *menD1*<sup>280</sup>. Based on this result, it is tempting to propose that the mechanism by which ABC1K1/ABC1K3 maintains a constant photoactive PQ(H₂) pool size depends on "quantification" of the total number of PQ(H₂) molecules available per electron flow chain. If this were the case, the size of the pool would be unresponsive in *menD1*, as the ABC1K1/ABC1K3 machinery will not deliver two additional PQ(H₂) per chain to the thylakoid membrane from the plastoglobuli to replace the PSI bound PQ.

## 4.2 Cytochrome *c* and plastocyanin as electron acceptors

Electron donation from both *Cytbc*₁ and *Cytb6f* to soluble proteins at the *p* side has been extensively studied. These molecular events have often served as pioneering models for electron transfer between different proteins, and reflect the development of techniques and theories of how such processes occur. In both cases, a single electron passes from a cytochrome in the complex (Cytc₁ in *Cytbc*₁ or Cytf in *Cytb6f*) to a small, soluble redox protein that is specifically, but transiently bound.
Once reduced, the electron carrier protein must rapidly dissociate (so as not to block the catalytic cycle), and diffuse through the *p* side space to another large membrane bound protein complex. In mitochondrial respiration this is cytochrome *c* oxidase (CcO, complex IV) <sup>281</sup>, while in plant photosynthesis this is photosystem I <sup>282</sup>. Protein acceptors vary between Cytc<sub>1</sub> and Cytf. In the case of the respiratory complex the acceptor is a c-type heme containing protein, cytochrome *c* (Cytc) <sup>283</sup>, perhaps one of the most studied of all redox proteins. Cytbc<sub>1</sub> in purple bacteria uses a homologue of Cytc, soluble Cytc<sub>2</sub>, and/or membrane-anchored Cytc<sub>y</sub> <sup>284,285</sup> as an electron acceptor, although this organism is photosynthetic and the final electron acceptor for the soluble electron carrier is a photosynthetic reaction center <sup>286</sup>. In the case of Cytb<sub>6f</sub>, the dominant electron acceptor is a copper-containing protein called plastocyanin (PC) <sup>287,288</sup>. It is generally accepted that PC has evolved to replace a Cytc homologue - Cytc<sub>6</sub>, the primordial acceptor for electrons coming from the Cytf subunit of Cytb<sub>6f</sub> (see De la Rosa et al., and references therein <sup>289</sup>). Following oxygenation of the atmosphere, oxidation of Fe<sup>2+</sup> to Fe<sup>3+</sup> in the ocean would have made it considerably less bioavailable, whilst oxidation of Cu to Cu<sup>1+</sup> would correspondingly have made it more soluble and therefore bioavailable. Interestingly, some prokaryotic photosynthetic organisms retain a cytochrome, Cytc<sub>6</sub> as their Cytf electron acceptor, and in primitive oxygen evolving cyanobacteria this can be the only electron acceptor from Cytf. Where both genes are present, Cytc<sub>6</sub> replaces PC under conditions of limiting copper <sup>290</sup>. Cytc<sub>6</sub> also occurs in some algae where it is also upregulated to act as an electron acceptor of the Cytb<sub>6f</sub> under copper limiting growth conditions <sup>291</sup>. A Cytc<sub>6</sub> homologue gene is present in plant genomes <sup>292</sup>. However, despite initial reports, the surface charge distribution on higher plant Cytc precludes its function as an effective *in vivo* electron acceptor from Cytf<sup>293</sup>.

The basic pattern of interaction is thought to be the same for electron transfer processes between Cytbc₁: Cytc and Cytb₆f:PC. The redox centers are oriented for optimal electron transfer by long distance charge effects in an encounter complex, before formation of a transiently stabilized productive complex, dominated by hydrophobic interactions. Mismatched surfaces between the partners, along with the retention of some water molecules at the interface, prevents interaction being so strong that it inhibits dissociation following the electron transfer event. The following sub-sections will describe the soluble electron carrier proteins in more detail, and outline the mechanisms by which soluble electron carriers interact with, and oxidize the Cytbc₁ and Cytb₆f complexes.
### 4.2.1 Redox properties and Interaction of Cytochromes c with Cytochrome $bc_1$

Cytochrome c is a small globular heme protein (90-120 amino acids, 10-13 kDa) which in bioenergetic systems serves as a water-soluble electron carrier. This definition encompasses the representatives found in eukaryotic (mitochondrial Cytc, yeast iso-Cytc) and prokaryotic (Cytc₂ present in several bacteria species) respiratory chains. A more distinct relative — Cytc₆ — is also a member of this category. However, due to its occurrence in plant-related systems, Cytc₆ is discussed in the chapter devoted to plastocyanin. For the scope of this review we will briefly go through the most important properties of Cytc with respect to its function and research in bioenergetic systems. More detailed information about the c-type family of cytochromes can be found in excellent reviews on cytochromes ²⁹⁴,²⁹⁵. In the Cytc molecule the heme is bound via two thioether linkages to the peptide. The covalent attachment of heme is a distinctive property of all c-type family cytochromes ²⁹⁶. The bond is formed between a cysteine residue (C) and the penultimate carbon of a vinyl side-group of the heme moiety. Comparison analyses revealed a specific heme binding motif – CXXCH (present also in Cytc₁ and Cytf, see sections 3.5.2 and 3.6.2, respectively). The cysteine residues (C) involved in thioether bonds are separated by any two residues (XX). This four amino acid subsequence is immediately followed by a histidine residue, which is an axial ligand to the heme iron. The second axial ligand is the sulphur atom of a methionine residue which together with four pyrrole nitrogen atoms of porphyrin completes the octahedral ligand geometry of the heme iron.

Redox properties of Cytc are related to the ability of heme iron to undergo reversible oxidation and reduction of ferrous ($Fe^{2+}$) and ferric ($Fe^{3+}$) states, respectively. The presence of a methionine ligand significantly elevates the redox potential of Cytc compared to bis-His coordination ²⁹⁷. Thus among biological compounds the redox midpoint potential of Cytc, and its counterparts, is considered high — in the range of +260 mV for horse heart Cytc ²⁹⁸ and bacterial Cytc₂ +360 mV ²⁹⁹. This is consistent with a location of Cytc in electron transport chains as an acceptor of electrons from quinol oxidising membrane complexes (i.e. Cytc – $Cytbc_1$; Cytc₂ – $Cytbc_1$; Cytc₆ – $Cytb_6f$) ²,¹³,²¹¹.

Upon reduction Cytc donates electrons to a terminal oxidase of the mitochondrial electron transport chain – CcO. In related systems cytochrome c-type electron carriers are oxidised by photosynthetic complexes of high oxidising power i.e. the bacterial photosynthetic reaction centre in case of Cytc₂ ³⁰⁰ or PSI for Cytc₆ in the case of cyanobacteria and some algae ³⁰¹. Beyond its primary role as a carrier shuttling electrons between membrane complexes, mitochondrial Cytc, is also involved in the process of cellular apoptosis in higher organisms ³⁰².

The amino acid sequence of Cytc varies between different organisms. The degree of identity spans quite high values (70%) among vertebrates ³⁰³ to low values (20%) when mitochondrial and
prokaryotic Cytc are compared <sup>304</sup>. Nevertheless general structural features are evolutionarily conserved – the number of helices (5) and their general spatial arrangement <sup>305</sup>. The heme is buried in a hydrophobic pocket within the apoprotein with only one edge being exposed to the surface of the protein. Such an asymmetric cover insulates most of the heme moiety from the water environment and only exposes the heme to the area where the binding occurs with physiological partners.

