Signal-Switchable Electrochemical Systems E-Book

Evgeny Katz

• Herausgeber: Wiley-VCH Verlag GmbH & Co. KGaA
• Kategorie: Wissenschaft und neue Technologien
• Sprache: Englisch

Seitenzahl: 620

Veröffentlichungsjahr: 2018

Beschreibung

A guide to the biological control over electronic systems that lead the way to wearable electronics and improved drug delivery

In recent years, this area of electrochemical systems has developed rapidly and achieved significant progress. Signal-Switchable Electrochemical Systems offers an overview to the wide-variety of switchable electrochemical systems and modified electrodes. The author?a noted researcher and expert on the topic?summarizes research efforts of many groups in a range of universities and countries. The book explores various types of external signals that are able to modify electrode interfaces, for example electrical potential, magnetic field, light, as well as chemical and biochemical inputs.

Multifunctional properties of the modified interfaces allow their responses to complex combinations of external signals. These are integrated with unconventional biomolecular computing systems logically processing multiple biochemical signals. This approach allows the biological control over electronic systems. The text explores the applications in different areas, including unconventional computing, biofuel cells and signal-triggered molecular release in electrochemical systems. This important guide:

-Provides an overview to the biological control over electronic systems and examines the key applications in biomedicine, electrochemical energy conversion and signal-processing
-Offers an important text written by a highly cited researcher and pioneer in the field
-Contains a summary of research efforts of an international panel of scholars representing various universities and countries
-Presents a groundbreaking book that provides an introduction to this interdisciplinary field

Written for scientists working with electrochemical systems and applications with signal-responsive materials, Signal-Switchable Electrochemical Systems presents an overview of the multidisciplinary field of adaptable signal-controlled electrochemical systems and processes and highlights their key aspects and future perspectives.

Leseprobe

Table of Contents

Cover

Preface

References

Chapter 1 Introduction

References

Chapter 2 Magneto‐switchable Electrodes and Electrochemical Systems

2.1 Introduction

2.2 Lateral Translocation of Magnetic Micro/nanospecies on Electrodes and Electrode Arrays

2.3 Vertical Translocation of Magnetic Micro/Nanospecies to and from Electrode Surfaces

2.4 Assembling Conducting Nanowires from Magnetic Nanoparticles in the Presence of External Magnetic Field

2.5 Vertical Translocation of Magnetic Hydrophobic Nanoparticles to and from Electrode Surfaces

2.6 Repositioning and Reorientation of Magnetic Nanowires on Electrode Surfaces

2.7 Integration of Magnetic Nanoparticles into Polymer‐Composite Materials

2.8 Conclusions and Perspectives

2.9 Appendix: Synthesis and Properties of Magnetic Particles and Nanowires

References

Chapter 3 Modified Electrodes and Electrochemical Systems Switchable by Temperature Changes

3.1 Introduction

3.2 Thermo‐sensitive Polymers with Coil‐to‐Globule Transition

3.3 Electrode Surfaces Modified with Thermo‐sensitive Polymers for Temperature‐controlled Electrochemical and Bioelectrochemical Processes

3.4 Electrode Surfaces Modified with Multicomponent Systems Combining Thermo‐sensitive Polymers with pH‐, Photo‐ and Potential‐Switchable Elements

3.5 Electrodes Modified with Thermo‐switchable Polymer Films Containing Entrapped Metal Nanoparticles – Inverted Temperature‐dependent Switching

3.6 Conclusions and Perspectives

References

Chapter 4 Modified Electrodes and Electrochemical Systems Switchable by Light Signals

4.1 Introduction

4.2 Diarylethene‐based Photoelectrochemical Switches

4.3 Phenoxynaphthacenequinone‐based Photoelectrochemical Switches

4.4 Azobenzene‐based Photoelectrochemical Switches

4.5 Spiropyran–merocyanine‐based Photoelectrochemical Switches

4.6 Conclusions and Perspectives

References

Chapter 5 Modified Electrodes Switchable by Applied Potentials Resulting in Electrochemical Transformations at Functional Interfaces

References

Chapter 6 Electrochemical Systems Switchable by pH Changes

6.1 Introduction

6.2 Monolayer Modified Electrodes with Electrochemical and Electrocatalytic Activity Controlled by pH Value

6.3 Polymer‐Brush‐Modified Electrodes with Bioelectrocatalytic Activity Controlled by pH Value

6.4 pH‐Controlled Electrode Interfaces Coupled with

in situ

Produced pH Changes Generated by Enzyme Reactions

6.5 pH‐Triggered Disassembly of Biomolecular Complexes on Surfaces Resulting in Electrode Activation