Cytc, as for other hemeproteins, absorbs strongly in the visible spectrum of electromagnetic radiation – in the range of 500-600 nm and the Soret band (around 400 nm). Reduced Cytc (Fe<sup>2+</sup>) shows a sharp peak at 550-556 nm (α band) and a slightly broader peak around 520 nm (β band). A unique feature of ferriheme c (Fe<sup>3+</sup>) is the presence of a weak absorbance band around 700 nm due to interaction with the sulfur of the methionine ligand. The peak in the Soret band (γ band) of Cytc shifts slightly when the redox state of heme c changes. The Soret peak is often used as a mark of the protein condition because when the protein denatures, significant changes in the γ band peak are observed. This was exploited in studies of Cytc folding stages and its stability <sup>306,307</sup>. The heme c also possesses peaks in the UV region around 320 nm and 280 nm (δ and ε band, respectively) <sup>308</sup>, which are, however, of little use due to overlapping signals from aromatic amino acids residues.

Octahedral heme iron in Cytc is in a low-spin configuration, hence, the reduced Cytc (Fe<sup>2+</sup>) is diamagnetic (S = 0) while oxidized (Fe<sup>3+</sup>) is paramagnetic (S = 1/2). Oxidized Cytc molecules from different organisms are the source of the EPR spectrum with significant g-factor anisotropy that usually exhibit rhombic symmetry with well resolved g-tensor principal values<sup>309</sup>. In some cases the EPR spectrum of Cytc is a HALS type<sup>310</sup>. These differences originate from structural changes in the mutual orientation of the His and Met ligands. However, no simple rules have been found so far which allow prediction of the type of EPR spectrum <sup>148</sup>. Such peculiarities are assigned to rather weak interaction with Met that under some circumstances can be disrupted. It was shown that pH changes, denaturation stress, or interaction with lipids (cardiolipin) can even lead to the breakage of the coordination bond changing heme c from hexa- to pentacoordinated high-spin geometry <sup>311-313</sup>.

Several studies involving measurement of the Cytc reduction rate by Cytc<sub>1</sub> and kinetics of electron transfer showed sensitivity to ionic strength <sup>314-319</sup>. This led to the notion that electrostatics is a key player in a "find-and-bind" game between the two protein contenders.

Chemical modification and site directed mutagenetic studies allowed identification of the crucial amino acid residues responsible for Cytc-Cytc<sub>1</sub> interaction. Indeed, it was established that the interaction is possible due to electrically charged surfaces. The surface of the Cytc molecule around the heme pocket has a net positive charge due to the presence of basic amino acid residues (mostly lysine residues), while the binding domain on the Cytc<sub>1</sub> surface includes complementary (negative) charges of acidic residues <sup>320-325</sup>. These long range interactions are thought to orient the
Cytc molecule in an encounter complex which enables maintenance of an electron-transfer-productive orientation at the early stages of interaction. Additionally the Cytc molecule itself possesses negatively charged residues on the side opposite to the interaction domain. This spatial separation of charges makes the Cytc molecule an electric dipole. It was shown that the presence of a dipole moment further facilitates the orientation of Cytc in the Cytc – Cytbc₁ complex that is proper for physiologically efficient electron transfer ³²⁶.

Following initial orientation, the proteins form a productive complex, knowledge of which is based on the structure of co-crystalized yeast proteins, either both in the oxidised, or both in the reduced state ⁷²,³²⁷. Only subtle differences in structure are observed between the different redox states. In these complexes, the oppositely charged side-chain pairs on Cytc₁ and Cytc do not directly interact, being 4 to 9.6 Å apart and therefore beyond the Debye length in physiological salt concentrations. Rather, the interface is dominated by non-polar interactions. The core surface area of interaction is 880 Å² around the heme clefts, and contains a core of 4 pairs of amino acids in non-polar interactions, including a cation-π interaction between a Cytc₁ Phe residue and a Cytc Arg residue. The interface contains 30 water molecules, only 2 of which coordinate H bonds between the proteins. This highly solvated surface aids rapid dissociation of the complex. The two heme cofactors have an inter-planar angle of 55° and are very close – only 4.1 Å between the closest thioester bonded carbons of the tetrapyrrole rings, and 17.4 Å between the Fe centers. This short distance means that, in principal, electron transfer could already occur when Cytc is approaching the Cytbc₁. It is possible that many other, less stable, productive complexes are also possible aside from those in the crystallized structure. A molecular dynamics approach has attempted to rationalize the apparent importance of charged side chains in mutational studies with the dominance of non-polar interactions in the crystal structures ³²⁸. This analysis indicates that many of the lysine side chains on Cytc, which surround the hydrophobic interface, are highly dynamic, and capable of forming H-bonds or even salt bridges with Cytc₁. Based on this, the authors suggest a more dynamic mixture of interaction modes between Cytc and Cytc₁.

No significant structural changes seem to occur between non-complexed and complexed proteins in the crystal structures, although one loop region that is poorly resolved in the crystal structure does change orientation in the molecular dynamics experiments ³²⁸. This in combination with the small surface area of interaction and short distance between the cofactors probably contributes to the rapid rate of electron transfer by the complex. Interestingly, only one Cytc₁ site in the dimer is occupied by Cytc in the crystal structures (Fig. 14 A), and this coincides with an increase in Qᵢ site occupancy in the same monomer, indicating binding of the substrates could be coordinated. Indeed, a change in the conformation of ISP-HD head group is observed on Cytc binding to the
Cytbc₁. The molecular dynamics studies on Cytc₁—Cytc interaction show that Cytc binding to one monomer induces structural changes at the Cytc binding site of Cytc₁ in the other monomer ³²⁸, perhaps explaining this. Co-crystalisation of yeast Cytc₁ with a second yeast isoform of Cytc (Cytc isoform 2), showed a nearly identical structural arrangement, although some of the charged pairs vary slightly ⁷².

Interestingly, an inspection of the electrostatic surface potential of crystal structures with different ISP-HD positions indicates that a negatively-charged area accessible to Cytc is significantly larger when ISP-HD is in the b-position compared to the case when it is in the c-position (Fig. 14 B). This could potentially influence the molecular organization of Cytc near the binding domain.

The idea of one Cytc per dimer stoichiometry attracted some attention and brought some productive scientific effort. On one hand this idea suggested that binding of Cytc to the one monomer disables the binding site in the other monomer as explored in context of *half-of-the-site* activity model of Cytbc₁ regulation. This proposed that during enzyme turnover only one monomer of Cytbc₁ is active at a time ³²⁹. On the other hand it provoked other authors to investigate the stoichiometry issue by other experimental approaches. Titration data obtained by plasmon resonance spectroscopy suggested that Cytc₂ interacts with the Cytbc₁ dimer in a biphasic manner which was interpreted as a modulating effect of ISP-HD on the binding ³³⁰. The data obtained by NMR and ITC for Cytc interaction with the soluble Cytc₁ head domain in low ionic strength (both in plant and bovine system) proteins revealed two binding sites of different affinity on Cytc₁ ³³¹. The authors proposed that Cytc₁ exposes a *proximal* site at which electron transfer takes place, and the *distal* site, which is too far for electron transfer, but maintains another molecule prior to electron transfer. In this way a subpool of Cytc is trapped close to the surface of the Cytbc₁–CcO supercomplex. This restricts diffusion of Cytc in the vicinity of the Cytbc₁–CcO complex leading to the enhancement of electron flow rate between the two complexes. The Cytc then does not need to return to the intermembrane bulk phase. A somewhat similar finding was revealed by experiments in which titration of Cytbc₁ binding sites was done by means of spin-labelling and pulse EPR ³³². Titration curves obtained at low ionic strength (no NaCl in buffer) were explained by a multiple-site-binding model. When electrostatic interactions are strong, Cytc₁ attracts many Cytc molecules which organize in a molecular cloud near the Cytc₁ binding surface (Fig. 14 C).

The putative presence and a role of such supramolecular structures in the Cytc pool are intriguing. However as the authors state in both cases (NMR, ITC) and (pulse EPR) the titration curves at higher ionic strength (25-50 mM NaCl) could be readily explained with one binding site per Cytc₁ ³³²,³³³. This means that binding of Cytc to the proposed additional sites under physiological conditions is much weaker than binding to the primary (electron transfer relevant) site on Cytc₁.
Hence in order for these additional collective interactions in the Cytc subpool to be physiologically relevant, other factors including macromolecular crowding, may be involved. This matter remains to be determined.