6.6 pH‐Stimulated Biomolecule Release from Polymer‐Brush Modified Electrodes

6.7 Conclusions and Perspectives

References

Chapter 7 Coupling of Switchable Electrodes and Electrochemical Processes with Biomolecular Computing Systems

7.1 Introduction

7.2 Electrochemical Analysis of Output Signals Generated by Enzyme Logic Systems

7.3 Summary

References

Chapter 8 Biofuel Cells with Switchable/Tunable Power Output as an Example of Implantable Bioelectronic Devices

8.1 General Introduction: Bioelectronics and Implantable Electronics

8.2 More Specific Introduction: Harvesting Power from Biological Sources – Implantable Biofuel Cells

8.3 Biofuel Cells with Switchable/Tunable Power Output

8.4 Summary

References

Chapter 9 Signal‐triggered Release of Biomolecules from Alginate‐modified Electrodes

9.1 Introduction – Signal‐activated Biomolecular Release Processes

9.2 Alginate Polymer Cross‐linked with Fe Cations – The Convenient Matrix for Molecular Release Stimulated by Electrochemical Signal

9.3 Self‐operating Release Systems Based on the Alginate Electrodes Integrated with Biosensing Electrodes

9.4 Conclusions and Perspectives

References

Chapter 10 What is Next? Molecular Biology Brings New Ideas

10.1 Switchable Enzymes and Their Use in Bioelectrochemical Systems – Motivation and Applications

10.2 Electrocatalytic Function of the Ca‐Switchable PQQ‐GDH‐CaM Chimeric Enzyme

10.3 Integration of the Ca‐Switchable PQQ‐GDH‐CaM Chimeric Enzyme with a Semiconductor Chip

10.4 A Ca‐Switchable Biofuel Cell Based on the PQQ‐GDH‐CaM Chimeric Enzyme

10.5 Substance Release System Activated with Ca Cations and Based on the PQQ‐GDH‐CaM Chimeric Enzyme

10.6 Summary

References

Chapter 11 Summary and Outlook: Scaling up the Complexity of Signal‐processing Systems and Foreseeing New Applications

References

Index

End User License Agreement

List of Illustrations

Chapter 02

Figure 2.1 Magneto‐controlled switchable bioelectrocatalytic process – a general concept.

Figure 2.2 Reversible switching of the DNA oxidation upon magneto‐induced lateral translocation of the DNA‐functionalized Fe

3

O

4

magnetic particles. Chronopotentiometric responses of the “left” and “right” electrodes are shown in the presence and absence of the particles. Amount of magnetic particles, 100 μg; DNA‐oligomer (

1

) structure is shown (note that the used oligonucleotide is rich with oxidizable guanine bases); pre‐treatment potential, +1.7 V for 10 s; stripping current, +5 μA (between 0.6 and 1.2 V vs Ag/AgCl reference electrode).

Figure 2.3 (A) Modification of magnetic Fe

3

O

4

microparticles (ca. 1 μm diameter) with naphthoquinone (

2

). (B) Magneto‐controlled patterning of a Au electrode surface upon formation of an insoluble product (

4

) of a biocatalytic reaction triggered by the electrocatalytic formation of H

2

O

2

in the presence of naphthoquinone (

2

)‐functionalized magnetic particles. (C) Pattern produced on the Au electrode by the electrocatalytic process using the naphthoquinone (

2

)‐functionalized magnetic particles, 10 mg, HRP, 1 mg mL

−1

, and substrate (

3

), 3 × 10

−4

M. Background electrolyte: 0.1 M tris‐buffer, pH 7.5, was saturated with air. The potential, −0.5 V (vs SCE), was applied for 3 min on the electrode to produce the first spot, then the potential was switched off, the magnetic particles were moved by the external magnet, and the potential −0.5 V was reapplied for 3 min to produce the second spot.

Figure 2.4 (A) Magneto‐controlled electrobiochemiluminescence. Note that the light emission was observed only when the quinone‐functionalized magnetic particles were located on the electrode surface. (B) Cyclic voltammograms of: (a) the cystamine‐modified Au electrode under Ar (dashed line); (b) the naphthoquinone (

2

)‐modified particles (10 mg) magnetically attracted to the electrode surface under Ar (solid line); (c) the cystamine‐modified Au electrode under air in the absence of magnetic particles; and (d) the naphthoquinone (

2

)‐modified particles (10 mg) magnetically attracted to the electrode surface under air. The vertical dash line shows the potential of −0.4 V (vs SCE), which was applied to induce the electrobioluminescence – compare electrocatalytic (curve d) and non‐catalytic (curve c) currents at this potential. All experiments were recorded in 0.1 M phosphate buffer, pH 7.0, potential scan rate, 10 mV s