All available titration data report dissociation constants in the range of several to tens of µM, indicating that Cytc — Cytbc₁ affinity is not very strong <sup>330-332</sup>. What is even more striking is that EPR data indicates that when the ionic strength is increased to the physiological levels (100-150 mM NaCl) binding of Cytc to the primary interaction site is not even detectable <sup>334</sup>. Even though the stationary concentration of stable long-lived Cytc — Cytbc₁ complex is low, the proteins do interact because the electron transfer between them is not stopped <sup>314</sup>. Therefore this interaction is transient and the life-time of the complex is much shorter than the timescale of electron transfer. This would also explain the discrepancy between the measured rate of Cytc — Cytc₁ electron transfer that turned out to be several orders of magnitude lower<sup>314</sup> than the calculated electron transfer rate based on crystallographic data <sup>327</sup>. This supports a diffusion-coupled (in contrast to diffusion limited) mechanism of electron transfer between Cytc and Cytbc₁, in which the proteins constantly collide and several collisions are needed before one ET event occurs (Fig. 14 C) <sup>332,334</sup>. Therefore, the dynamic aspect of the Cytc — Cytbc₁ interaction has tremendous impact on the flow of electrons from Cytbc₁ to Cytc pool and it cannot be neglected.
<figure><img src="image_18.png" /></figure>

Figure 14. Binding of Cytc to Cytbc₁. A) Fragment of the structure of Cytbc₁ co-crystallized with Cytc (PDB: 1KYO). B) Comparison of electrostatic potential of the surface of Cytbc₁ accessible to Cytc in two different positions of ISP-HD. C) Schematic representation showing different molecular organization of Cytc near the binding domain of Cytbc₁ at low and high ionic strength.

### 4.2.2 Redox properties and Interaction of Plastocyanin with Cytochrome

#### b₆f

A structure of co-crystallized Cytf and PC is lacking, and so our understanding of their interaction is based on a combination of mutagenesis studies, nuclear magnetic resonance (NMR) and molecular dynamics. These have recently been augmented with atomic force microscopy (AFM) and 2D infra-red spectroscopy (2D-IR) experiments, meaning that although we lack a perfect snap-shot of any one interaction mode, we do have an excellent understanding of the dynamic events that occur during oxidation of Cytb₆f.

While the C-teminal domain of Cytf forms a membrane spanning domain, the soluble, N-terminal portion contains the heme group. The heme lies below the surface for interaction with PC or Cytc₆¹⁷¹. This is composed of a hydrophobic surface with a ridge of associated charged residues. The
hydrophobic surface is highly conserved between Cytf proteins from different species, while there is great variation in the ridge residues, and even their charge, between different groups of photosynthetic organism ³³⁵⁻³³⁷. In higher plants and algae, the ridge is functionalized by basic residues, and mutation of these disrupts interaction with PC ¹⁷¹,³³⁸⁻³⁴⁰. The charged residues on the Cytf ridge in cyanobacteria are by contrast usually acidic ³⁴¹,³⁴².

The secondary structure of PC is fundamentally different from Cytc. It is an 11 kDa protein composed of eight β strands and a small α-helix ³⁴³⁻³⁴⁶. It contains a single copper center, coordinated by one cysteine, one methionine and two histidine residues. There are two areas on the surface of PC involved in association with Cytf ¹⁷¹,³⁴²,³⁴⁷⁻³⁵⁰. The first (site 1) is a hydrophobic patch over the copper-His ligand, while the second (site 2) is functionalized by charged residues complementary to those on the Cytf. These therefore vary between photosynthetic organisms, being predominantly acidic in plants and algae, but basic in cyanobacteria. Long distance attraction between the charged residues on the surface of Cytf and PC are proposed to orient the molecules for optimal electron transfer, prior to formation of the catalytic complex. This is supported by the observation that low ionic strength results in a strongly bound complex incapable of turnover, while high ionic strength disrupts complex formation ³⁵¹. In addition, AFM experiments, with the Cytb6f complex tethered to the surface and PC attached to the tip, indicate that ionic strength strongly impacts both frequency of initial interactions and unbinding forces ³⁵². Despite this evidence supporting the importance of electrostatics for Cytf: PC interaction *in vitro*, the only *in vivo* analysis to date, on the alga *Chlamydomonas reinhardtii*, indicates that mutation of the basic ridge residues on Cytf has little impact on its oxidation rate ³³⁹,³⁴⁰.

While a recent molecular dynamics study supported a role for the charged amino acid side chains at site 2 in the initial complex formed between higher plant Cytf and PC ³⁵³, it also found no evidence for electrostatic pre-orientation in the cyanobacterium *Nostoc*. Moreover, there is little impact of electrostatics on formation of the Cytf:PC complex from the thermophilic cyanobacterium *Phormidium laminosum*, where non-polar interactions appear to both orient the complex and stabilise it ³⁴². Recent 2D-IR experiments on cyanobacterial (*Nostoc*) PC:Cytf interaction used site specific labelling of PC with cyanophenylalanine probes to measure very fast changes in conformation and orientation ³⁵⁴. This indicated that a wide variety of initial encounter complexes are formed, with rapid (1-2 ps) changes in orientation. It should be noted that this experiment was performed with the proteins in redox states opposite to that expected in electron flow (oxidized Cytf and reduced PC).

Following correct orientation of the electron transfer partners, interaction occurs mainly through non-polar interactions, with exclusion of some water from the interface. Chemical shift perturbation experiments indicate that in plants and cyanobacteria, the surface area of interaction is
around 600-850 Å², with the Cu-His ligand close to the Fe coordinating Tyr reside in Cytf³⁴⁹,³⁵⁰,³⁵⁵,³⁵⁶. Different plants appear to have the PC rotated slightly relative to Cytf, although the distance between the two redox centers remains around 11-14 Å. Although the surface charge distributions are reversed in cyanobacterial complexes, the orientation of PC relative to Cytf is conserved between higher plants and the cyanobacterium *Nostoc* ³⁴⁷. However, NMR studies ³⁴⁷ and modelling approaches ³⁵³,³⁵⁷ indicate that other cyanobacteria show different patterns of interaction: in *Phormidium laminosum*, where Cytf lacks the charged ridge, computer modeling indicates that PC is highly mobile, with only the site 1 hydrophobic patch interacting. This also seems to be the case for *Prochlorothrix hollandica*, where NMR studies show that although electrostatics make a modest contribution, site 1 dominates the interaction, although PC is oriented slightly differently ³⁵⁸. On binding to Cytf, X-ray absorption spectroscopy indicates that the geometry of the co-ordination sphere around the Cu ligand changes dramatically ³⁵⁹. This may be responsible for the -30 mV negative shift in PC redox potential of PC on Cytf binding ³⁶⁰, which counterintuitively makes electron transfer to PC less energetically favourable. Nevertheless, electron transfer is rapid, on the order of 104 kΩ s⁻¹ ³⁶¹,³⁶². It appears that the relative charge of the interaction partners also plays a large role in interaction, presumably in terms of the electrostatics of initial interaction. AFM experiments show a 5 fold increase in interaction frequency between the two proteins when redox states are opposite (irrespective of whether Cytf or PC is reduced or oxidized) ³⁵². This suggests that the rate of PC dissociation might be enhanced by rapid re-reduction of Cytf.

Cytb₆f also uses the Cytc homologue, Cytc₆ as an electron acceptor in cyanobacteria and algae grown under copper deficiency condition ²⁹⁰. It is reported to show a faster rate of electron transport between Cytf and terminal oxygenases than does PC ³⁶³. Intriguingly, Cytc₆ is also the principle electron acceptor in the heterocysts of the cyanobacterium *Anabaena*, even when Cu is replete ³⁶³. These cells are specialised for N-fixation, a process dependent on an O₂ sensitive enzyme (nitrogenase), and so PSII activity is very low, and the environment anaerobic ³⁶⁴. In heterocysts, the majority of electron transport is therefore thought to result in oxygen reduction or in cycling between PSI and Cytb₆f via PQH₂ and Cytc₆. *Anabaena* could no longer grow under diazotrophic conditions (obligatory N-fixing), when the gene for Cytc₆ was knocked out ³⁶³.