−1

. (C) Magneto‐switchable electrocatalytic generation of biochemiluminescence in a system consisting of the naphthoquinone (

2

)‐functionalized magnetic particles (10 mg), luminol (1 μM), and HRP (1 mg mL

−1

). Top curves: chronoamperometric transients upon application of potential steps from 0.0 to −0.4 V (vs SCE) on the cystamine‐modified Au electrode: (a) current transient measured when the naphthoquinone (

2

)‐functionalized particles are positioned on the Au electrode by the external magnet; (b) current transient measured when the particles are translocated to the nonconductive glass support by the external magnet. Note that the short current pulse observed in the absence of the quinone‐modified particles corresponds mostly to the capacitance current. Bottom curves: light (measured by the light detector as power) emitted from the system upon the application of the respective chronoamperometric transients: (c, d) in the presence and absence of the particles on the Au electrode, respectively. Note that the light emission intensity correlates with the produced current. All measurements were performed in 0.1 M phosphate buffer, pH 7.0, system equilibrated with air. (D) Structures of luminol, 3‐aminophthalate (excited state) and naphthoquinone (

2

)‐functionalized magnetic particles.

Figure 2.5 (A) Reversible activation and deactivation of electrocatalytic processes on a screen‐printed electrode (SPE) upon magneto‐induced lateral translocation of iron‐doped thermally reduced graphene oxide (Fe‐TRGO). Ascorbic acid (AA,

7

) is electrocatalytically oxidized to dehydroascorbic acid (DAA,

8

) when Fe‐TRGO is located on the SPE surface (left) and non‐catalytically oxidized when Fe‐TRGO is removed from the electrode surface (right). (B) Cyclic voltammogram corresponding to the AA electrocatalytic oxidation in the presence of Fe‐TRGO. (C) Cyclic voltammogram corresponding to the AA non‐catalytic oxidation in the absence of Fe‐TRGO (note smaller current and more positive anodic peak comparing with the cyclic voltammogram show in B). Potential scan rate, 100 mV s

−1

. The potentials are shown vs Ag/AgCl reference electrode. (D) Reversible variation of the peak current upon activation (ON) and deactivation (OFF) of the electrocatalytic process. (C) Reversible variation of the peak potential upon activation (ON) and deactivation (OFF) of the electrocatalytic process.

Figure 2.6 (A) Reversible activation/inhibition of the electrochemical reaction of PQQ (

9

) covalently bound to Fe

3

O

4

microparticles (ca. 1 μm diameter) upon their vertical translocation up and down with help of an external magnet. Note that the PQQ‐modified magnetic particles are disconnected from the electrode surface in the up‐position of the magnet, while their translocation down results in their deposition on the electrode surface. (B) Cyclic voltammograms corresponding to the electrode surface with the magnetically attracted PQQ‐modified particles (a) and to the electrode surface after their magneto‐induced removal (b); the small wave observed in the latter cyclic voltammogram (b) is due to the incomplete removal of the particles. The data were obtained in Tris buffer (0.1 M, pH 8.0) in the presence of CaCl

2

(10 mM) under argon. Potential scan rate, 100 mV s

−1

. The potentials are shown vs SCE. (C) Dependence of the peak‐to‐peak separation, Δ

E

p

, on the potential scan rate,

v

, observed in the cyclic voltammograms of PQQ‐functionalized magnetic particles attracted to the Au electrode. Inset: dependence of the peak current, I

p

, on the potential scan rate.

Figure 2.7 Reversible activation/inhibition of electrochemical reactions of the redox species covalently bound to Fe

3

O

4

microparticles (ca. 1 μm diameter) upon their vertical translocation with help of an external magnet: translocation to and from the electrode surface is shown for naphthoquinone (

2

)‐modified particles (A) and ferrocene (

10

)‐modified particles (B) in the left and right schemes, respectively. (C) Differential pulse voltammograms (DPVs) recorded for the naphthoquinone (

2

)‐modified particles magnetically attracted to the electrode surface (curve a) and removed from it (curve b). (D) DPVs recorded for the ferrocene (

10

)‐modified particles magnetically attracted to the electrode surface (curve a) and removed from it (curve b). Potential scan rate, 20 mV s

−1

. The potentials are shown vs SCE. Insets show reversible variation of the peak current upon stepwise attraction of the particles to (a) and removal from (b) the electrode surface.