Cytc₆ is a 10 kDa single c-heme type cytochrome bound by a typical CXXCH motif, where the His and a Met act as axial coordinators ³⁶⁵. The protein has a midpoint potential of +335 mV ³⁶⁶, close to that of PC. In cyanobacteria, Cytc₆ has a hydrophobic surface area around the heme, and a positively charged surface patch of amino acid side chains analogous to sites 1 and 2 in PC ³⁶⁷. Although modelling of the *Chlamydomonas Cytf:Cytc₆* complex indicates that a charged patch on the Cytc₆ is important, many orientations are possible for the encounter complex ³⁶⁸. A combination of NMR and
molecular dynamics<sup>367,369</sup> has shown that the heme edge region (where the cofactor breaks the surface of the protein) is critical for interaction with Cytf, and long distance charge interactions approximately orient the two proteins in an encounter complex, as for PC. Many orientations of Cytc<sub>6</sub> result in cofactor-cofactor distances that are close enough for electron transfer, and thus no specific interaction complex is expected <sup>369</sup>.

Our knowledge of the interactions between Cytbc₁ and Cytb₆f carriers and their *p* side electron carrier is not equivalent – in the case of the respiratory complex, co-crystalization has given a wealth of molecular detail about one particular productive complex. The absence of such a structure for the photosynthetic complex has lead to the innovative use of NMR and molecular dynamics, along with other recent techniques to provide abundant information on dynamic events. It is to be hoped that we will reach an equivalence of knowledge between these two fascinating systems soon, so that more detailed comparisons can be made.

## 4.3 FNR/Fd interactions with cytochrome b₆f

### 4.3.1 Potential *n* side donors

In the initial model of the Q cycle proposed by Mitchell, one of the electrons reducing the heme *b*<sub>H</sub> site (heme *b*<sub>n</sub>) in the Cytbc<sub>1</sub> complex came from a soluble *n* side electron donor<sup>33</sup>. He revised this idea not long after proposing the classical model of the Q cycle<sup>32</sup>, and more recent studies have given us a much firmer grasp on the classical Q cycle mechanism (see <sup>216,259</sup> for detailed reviews). However, solution of the crystal structure of Cytb<sub>6</sub>f complexes<sup>41,42</sup> prompted reassessment of whether an *n* side soluble electron donor might contribute to a modified Q cycle. The discovery of an additional heme group, named heme *c*<sub>i</sub>, or heme *c*<sub>n</sub>, 6 Å closer than heme *b*<sub>n</sub> to the *n* side solute interface and in contact with the Q<sub>n</sub> site has re-ignited the debate as to whether the Cytb<sub>6</sub>f might indeed receive electrons from a soluble, *n* side source. By contrast, the Cytbc<sub>1</sub> lacks heme *c*<sub>n</sub>, and it is reasonable to assume that such a mechanism is precluded. This may be a reflection of the different bioenergetic environments of the two complexes, where the Cytbc<sub>1</sub> can rely on consistent availability of quinols, while the Cytb<sub>6</sub>f is maintaining a proton gradient in the face of variable quinone reduction by PSII, due to fluctuations in light intensity.

So, where could such *n* side soluble reductant come from? In photosynthetic systems, the final steps in electron transport involve the donation of electrons from photoexcited PSI to the soluble, single electron carrier protein ferredoxin (Fd), and subsequent oxidation of Fd by the Fd:NADP(H) reductase (FNR)<sup>370</sup>. In turn FNR transfers these electrons over a flavin moiety (flavin adenine dinucleotide, FAD) to the 2 electron carrier NADP<sup>+</sup>, making NADPH (see <sup>371,372</sup> for a detailed
description of the reaction mechanism). Both Fd and NADPH are used as electron donors by multiple
enzymes, supplying the cytosol (bacteria) or stroma (chloroplasts) with reducing equivalents.
Crucially, the FAD in FNR enables the relatively safe storage of a single electron from Fd, before a
second reduced Fd can donate the second electron necessary for NADP+ reduction. The injection of
these electrons back into the quinone pool from soluble, *n* side carriers in CET therefore depends
upon events involving Fd and FNR.

The two dominant mechanisms of CET have been historically separated on the basis of their sensitivity to the inhibitor antimycin A. Curiously, despite the sensitivity of the *Cytbc*₁ to this inhibitor, the *Cytb*₆f was until recently thought to be insensitive ³⁷³, and linear electron transport to NADPH was not found to be impeded by antimycin A. For many years it proved extremely difficult to identify the proteins involved in quinone reduction by the antimycin A sensitive CET pathway. Three independent mechanisms have been proposed: 1) direct electron donation from Fd to the *Cytb*₆f ¹⁹,³⁷⁴,³⁷⁵; 2) electron donation from FNR to the *Cytb*₆f ¹⁸⁰,³⁷⁶; 3) electron donation from Fd via *Cytb*₆f associated Pgr5 and PgrL1 pathways ¹⁹⁹. It remains a possibility that all three of these pathways occur, depending on the bioenergetic situation. Although robustly debated ²³⁴,³⁷⁷⁻³⁸⁰, it has been the consensus among many researchers that a pathway involving Pgr5/PgrL1 catalyses the bulk of CET in most algae and higher plants ¹⁹⁷,³⁸¹⁻³⁸⁵. However, results in *Arabidopsis* ³⁸⁶ and the green algae *Chlammydomonas reinhardtii* ³⁸⁷ lead the authors to propose that the role of these proteins is regulatory, rather than mechanistic ³⁷⁴,³⁷⁷,³⁸⁶,³⁸⁷. Moreover, the PgrL1 protein appears to be significantly sub-stoichiometric to the *Cytb*₆f, and would therefore only be able to drive electron flux into a small proportion of *Cytb*₆f complexes present in the membrane²⁰¹. Finally, it has recently been reported that Antimycin A can indeed act as an inhibitor of the Qi site on Cyt b6f in conditions of highly reduced PQ, dependent on the presence of the STT7 kinase and pgr5.³⁸⁸ Given the controversy, it is timely to discuss how the relationship between these different candidates and the *Cytb*₆f might help to resolve this question.

**4.3.2 Direct electron donation from Fd to cytochrome b<sub>6</sub>f**

As mentioned previously, the original Q-cycle proposed by Mitchell involved a single electron
from the soluble *n* side ³², and if this were to be the case for the *Cytb*<sub>6</sub>f, Fd would be an obvious
candidate for such a donor. The advantages of such a system in amplifying the proton motive force
(pmf) are evident, with additional electrons from the *n* side enabling immediate quinone reduction at
the Q<sub>n</sub> site on oxidation of a single quinone at the Q<sub>p</sub> site, and also providing the means to close
"incomplete" catalytic cycles in the *Cytb*<sub>6</sub>f that would otherwise leave *Cytb*<sub>6</sub> in a reduced state ¹⁹. It
is possible that this could be automatically poised by the redox state of the quinone and stromal
electron carriers, but seems more likely that some regulation would be required, particularly in the
dark, when reduced Fd is required for multiple other metabolic reactions ³⁸⁹ and continuous input of
*n* side electrons to the *Cytb*<sub>6f</sub> undesirable.

Nearly all photosynthetic organisms possess a suite of Fd proteins that are presumed, and in
some cases proven, to have variable affinity for different electron acceptors ³⁹⁰⁻³⁹⁵. Such a system
means that, depending on the relative abundance of these Fd iso-proteins, electron flux into different
pathways could either be increased or decreased. There is abundant evidence that Fd iso-proteins
specific for CET exist (see ³⁹⁶ for a full discussion). For example, maize plants perform an adapted
photosynthetic pathway known as C4, in which one cell type has a greatly increased ATP demand and
low PSII activity – a situation that demands high CET. A specific Fd iso-protein (FdII) is exclusively
present in this ATP demanding cell type ³⁹⁷ and when this is used to replace the native Fd in
cyanobacteria it prompts massive CET ³⁹⁸. Pea plants also possess an Fd iso-protein that is expressed
only in times of high ATP demand ³⁹⁹⁻⁴⁰¹ and when this is over-expressed in tobacco it also results in
elevated CET ⁴⁰². At this point it is not clear whether these CET specific Fds operate in the antimycin
A sensitive or insensitive pathways.

After the characterisation of Fd-dependent CET by Arnon and colleagues ⁴⁰³, the possibility
of direct electron donation from Fd to the *Cytb*<sub>6f</sub> was investigated in isolated spinach thylakoids ⁴⁰⁴.
The authors showed that although *Cytb*-563 (heme *b*<sub>n</sub>) is only slowly reduced by dithionite, addition
of hydrophobic quinone mediators accelerated this reduction, while Fd had no effect. Moreover, the
addition of NADPH and Fd in the dark had little effect on the redox state of the cytochrome. In these
experiments, rapid re-reduction of the oxidized P700 center of PSI was measured in the presence of
Fd and NADPH, indicating that Fd-CET was operating very efficiently. This result indicates that, in
this system at least, Fd is a poor direct electron donor to the *Cytb*<sub>6f</sub>. It remains possible, that direct
electron donation from Fd to heme *c*<sub>n</sub> is downregulated in the dark adapted membranes used in this
experiment, or that it might not result in reduction of the heme *b*<sub>n</sub>. Interestingly, the broken chloroplast
assay, used extensively to characterize the *pgr5* and *pgrL1* mutant *Arabidopsis* plants ²³⁵,²³⁶ does not
work if attempted on washed thylakoid membranes (personal observation, GH). The possibility
therefore also remains that components only weakly associated with the *Cytb*<sub>6f</sub> complex are required
to regulate direct Fd: *Cytb*<sub>6f</sub> interaction, and the obvious candidates for such proteins are *pgr5* and
*pgrL1*, although documentation of the role of these proteins in competent electron transport reactions
is minimal and their presence sub-stoichiometric. ²⁰¹ If this is not the case then further experiments
are necessary to identify 1) the interaction partners and 2) their mechanism of function.
As part of a possible model involving direct electron donation from Fd to the Cytb6f<sup>374</sup>, one can propose a docking model of Fd binding to the *n* side of the Cytb6f. Although binding sites are notoriously difficult to predict, interactions between Fd and Fd-dependent enzymes are characterized by charge interactions between the proteins, whose purpose is to optimally orient the redox centers for electron transfer <sup>405-407</sup>. For example, Figure 15 shows the Fd-binding sites (highlighted in black) on two classical Fd-dependent enzymes (C, nitrite reductase; D, FNR). There is an obvious ring of basic charges around the active center to which Fd can dock. When the *n*-facing surface of the Cytb6f is examined, such a patch does appear to exist on the *C. reinhardtii* complex, used in the model proposed by Nawrocki et al.<sup>374</sup>, highlighted with a hashed circle (Figure 15 B). However, such a mechanism would be expected to be universal throughout CET performing organisms, and when the same surface of the cyanobacterial *M. Laminosus* complex is examined (Figure 15 A) the location of such a binding site is much less clear, although co-evolution of Fd iso-proteins with the Cytb6f complex and auxiliary subunits could result in differences in Fd-Cytb6f interactions.

<figure><img src="image_19.png" /></figure>

**Figure 15.** Comparison of charge distribution on the *n* side surface of the Cytb6f with known Fd-binding sites. A) Cytb6f monomer from *M. laminosa* (taken from 2D2C), seen as looking down from the *n* side B) Cytb6f monomer from *C. reinhardtii* (taken from 1Q90), seen as looking down from the *n* side. C) Fd-dependent nitrite reductase (2AKJ), looking down on the Fd binding site, as defined by site directed mutagenesis studies. D) Fe:NADP(H) oxidoreductase (FNR, 1GAW), looking down on the Fd binding site, as defined by site directed mutagenesis studies and co-crystalization. Known Fd binding sites are highlighted with a black circle. The putative Fd binding site on Cytb6f suggested by Nawrocki et al (2019)<sup>374</sup> is indicated by a hashed line. Positively charged groups in blue, negatively charged groups in red.
### 4.3.3 Direct electron donation from FNR to the cytochrome *b*<sub>6f</sub>

Involvement of the FNR enzyme in re-injection of electrons into the quinone pool from the *n* side of the membrane has long been debated, and there is plenty of circumstantial evidence to suggest that this is the case. For example, CET appears inhibited in cyanobacteria when FNR is no longer tethered to the membrane<sup>408</sup>, FNR is more strongly membrane bound in higher plant cells performing enhanced CET<sup>409</sup> and plants with diminished FNR content also show perturbed CET<sup>410</sup>. Moreover, FNR has been consistently co-localised with PgrL1<sup>197,236</sup>, and FNR is no longer recruited efficiently to the membrane in the classic pgr5 mutant of the alga *C. reinhardtii*<sup>411</sup>. Inhibitors of FNR indicate that the enzyme is essential to some CET pathways<sup>233,375,412–416</sup>, although FNR does not bind antimycin A in solution<sup>233</sup>, and is relatively insensitive to this CET inhibitor<sup>417</sup>. Immuno-inhibition studies show that blocking Fd to NADP(H) electron transfer by FNR does not affect CET activity<sup>414,418</sup>, indicating that any CET role for FNR does not involve the NADP<sup>+</sup> binding site. On the other hand, It has been reported that *Cytb*<sub>6f</sub> complexes purified from spinach and incorporated into liposomes are only capable of NADPH mediated quinone reduction if the purification method is mild enough to preserve FNR binding<sup>419</sup>. It should be noted that the severe washes employed to remove FNR in this paper may also have removed other proteins important for this function, such as Fd and/or PgrL1. Importantly, addition of NADPH to the purified FNR:*Cytb*<sub>6f</sub>complex only results in reduction of *Cytb*<sub>6</sub> in the presence of Fd<sup>180</sup>.

Mutant studies on FNR would provide a definitive answer, but are lacking due to the difficulty in fully knocking out the gene for a protein that is essential for photo-autotrophic growth. Plants with minimal FNR downregulate the entire electron transport chain to compensate, leaving an extremely severe phenotype<sup>420</sup>, making interpretation of CET measurements problematic.

The discovery that FNR is co-purified with the *Cytb*<sub>6f</sub> complex<sup>180,421,422</sup> prompted the suggestion that FNR could act as a conduit of electrons from Fd into the *Cytb*<sub>6f</sub>, potentially via the heme *c*<sub>n</sub><sup>180,376</sup>. As with Fd, the absence of information about where and how FNR docks to the *Cytb*<sub>6f</sub> is currently the major impediment to our understanding of a functional mechanism of electron transfer from FNR to the complex. It seems clear that the PgrL1 and Pgr5 proteins are involved in FNR recruitment to the membrane (and therefore probably the *Cytb*<sub>6f</sub>), at least in algae<sup>411</sup>, but in higher plants FNR is recruited to the membrane by two tether proteins, Tic62<sup>423</sup> and TROL<sup>424</sup>. Based on blue native PAGE analysis, it has been suggested that these are the only sites of FNR interaction<sup>424,425</sup>. However, this is likely a function of detergent disruption of interactions between FNR in complex with its tether proteins, and other thylakoid membrane components such as the *Cytb*<sub>6f</sub>. Indeed, in a blue native PAGE experiment using different solubilisation conditions, FNR co-migrated with several
complexes, including the *Cytb*6f <sup>426</sup>. It remains unknown whether the tethering proteins Tic62 and TROL also play a role in FNR association with the *Cytb*6f, or whether release of FNR from these tethers (which happens in response to the generation of ΔpH across the membrane) might prompt association with the *Cytb*6f. Interestingly, only FNR iso-proteins capable of binding to the TROL tether were found co-purified with the *Cytb*6f from maize <sup>422</sup>.