Figure 2.8 Reversible activation/inhibition of electrocatalytic reactions mediated by the redox species covalently bound to Fe

3

O

4

microparticles (ca. 1 μm diameter) upon their vertical translocation with help of an external magnet: translocation to and from the electrode surface is shown for PQQ‐modified particles in the presence of NADH (A) and ferrocene‐modified particles in the presence of GOx and glucose (B) in the left and right schemes, respectively. (C) Cyclic voltammograms recorded for the PQQ‐modified particles in the presence of NADH (50 mM) magnetically attracted to the electrode surface (curve a) and removed from it (curve b). Potential scan rate, 10 mV s

−1

. (D) Cyclic voltammograms recorded for the ferrocene‐modified particles in the presence of GOx (1 mg mL

−1

) and glucose (10 mM) magnetically attracted to the electrode surface (curve a) and removed from it (curve b). Potential scan rate, 5 mV s

−1

. The potentials are shown vs SCE. Insets show reversible variation of the electrocatalytic current upon stepwise attraction of the particles to (a) and removal from (b) the electrode surface.

Figure 2.9 (A) Magneto‐switched bioelectrocatalytic oxidation of lactate in the presence of lactate dehydrogenase (LDH) and magnetic particles functionalized with PQQ‐NAD

+

. (B) Formation of pyruvate upon the bioelectrocatalytic oxidation of lactate (0.1 M) in the presence of LDH (2 mg mL

−1

) and PQQ‐NAD

+

‐functionalized magnetic particles with a potential of

E

 = 0.05 V (vs SCE) applied on the electrode. Domains started from points “a”: the magnetic particles are attracted to the electrode surface. Domains started from points “b”: the magnetic particles are retracted from the electrode surface. Inset: reversible changes in the rate of the bioelectrocatalytic formation of pyruvate upon the magnetic switching the biocatalytic process ON and OFF by attraction and retraction of the magnetic particles to and from the electrode surface, respectively. The data were recorded in Tris buffer (0.1 M, pH 7.0) and CaCl

2

(10 mM) under argon.

Figure 2.10 Magneto‐controlled selective biosensing of glucose or lactate in the presence of glucose oxidase (GOx), lactate dehydrogenase (LDH), magnetic particles functionalized with PQQ‐NAD

+

, and a Au electrode modified with a monolayer of ferrocene‐mediator units (

12

). Note that the system is activated for the lactate sensing in the “down” magnet position and for the glucose sensing in the “up” magnet position (the left and right parts of the scheme, respectively). Insets (top left) show amperometric analysis of glucose and lactate for the “up” and “down” magnet positions corresponding to the magnetic particles removed from and attracted to the electrode surface, respectively. Note that the system responds to the glucose variable concentrations and is unaffected by the lactate concentrations when the magnet is up (the magnetic particles are removed from the electrode surface); the potential applied on the electrode was 0.5 V (vs SCE). On the contrary, the system responds to the lactate variable concentrations and is unaffected by the glucose concentrations when the magnet is down (the magnetic particles are attracted to the electrode surface); the potential applied on the electrode was 0.05 V (vs SCE). Note the different potentials applied on the electrode for the “up” and “down” magnet positions.

Figure 2.11 (A) Dual magneto‐chemical switching ON–OFF electrode activity: the scheme shows changes of the P4VP‐polymer brush between hydrophilic swollen and hydrophobic collapsed states upon

in situ

pH changes produced at the surface by GOx‐functionalized magnetic nanoparticles in the presence of glucose. (B) Impedance spectra (Nyquist plots) for the ABTS redox probe obtained for the electrode in the ON and OFF states (bias potential, 0.62 V vs Ag/AgCl reference). Note the different scales for the impedance measured in the ON and OFF states. The inset shows stepwise changes of

R

et

upon reversible transition between the ON and OFF states.

Figure 2.12 Magneto‐switchable electrochemical reaction observed with a single Fe

3

O

4

magnetic microparticle modified with Prussian blue redox shell. (A) Magneto‐controlled vertical translocation of a single magnetic particle between the electrode surface‐state and solution‐suspended state (electrochemical reaction ON and OFF, respectively). (B) Quartz‐crystal microbalance (QCM) analysis of the microparticle translocation. (C) Cyclic voltammograms (CVs) measured when the microparticle is connected to (a) and disconnected from (b) the electrode surface. Potential scan rate, 25 mV s

−1

. (D) DPV obtained for the microparticle contacting the electrode surface. Potential scan rate, 25 mV s

−1

. (E) Chronoamperometric analysis showing the electrochemical process activation/deactivation (on/off) in real time. Applied potential, 0.12 V vs pseudo‐Ag reference.