If FNR acts as a conduit of electrons from Fd into the heme c<sub>n</sub>, it prompts some energetic questions. The FAD moiety of FNR can be reduced by one electron from Fd to the flavin radical state (FNR<sub>ox</sub> to FNR<sub>sq</sub>), and a second electron can then fully reduce the FAD (FNR<sub>sq</sub> to FNR<sub>red</sub>), with redox midpoint potentials of -338 mV and -312 mV respectively measured at pH 8 <sup>427</sup>, or NADPH can directly donate two electrons to fully reduce the FAD (FNR<sub>ox</sub> to FNR<sub>red</sub>), with a redox midpoint potential around -380 mV <sup>428,429</sup>. Any of these reduced centers would be adequate to reduce heme c<sub>n</sub>, irrespective of whether the quinone site was occupied or not <sup>430</sup>. So, is the mechanism of FNR to *Cytb*6f reduction a single or double electron transfer to the Q<sub>n</sub> site from FNR? The dependence on Fd of NADPH mediated reduction of *Cytb* <sup>180</sup>, and the seeming independence of CET on the NADPH binding site of FNR <sup>414,418</sup> implies that single electron reduction and oxidation of the FNR FAD moiety is sufficient for CET. It has also been suggested that rather than having an electron transfer function, FNR could simply act as an Fd recruiting module – providing a site for Fd to bind on the *Cytb*6f complex before transfer of electrons directly to heme c<sub>n</sub> or via PgrL1 <sup>199</sup>. A number of experiments are required to clarify the role of FNR recruitment to *Cytb*6f in CET, principally: 1) more information is needed regarding the binding site of FNR on *Cytb*6f, and its orientation; 2) the capacity for CET has to be correctly measured in membranes or cells where the FNR protein has no capacity to interact with the *Cytb*6f or is missing. Finally, if electron donation from Fd or FNR directly to *Cytb*6f is the major route of antimycin A sensitive CET, rates of electron transfer for this complex close to those estimated for the CET pathway – in the range of 10 e<sup>-1</sup> s<sup>-1</sup> PSI<sup>-1</sup> to 130 e<sup>-1</sup> s<sup>-1</sup> PSI<sup>-1</sup> <sup>431</sup> – would be the final proof.

# 5 Inhibitors

## 5.1 Cytochrome *bc*₁ specific inhibitors

Microorganisms often produce and secrete chemical substances in order to restrain or kill other species or strains. As *Cytbc*₁ is a crucial component of the respiratory chain in many bioenergetic systems, its quinone binding catalytic sites have become a critical target in this chemical warfare. Indeed, many compounds that bind to the catalytic sites and act as *Cytbc*₁ inhibitors can be
found in bacteria and fungi. Over the past few decades, these naturally-occurring toxins became popular templates for man-made chemicals, with many proposed functions, from medicinal drugs and treatments for parasitic infections, to fungicides in agriculture.

Depending on the site of action, the Cytbc₁ inhibitors are often divided into two groups. The first group is considered to be small, and consist of compounds with the ability to inhibit the Qᵢ site. The second group consists of the Q₀ site inhibitors and is considered to be much larger, with numerous synthetic compounds developed over the years of research. Moreover, with recent developments, dual-mode inhibitors, capable of binding to both active sites, have been found or synthetized. It is important to note that this classification also excludes the inhibitors that do not act on the quinone-binding sites, such as cyanide, which has been shown to inhibit bacterial cytochrome bc1¹³⁶ or 2,3-dimercaptopropanol (also known as BAL or British anti-Lewisite) which is known as a respiratory inhibitor since the early studies on mitochondrial material⁴³² and later has been shown to destroy the Rieske cluster⁴³³.

*Inhibitors of the Qᵢ site.* The best known compound that inhibits the Qᵢ site is antimycin A. However, this group consists also of naturally occurring ilicicolin H, funiculosin, funiculosin-related compounds, and various synthetic inhibitors (Fig. 16 and 17).

**Antimycin A** (Fig. 16 A) is a depsipeptide with an acyl- (R₁) and alkyl-substituted (R₂) dilactone ring with an amide linkage to a 3-formamidosalicylate moiety. In fact, antimycin A is a whole family of compounds (about 29)⁴³⁴, produced by actinobacteria from the *Streptomyces* genus, with the first members isolated and identified in 1948.⁴³⁵,⁴³⁶ Antimycin A1 has a high binding affinity to the Qᵢ site, with a dissociation constant (K_d) of 32 pM in bovine Cytbc₁.⁴³⁷ It has been shown that antimycin is able to displace quinones and even other inhibitors from the active site.⁴³⁸⁻⁴⁴⁰ Based on the crystal structures ⁶³,⁷⁹,¹²³ it is known that the 3-formamidosalicylate group acts as the toxophore and forms hydrogen bonds with highly conserved residues of Qᵢ (Fig. 16 B), including the His, Asp and Lys, that serve as proton donors in Q reduction¹⁶³, and a conserved Ser. The dilactone ring of antimycin appears to control its solubility and hydrophobicity.⁴⁴¹ There have been attempts to create synthetic analogues (example in Fig. 16 C top) with the dilactone replaced by various other moieties⁴⁴²⁻⁴⁴⁴, with promising *in vitro* results for biphenyl and biphenyl ether groups.⁴⁴⁴ Some of synthetic inhibitors where the 3-formamidosalicylate moiety is replaced with benzazoles and the dilactone rings are replaced with a trifluoromethyl-substituted biphenyl ether moiety (Fig. 16 C, middle and bottom), were shown to be almost as potent as the natural antimycin A in *in vitro* inhibitor assays. However, they exhibited a negligible *in vivo* activity because of low uptake into the living cell.⁴⁴² Recently **UK-2A** (Fig. 16 D), which belongs to a group of dilactone compounds isolated from *Streptomyces* sp. 517-02⁴⁴⁵,⁴⁴⁶, has been revisited and shown to bind to the Qᵢ site in a similar mode
to antimycin A.<sup>447</sup> In contrast to antimycin, UK-2A comprises a 4-methoxypyridin-3-ol moiety as its toxophore. Its inhibitory potency was estimated to be about 3-fold lower than that of antimycin.<sup>448</sup>

<figure><img src="image_20.png" /></figure>

**Figure 16.** Structural features of antimycin and related compounds. **A)** Structural formulae of compounds belonging to the antimycin A family. The upper formula represents the general structural elements of antimycins A, the lower is a formula of antimycin A1<sub>b</sub>. **B)** Stick/ribbon representation of binding of antimycin A in the Q<sub>i</sub> site, as revealed by X-ray crystallography. Antimycin A (*pale green sticks*) forms a direct hydrogen bond to <sup>Bt</sup>Asp228 and additional, water-bridged bonds to <sup>Bt</sup>Lys227, <sup>Bt</sup>His201 and <sup>Bt</sup>Ser35 (*gray sticks*). Oxygen and nitrogen atoms are colored *red* and *blue*, respectively. Heme *b*<sub>H</sub> shown in *brown*. The model is based on PDB entry 1PPJ. **C)** Structural formulae of synthetic antimycin analogues. **D)** Structural formula of UK-2A. In **A**, **C** and **D**, *pale-red*, *dotted* frames indicate the toxophore moiety of each compound.

**Funiculosin** (Fig. 17 A top) is an N-methyl-4-hydroxy-2-pyridone derivative, substituted with a cyclopentanetetrol moiety. It is a natural compound with antifungal properties, isolated from *Penicillium funiculosum*.<sup>449,450</sup> Funiculosin was shown to inhibit Cytbc<sub>1</sub> by a similar mechanism to antimycin A.<sup>164,451,452</sup> However, there is no structural data on funiculosin binding mode to the Q<sub>i</sub> site. It is speculated that it binds in close proximity to heme *b*<sub>H</sub>, as a rise in *E*<sub>m</sub> of this heme by about 100 mV was observed in the inhibited enzyme.<sup>164</sup> Moreover, studies on yeast mutants of <sup>Sc</sup>Asp208 revealed that this residue can possibly be involved in funiculosin binding, which distinguishes it from antimycin.<sup>452</sup> Recently, funiculosin analogues were shown to be produced by other microorganisms, for example the compound **AS2077715** (Fig. 17 A, upper middle), isolated from fungal strain *Capnodium* sp. 33985. The AS2077715 has been shown to exhibit selective and highly inhibitory properties towards *Trichophyton mentagrophytes* Cytbc<sub>1</sub>, while being a weak inhibitor of the
mammalian counterpart. Therefore, it has been proposed as a potential drug for *Trichophyton*
infections.⁴⁵³

Another 2-pyridone deriviative, **Ilicicolin H** (Fig. 17 A, lower middle), was isolated from *Cylindrocladium ilicicola*⁴⁵⁴ and *Gliocladium roseum*.⁴⁵⁵ Despite some structural differences to funiculosins, ilicicolin was shown to act in similar way, being a potent and broad-spectrum antifungal agent⁴⁵⁶, inhibiting yeast and fungal *Cytbc*₁ with high selectivity.⁴³⁹,⁴⁵⁶ Thus, a 2-pyridone moiety is the suggested toxophore in both funiculosins and ilicicolins.⁴³⁹ However, a closer examination of the influence of antimycin A, funiculosin and ilicicolin H on *Cytbc*₁ isolated from distinct organisms and yeast Q₁-site mutants, reveals some differences in sensitivity between the enzymes, which suggests that there is some variation in the binding mode of these inhibitors.⁴⁵⁷

The same 2-pyridone moiety as in funiculosin and ilicicolin H can be found in **sambutoxin** (Fig. 17 A, bottom) and related compounds isolated from *Fusarium sambucinum*⁴⁵⁸,⁴⁵⁹ and other strains of *Fusarium*.⁴⁶⁰,⁴⁶¹ Sambutoxins have been also shown to be toxic to mitochondrial respiration and specific towards *Cytbc*₁.⁴⁵⁹ Although their binding mode was not specified, they can be expected to bind to the Qᵢ site as well. More recently, several synthetic antimalarial compounds comprising a 4-pyridone ring with a biphenyl ether tail were also shown to inhibit the Qᵢ site, eg. **GSK932121**, **GW844520** (Fig. 17 B, top and middle)⁴⁶² or **MJM170** (Fig. 17 B, bottom)⁴⁶³. The co-crystal structures of bovine enzyme with these three novel inhibitors indicate that they all bind close to heme *b*H, with the carbonyl group of the pyridone toxophore in a direct contact to the conserved *BtSer35* ⁴⁶²,⁴⁶³.
<figure><img src="image_21.png" /></figure>

**Figure 17.** Structural formulae of Qᵢ site inhibitors: A) Naturally occurring 4-hydroxy-2-pyridone derivatives. B) Synthetic 4-pyridone-based antimalarial drugs. C) Diuron. D) Sulfonamide-based commercial fungicides. *Pale-red*, *dotted* frames indicate the toxophore moiety of each compound.

**Diuron** (Fig. 17 C), 3-(3,4-dichlorophenyl)-1,1-dimethylurea, is a relatively simple synthetic compound which was shown to have strong inhibitory properties in photosynthesis and weak in mitochondrial respiration.⁴⁶⁴ No structural data is available for its binding mode. The mechanism of action, through inhibition of the Qᵢ site, is known from genetic analysis of diurone- and antimycin-resistant mutations in the yeast model⁴⁶⁵,⁴⁶⁶ and comparative kinetic studies.⁴⁶⁷

**Cyazofamid** (Fig. 17 D left) and **amisulbrom** (Fig. 17 D right) are commercial, synthetic compounds which are imidazole and triazole sulfonamides, respectively. Both are fungicides with high selectivity towards oomycetes.⁴⁶⁸⁻⁴⁷⁰ Cyazofamid was shown to be almost inactive towards yeast, rat liver and potato tuber *Cytbc*₁⁴⁷⁰ and a weak inhibitor of porcine *Cytbc*₁ Qᵢ site when compared to antimycin A.⁴³⁷ Little is known about the binding mode of these inhibitors to the Qᵢ site.
However, based on computational docking studies, it has been suggested that cyazofamid forms a
hydrogen bond to $^{Bt}$Asp228.$^{437}$

*Inhibitors of the Q<sub>o</sub> site.* They are commonly divided into two subclasses. The compounds belonging to the first subclass, of which stigmatellin is the best known example, are capable of immobilizing the ISP-HD in a position near the Q<sub>o</sub> site (in the *b*-position). The inhibitors belonging to the second subclass, such as myxothiazol or MOA-type compounds, bind deeper in the Q<sub>o</sub> site cavity and do not fix the ISP-HD position. Therefore, these two subclasses are often refered to as P<sub>f</sub> (with *f* originating from “fixed”) and P<sub>m</sub> (with *m* originating from “mobile”) inhibitors, respectively. The influence of these inhibitors on movement of ISP-HD is explained in detail in section 3.4.

**Stigmatellin A** (Fig. 18 A, upper) is a natural compound isolated from the myxobacterium *Stigmatella aurantiaca*⁴⁷¹ and the inhibitor of Q₀ site exhibiting the lowest K_d (<0.01 nM).⁴⁷² It consists of a chromone moiety, acting as its toxophore, substituted with a lipophylic alkenyl chain. Crystallographic studies have revealed that stigmatellin fixes the position of the ISP-HD near the Q₀ site, by formation of hydrogen bond between carbonyl oxygen of chromone ring and one of the His residues acting as a ligand to the 2Fe2S (Fig. 18 B).⁶¹,⁶³,¹²³,⁴⁷³ The second hydrogen bond is formed between the phenolic hydroxyl group of the compound and the carboxylate side chain of a conserved Glu residue (belonging to the PEWY motif) of the *Cytb*.⁶³,⁴⁷⁴ The alkenyl tail of stigmatellin is not involved in any specific interactions, but studies on synthetic analogs indicate that modification of this tail can decrease the inhibitory potency of the compound.⁴⁷² However, such a modification was also made in the case of **tridecyl-stigmatellin** (Fig. 18 A, lower), which has the native tail replaced with tridecyl chain but retains its potency. It was developed as a stigmatellin replacement and a potential inhibitor of *Cytb*₅f.⁴⁷⁵ Tridecyl-stigmatellin has been shown to bind to the *Cytb*₅f at both active sites⁶⁶ and also to inhibit the Q₀ site of the bacterial *Cytbc*₁¹⁵¹,²⁶³,⁴⁷² in similar way to the natural compound.
<figure><img src="image_22.png" /></figure>

Figure 18. Q₀ site inhibitors that fix ISP-HD at *b*-position. **A)** Structural formulae of natural stigmatellin A and synthetic tridecyl-stigmatellin. *Pale-red, dotted* frames indicate the toxophore moiety of each compound. **B)** Stick/cartoon representation of binding of stigmatellin A in the Q₀ site, as revealed by X-ray crystallography. Stigmatellin A (*yellow sticks*) forms hydrogen bonds to <sup>Bt</sup>Glu271 of Cytb (*gray sticks*) and to <sup>Bt</sup>His161 of ISP (*pale cyan sticks*). Oxygen and nitrogen atoms are colored *red* and *blue*, respectively. Heme *b*<sub>L</sub> is shown in *brown*. Model is based on PDB entry 1PPJ. **C)** Structural formulae of atovaquone (left) and DBMIB (right). The toxophore of atovaquone is marked with a frame.