Figure 2.13 (A) Formation of the conducting nanowires upon self‐assembling of the Au‐coated CoFe

2

O

4

magnetic nanoparticles along the magnetic field lines and their use as a nanostructured electrode for electrochemical oxidation of ferrocene monocarboxylic acid (Fc;

16

) coupled with the glucose oxidation biocatalyzed by GOx. (B) Cyclic voltammograms of ferrocene monocarboxylic acid (

16

), 0.1 mM, obtained before (a) and after (b) formation of self‐assembled nanowires (in the absence and presence of the magnetic field, respectively). Potential scan rate, 200 mV s

−1

. (C) AFM image of an individual nanowire self‐assembled from the Au‐coated magnetic nanoparticles in the presence of the external magnetic field. (D) AFM image of the nanostructured array obtained on the electrode surface upon magneto‐assisted self‐assembling of the Au‐coated magnetic nanoparticles. (Note that the AFM imaging was performed in the absence of the magnetic field when the original alignment of the nanowires along the magnetic field lines was not preserved.) The cross‐section of the nanostructured array is shown at the bottom.

Figure 2.14 Reversible formation of the hydrophobic thin film on the electrode surface by deposition of hydrophobic magnetic nanoparticles. Note that the magnetic nanoparticles transport toluene molecules entrapped in the hydrophobic shall. (A, B) Faradaic impedance spectra (Nyquist plots) of the electrode in the presence and absence of the hydrophobic magnetic nanoparticles. (A) High impedance measured on the electrode surface coated with the thin film of hydrophobic nanoparticles magnetically attracted to the surface. (B) Low impedance measured on the electrode surface facing the aqueous background solution after magneto‐induced removal of the hydrophobic magnetic nanoparticles. (Note different scales for the impedance spectra shown in A and B.) (C) Reversible changes of the electron transfer resistance,

R

et

, upon stepwise deposition–removal of the hydrophobic magnetic nanoparticles. (D) Reversible changes of the double‐layer capacitance,

C

dl

, upon stepwise deposition–removal of the hydrophobic magnetic nanoparticles. The impedance measurements were performed in the presence of a 1 mM (1 : 1) K

3

[Fe(CN)

6

]/K

4

[Fe(CN)

6

] mixture and upon biasing the working electrode at 0.17 V vs Ag/AgCl reference.

Figure 2.15 (A) The scheme showing reversible translocation of hydrophobic magnetic nanoparticles between the modified electrode surface and toluene phase. (B) Cyclic voltammograms obtained in the absence (a) and presence (b) of hydrophobic magnetic nanoparticles on the ferrocene (

12

) monolayer‐modified electrode surface (note the presence of a quinone derivative (

17

) soluble in the aqueous phase). Potential scan rate, 100 mV s

−1

. (C, D) Analysis of peak currents obtained at different potential scan rates,

v

: I ∝

v

characteristic of surface‐confined electrochemical process; I ∝

v

1/2

characteristic of diffusional electrochemical process.

Figure 2.16 (A) The scheme showing reversible translocation of hydrophobic magnetic nanoparticles between the modified electrode surface and toluene phase. (B) Cyclic voltammograms obtained in the absence (a) and presence (b) of hydrophobic magnetic nanoparticles on the quinone (

18

) monolayer‐modified electrode surface. (Note the presence of a ferrocene monocarboxylic acid (

16

) soluble in the aqueous phase.) Potential scan rate, 100 mV s

−1

. (C–D) Analysis of peak currents obtained at different potential scan rates,

v

: I ∝

v

characteristic of surface‐confined electrochemical process; I ∝

v

1/2

characteristic of diffusional electrochemical process.

Figure 2.17 (A) Magneto‐controlled reversible ON–OFF switching of the bioelectrocatalytic oxidation of glucose by GOx using the hydrophobic magnetic nanoparticles. (B) Cyclic voltamograms of the system consisting of the surface‐confined ferrocene monolayer (

12

), GOx, 1 mg mL

−1

, and glucose, 80 mM, dissolved in the aqueous phase: (a) when the magnetic nanoparticles are retracted from the electrode surface and (b) when the magnetic nanoparticles are attracted to the electrode surface. The data were recorded under Ar in a biphase system composed of 0.1 M phosphate buffer, pH 7.0 (lower phase), and toluene with the magnetic nanoparticles, 1 mg mL

−1

(upper phase). Potential scan rate 5 mV s

−1

. Inset: the reversible switch of the current generated by the system at

E

 = 0.5 V vs SCE. (a) The magnetic nanoparticles are retracted from the electrode surface. (b) The magnetic nanoparticles are attracted to the electrode surface.