**Atovaquone** (Fig. 18 C left) is 2-hydroxynaphthoquinone substituted with a cyclohexyl-chlorophenyl moiety, which was developed as a drug for malaria in 1990s.<sup>476</sup> Among numerous other naphthoquinone derivatives created at the time, it was shown to be the most active against apicomplexan parasites, including several species of *Plasmodium*.<sup>441,476-478</sup> It is the only drug against *Cytbc*₁ currently in clinical use.<sup>441</sup> Atovaquone inhibits the enzyme of *P. falciparum* with nanomolar affinity,<sup>476,479,480</sup> and has much lower toxicity to the human counterpart.<sup>480</sup> The results of early studies, based on EPR and redox potentiometry, indicated that atovaquone has a similar binding mode to stigmatellin, and also arrests ISP-HD at *b*-position.<sup>479</sup> Moreover, based on the complementary molecular modeling studies, atovaquone was suggested to bind within the Q₀ site, forming a hydrogen bond between its hydroxyl group and the 2Fe2S cluster-ligating His residue, and a water bridge between the carbonyl oxygen at position 1 of naphthoquinone ring and the conserved Glu of PEWY motif.<sup>479</sup> However, crystallographic studies on yeast showed that the only hydrogen bond is formed
between the ionized hydroxyl group of the inhibitor and the Nε atom of ⁵CHis181. The ⁵CGlu272 residue side chain was shown to face the opposite direction with respect to the carbonyl oxygen, and does not appear to be involved in atovaquone binding.⁷¹

**DBMIB** (Fig. 18 C, right), 2,5-dibromothymoquinone, is a halogenated benzoquinone derivative. Although DBMIB is known more as a Cytb6f inhibitor¹¹⁰ (see section 5.2), it has been shown to exhibit similar potency against Cytbc₁.⁴⁸¹ DBMIB should not be considered as a regular inhibitor competing with QH₂, as has been commonly assumed. Rather, it binds to the Q₀ site in the form of a semiquinone and there forms a spin-coupled state with 2Fe2S, which leads to locking of the ISP-HD at *b*-position.⁴⁸² (see section 7.6). The spin-coupled state can be detected in EPR experiments as the prominent g = 1.94 EPR signal in X-band at low temperatures.⁸⁴,¹⁰⁸,¹¹⁰,⁴⁸³

**UHDBT** and **HHDBT** (Fig. 19 A) are both 6-hydroxy-4,7-benzothiazolediones (HDBT) with undecyl and heptyl tails, respectively. HDBT compounds are synthetic inhibitors capable of binding to the Q₀ site.⁷⁰,¹¹⁶,⁴⁸⁴,⁴⁸⁵ As they comprise the hydroxybenzoquinone moiety, they exist as a mixture of ortho- and para-quinone tautomers (Fig. 19 B). The 6-hydroxy group is ionizable, with a pKₐ of 6.5, which is reflected in a distinctive color change from yellow to violet when pH changes from acidic to alkaline.⁴⁸⁶ These compounds are also redox active and, as is typical for quinones, the Eₘ exhibits a pH dependence with a slope of about 60 mV/pH unit.⁴⁸⁶ Thus, HDBTs may be regarded as substrate analogues. However, the Eₘ value of UHDBT was shown to be about -40 mV at pH 7⁴⁸⁶, thus much lower than the ubiquinone pool (about 100 mV).⁴⁸⁷,⁴⁸⁸ Cytochrome bc₁ inhibition by UHDBT was found to be pH dependent, with a 10-fold decrease in the binding rate constant at more alkaline pH.⁴⁸⁵ This had been initially associated with ionization of the hydroxyl moiety. However, this effect was also noted for 6-bromo substituted UHDBT.⁴⁸⁴ As indicated by the results of X-ray crystallography, HDBT compounds immobilize the ISP-HD in the *b*-position, via formation of a hydrogen bond with one of His ligands of the 2Fe2S cluster.⁷⁰,⁷⁷,⁴⁷⁴ However, there is no agreement on the detailed mode of hydrogen bonding for HHDBT and UHDBT within the Q₀ site. Based on the structure of yeast Cytbc₁ co-crystalized with HHDBT, hydrogen bonds are proposed to form between: 1) the oxygen from the ionized hydroxyl group of HHDBT and the protonated Nε of ⁵CHis181 of the ISP, 2) the carbonyl O4 atom and the water molecule that is itself bonded to the backbone nitrogen of the Glu within the PEWY motif.⁷⁰ A slightly different model has been proposed in the case of bovine Cytbc₁ interaction with UHDBT. It was suggested that the protonated His of ISP interacts with the carbonyl O7, while its deprotonated form interacts with the hydroxyl group of the inhibitor. Moreover, the carbonyl O4 and N3 of UHDBT were suggested to form a hydrogen bond with the hydroxyl group of the conserved ⁸Tyr131 of the Cytb (⁵Tyr132).⁷⁷
<figure><img src="image_23.png" /><figcaption>**Figure 19.** Structural features of 6-hydroxy-4,7-benzothiazolediones. **A)** General structural formulae of *n*HDBT compounds. **B)** Possible tautomeric states of *n*HDBT compounds.</figcaption></figure>

**Crocacin A and D** (Fig. 20 A, upper and middle) are naturally occurring dipeptide derivatives, comprising a Z-enamide moiety of glycine and either a aminohexadienoic or aminohexenoic acid (respectively), substituted with a polyketide acyl tail. They belong to a whole family of compounds isolated from the myxobacterium *Chondromyces crocatus*.⁴⁸⁹⁻⁴⁹¹ Some of the crocacins exhibit inhibitory properties against many strains of bacteria, yeasts and fungal plant pathogens, with (+)-crocacin D recognized as the most potent compound of the family.⁷⁵,⁴⁹¹ In studies on bovine *Cytbc*₁ in SMP's it has been shown that the inhibitory mechanism of crocacins is based on binding to the Q₀ site.⁴⁸⁹ The model for binding of crocacin A and D was developed based on EPR studies and crystallographic analysis of the crocacin D analogue (Fig. 20 A, bottom) bound to the avian *Cytbc*₁, combined with molecular modeling and docking studies. Crocacins were shown to immobilize the ISP-HD in the *b*-position, with the Z-enamide carbonyl forming a hydrogen bond to Nε of the 2Fe2S-ligating His. Moreover, it was suggested that the glycine residue of crocacin can form hydrogen bonds to the backbone nitrogen with the PEWY motif Glu and to the carbonyl group of the conserved Met (⁸¹Met138) of *Cytb*. However, the existence of a hydrogen bond to the Met residue is not supported by the crystal structure. Interestingly, the Z-enamide seems to adopt a hairpin conformation, with an intramolecular hydrogen bond between the two amide groups.⁷⁵

Because natural crocacins have been shown to exhibit a photostability of minutes under field conditions,⁷⁵ they are not suitable for use in agriculture, despite their inhibitory potency. However, in
recent years, several methods for synthesis of crocacins⁴⁹¹,⁴⁹² and their simplified, more stable
analogues have been developed.⁴⁹³

<figure><img src="image_24.png" /></figure>

**Figure 20.** Crocacins and famoxadone-like compounds. A) Structural formulae of natural crocacins A and D (top and middle, respectively) and synthetic derivative of crocacin D (bottom). B) Structural formulae of famoxadone (upper), JG144 (lower, left) and fenamidone (lower, right). *Pale-red, dotted* frames indicate the toxophore moiety of each compound.

**Famoxadone, JG144 and fenamidone** (Fig. 20 B) are synthetic fungicides. The toxophores of famoxadone and JG144 comprise 1,3-oxazolidine-2,4-dione, and that of fenamidone comprises imidazol-4-one ⁷⁸,¹¹⁷,⁴⁹⁴,⁴⁹⁵. The famoxadone molecule is chiral, and the S-(-)-enantiomer has been shown to be active against *Cytbc*₁.⁴⁹⁶ The crystal structures of famoxadone ⁷⁸,¹¹⁷,⁴⁹⁵ and its analogs⁷⁸,¹¹⁷,⁴⁹⁵ indicate that these inhibitors bind to the Q₀ site in the same fashion, with carbonyl oxygen at ring position 4 forming a hydrogen bond to the backbone nitrogen of the PEWY motif Glu. This resembles the binding mode of MOA-like inhibitors, but in contrast to those, famoxadone-like inhibitors are capable of immobilizing the ISP-HD near the Q₀ site, close to *b*-position (Pf inhibitors subclass).⁴⁷⁴ However, famoxadone and its analogs do not form a hydrogen bond with the His of the ISP as in the case of other ISP-fixing compounds.⁷⁸,¹¹⁷ Rather, the inhibitor binding is thought to induce conformational changes, which result in formation of a hydrogen bond between the 2Fe2S His ligand and a highly conserved Tyr residue of the *ef*α-helix of *Cytb*, thus immobilizing the ISP-HD in the near *b*-position.¹¹⁷