Figure 2.18 (A) Magneto‐controlled reversible ON–OFF switching of microperoxidase‐catalyzed reduction of cumene hydroperoxide (

20

) by means of the hydrophobic magnetic nanoparticles. (B) Cyclic voltammograms of the system consisting of the surface‐confined microperoxidase (

19

) and cumene hydroperoxide (

20

), 2 mM, dissolved in the toluene layer: (a) when the magnetic nanoparticles are retracted from the electrode surface and (b) when the magnetic nanoparticles are attracted to the electrode surface. The data were recorded under Ar in a biphase system composed of 0.1 M phosphate buffer, pH 7.0 (lower phase), and toluene with the magnetic nanoparticles, 1 mg mL

−1

(upper phase). Potential scan rate 5 mV s

−1

. Inset: the reversible switch of the current generated by the system at

E

 = −0.7 V vs SCE. (a) The magnetic nanoparticles are retracted from the electrode surface. (b) The magnetic nanoparticles are attracted to the electrode surface.

Figure 2.19 (A) Magneto‐switchable anodic/cathodic currents generated at the electrode functionalized with microperoxidase (MP;

19

) by using hydrophobic magnetic nanoparticles to gate the bioelectrocatalytic processes. (B) Cyclic voltammograms recorded in the presence of the MP‐functionalized electrode when the magnetic nanoparticles are retracted from the surface: (a) in the absence of H

2

O

2

and (b) in the presence of H

2

O

2

(50 mM); potential scan rate 5 mV s

−1

. Inset: cyclic voltammogram of the MP‐functionalized electrode recorded in the absence of H

2

O

2

; potential scan rate of 100 mV s

−1

; note the different potential range. (C) Cyclic voltammograms recorded in the presence of the MP‐functionalized electrode when the magnetic nanoparticles are attracted to the surface: (a) in the absence of decamethylferrocene (

22

) in the toluene phase, (b) in the presence of

22

(2 mM) in the toluene phase and the absence of glucose in the aqueous phase, and (c) in the presence of

22

(2 mM) in the toluene phase and glucose (50 mM) in the aqueous phase; potential scan rate 5 mV s

−1

. Vertical dash line shows the potential applied for the current measurements in D. (D) Switchable bioelectrocatalytic currents generated in the system upon application of a fixed potential of 0.25 V vs SCE, in the presence of H

2

O

2

(50 mM) and glucose (50 mM) in the aqueous phase and

22

(2 mM) in the toluene phase, when the magnetic nanoparticles are retracted from the surface (a) or attracted to the surface (b). The data were recorded in a solution of phosphate buffer (0.1 M, pH 7.0) containing GOx (1 mg mL

−1

).

Figure 2.20 (A) Magneto‐switchable anodic/cathodic photocurrents generated at the electrode functionalized with CdS nanoparticles (5 nm diameter) by using hydrophobic magnetic nanoparticles to gate the photoelectrochemical processes. (B) Switchable photocurrents generated in the system upon (a) retraction from the electrode and (b) attraction to the electrode of the magnetic nanoparticles associated with cumene hydroperoxide (

20

). The data were recorded in the solution of phosphate buffer (0.1 M, pH 7.0) containing TEOA (20 mM) upon application of a potential of 0 V (vs SCE) and irradiation of the electrode with visible light. (A semitransparent electrode with a very thin layer of Au was irradiated from the bottom.)

Figure 2.21 (A) Magneto‐switchable selective sensing of glucose or lactate using the hydrophobic magnetic nanoparticles for gating the bioelectrocatalytic processes. Left: the magnetic nanoparticles retracted from the electrode provide lactate sensing through the LDH biocatalyzed reaction mediated by PQQ‐NAD

+

‐monolayer proceeding at the electrode facing the aqueous solution. Right: the magnetic nanoparticles attracted to the electrode provide glucose sensing through the GOx biocatalyzed reaction mediated by decamethylferrocene (

20

) shuttling electrons through a hydrophobic thin film. Note that glucose oxidation and lactate oxidation are inhibited in the former and latter states, respectively. (B, C) Cyclic voltammograms recorded at the PQQ‐NAD

+

‐monolayer functionalized Au electrode in the presence of GOx and LDH in the aqueous solution and hydrophobic magnetic NPs in the toluene phase: (B) Upon retraction of the magnetic nanoparticles from the electrode surface and in the presence of different concentrations of lactate in the aqueous phase: (a) 0 mM, (b) 20 mM, and (c) 80 mM. (C) Upon attraction of the magnetic nanoparticles to the electrode surface: (a) in the absence of decamethylferrocene (

20

) in the toluene phase and in the presence of glucose, 50 mM, and lactate, 50 mM, in the aqueous phase. Note that the bioelectrocatalytic process is not activated in the absence of the decamethylferrocene mediator. In the presence of

20

in the toluene phase and in the presence of different concentrations of glucose in the aqueous phase: (b) 0 mM, (c) 20 mM, and (d) 80 mM.

Figure 2.22 (A) Magneto‐controlled quantum charging of the Au nanoparticle array associated with a Au electrode in the presence of hydrophobic magnetic nanoparticles and two‐phase liquid solution. (B) Linear sweep voltammograms recorded in the presence of: (a) magnetic nanoparticles retracted from the Au nanoparticle/thiol monolayer‐functionalized electrode (note the different current scale in the inset); (b) magnetic nanoparticles attracted to the Au nanoparticle/thiol monolayer‐functionalized electrode (the quantized charging peaks are marked by asterisks); (c) magnetic nanoparticles attracted to the thiol monolayer‐modified electrode in the absence of Au nanoparticles. Note the small capacitance, but the absence of the quantized charging peaks. The data were obtained under Ar in a biphase system consisting of 0.1 M phosphate buffer, pH 7.0, and toluene with the hydrophobic magnetic nanoparticles, 1 mg mL

−1

. Potential scan rate, 2 mV s

−1

. Arrows show the directions of the potential sweeps. (C) Plot of the variation of the formal charging potentials as a function of the Au nanoparticle charging states obtained from the linear sweep voltammogram shown in panel B, curve (b).

Figure 2.23 (A)

Scanning electron micrograph

,

SEM

, (top view) image of a typical hexagonally ordered nanoporous alumina template with a pore diameter of 70 nm and an interpore distance of 100 nm. (B) SEM cross‐sectional view of alumina membranes filled with Fe nanowires deposited from electrolytes containing: (a) 0.1 M FeSO

4

, (b) 1 M FeSO

4

, and (c) 0.5 M FeSO

4

 + 0.4 M H

3

BO

3

. (C) Schematic description of the membrane‐template electrochemical preparation of multifunctional nanowires.

Figure 2.24 Magneto‐controlled reversible activation–inhibition of electrochemical processes upon reorientation of adaptive magnetic nanowires functionalized with self‐assembled long‐chain alkanethiol monolayer (C

18

alkanethiol was used in the experiments; the scheme shows a shorter chain for illustration only). Also shown are the optical images (top view) of the glassy carbon disk electrode covered with the vertically (left) and horizontally (right) aligned nanowires.

Figure 2.25 (A) Magneto‐controlled reversible activation–inhibition of electrocatalytic glucose oxidation upon reorientation of adaptive magnetic Ni nanowires. (B) Amperometric response showing switchable current corresponding to glucose (1 mM) oxidation electrocatalyzed by the Ni nanowires oriented in the vertical (V) and horizontal (H) positions. Potential, +0.85 V (vs Ag/AgCl). (C) Tuning of the electrode activity through control of the angle of the nanowire orientation (a–e): amperometric response for 1 mM glucose recorded while changing slowly the orientation of the nanowires from the horizontal (a) through vertical (c), and back to horizontal (e).

Figure 2.26 (A) An SEM image of grouped Au/Ni nanowires illustrating the relative polarities of the nickel segment (a bright segment is gold). Scale bar: 1 μm. (B) Cyclic voltammograms corresponding to glucose (20 mM) oxidation mediated by the surface‐confined ferrocene and biocatalyzed by the GOx‐functionalized Au/Ni nanowires in the active‐horizontal position, low activity‐vertical position and “off”‐lifted position. Potential scan rate, 10 mV s

−1

. (C) Nanowire‐based magneto‐switchable bioelectrocatalytic processes shown schematically. In the experimental setup with the GOx‐functionalized Au/Ni nanowires and the ferrocene‐modified surface, the magnetic field can be oriented in the horizontal, vertical and “off” (lifted) positions, for activating, hindering, and blocking the communication between the nanowire‐confined GOx and the surface‐confined ferrocene relay.

Figure 2.27 (A) Magneto‐switchable redox properties of the composite material containing γ‐Fe

2

O

3

‐Au core–shell nanoparticles coated with lipoic acid and included in polyaniline (PAn) thin film. (B) Faradaic impedance spectra (Nyquist plots) corresponding to: (a) The PAn/nanoparticle composite film in the absence of the external magnet; (b) The PAn/nanoparticle film subjected to the external magnet located below the electrode surface; and (c) The control system consisting of PAn/polyacrylic acid film (note the absence of magnetic nanoparticles) subjected to the external magnet. Inset: interfacial electron transfer resistance,

R

et

, corresponding to the PAn/nanoparticle film in the absence (a) and presence (b) of the external magnet. Data recorded in phosphate buffer (0.2 M, pH 7.3) with the bias potential 0.04 V (vs SCE).

Figure 2.28 Cyclic voltammograms obtained on the electrode modified with γ‐Fe

2

O

3

‐Au core–shell nanoparticles coated with lipoic acid and included in polyaniline (PAn) thin film: (a) in the presence of GOx, without glucose; (b) in the presence of GOx and glucose, 80 mM, in the absence of the external magnet; and (c) in the presence of GOx and glucose, 80 mM, in the presence of the external magnet. All data were recorded in 0.2 M phosphate buffer, pH 7.3, in the presence of GOx (2 mg mL

−1

), scan rate 5 mV s

−1

, under argon.

Figure 2.29 Various magnetic nanoparticles coated with gold shells: (a–d) TEM images of Fe

3

O

4

‐core/Au‐shell magnetic nanoparticles synthesized according to different experimental procedures: (a) [140], (b) [141], (c) [140, 141], (d) [142]; see more details in [93]. (e, f) TEM and

scanning transmission electron microscopy

(

STEM

) images, respectively, of the γ‐Fe

2

O

3

‐core/Au‐shell magnetic nanoparticles [8].

Figure 2.30 Various magnetic nanoparticles coated with silica shells: backscattered electrons image (a) and TEM image (b) of Fe

3

O

4

‐core/SiO

2

‐mesoporous‐shell magnetic nanoparticles. TEM image (c) of Fe

3

O

4

‐core/SiO

2

‐mesoporous‐shell magnetic nanoparticles. TEM image (d) of Fe

3

O

4

‐core/SiO

2

‐shell magnetic nanoparticles.

Figure 2.31 (a, b) TEM images of Fe

3

O

4

‐Ag and FePt‐Ag hetero‐dimers composed for the magnetic nanoparticle and connected Ag nanoparticle. (c) Directed functionalization of the Fe

3

O

4

nanoparticle and Ag nanoparticle with different functional units, such as dopamine‐derivatized and thiol‐derivatized species, respectively. X and Y might be represented by different molecular and biomolecular species.

Figure 2.32

Field emission scanning electron microscope

(

FESEM

)

images of released strontium ferrite magnetic nanowires with diameter of (a) 60, (b) 50, (c) 40, and (d) 30 nm, after removal of alumina templates.

Figure 2.33 Scanning electron micrograph (SEM) images of Ni nanowires with average diameter of 98 nm and length of 17 μm after removal of alumina templates.

Chapter 03

Figure 3.1 Temperature‐controlled switchable electrochemical processes – a general concept.

Figure 3.2 (a) Poly(

N

‐isopropylacrylamide)‐based polymer

(

1

) undergoes rapid gelation from a soluble liquid at room temperature, (b) to form a stable, nonshrinking gel at body temperature (37 °C), (c) after 1 min.

Figure 3.3 Typical LCST polymers

are based on

N

‐isopropylacrylamide (NIPAM)

(

2

),

N

,

N

‐diethylacrylamide (DEAM) (

3

), methyl vinyl ether (MVE)

(

4

), and

N

‐vinylcaprolactam (NVCL)

(

5

) as monomers. A typical UCST system is based on a combination of acrylamide (AAm)

(

6

) and acrylic acid (AAc) (

7

). The monomers shown in the figure are used for preparation of thermo‐sensitive polymers.

Figure 3.4 Cyclic voltammograms of the PNIPAM‐brush modified

ITO electrode obtained in the presence of 1 mM [Fe(CN)

6

]

3−

in 0.1 M phosphate buffer (pH 7.0) and 0.1 M sodium perchlorate at different temperatures: (a) 12 °C and (b) 42 °C. Inset: Reversible changes of the peak current,

I

p

, derived from the cyclic voltammograms upon stepwise temperature changes: steps 1 and 3 correspond to the OFF state at 42 °C, while steps 2 and 4 correspond to the ON state at 12 °C. Potential scan rate, 10 mV s

−1

. Scheme at right shows the temperature‐controlled electrochemical reaction with the [Fe(CN)

6

]

3−

diffusional redox probe.

Figure 3.5 (A) Temperature‐switchable electrochemical reaction of soluble ferrocene monocarboxylic acid (Fc,

9

) on the PG electrode surface

modified with the PNIPAM (

8

) hydrogel film containing GOx. (B) Cyclic voltammograms of Fc, 0.5 mM, obtained on the PNIPAM‐GOx‐modified electrode at different temperatures; potential scan rate, 50 mV s

−1

. Inset: reversible changes of the electrochemical response (anodic peak current,

I

pa

, derived from the cyclic voltammograms) upon cyclic temperature changes between 25 and 37 °C. (C) Reversible time‐dependent changes of the

I

pa

after switching temperature from 37 to 25 °C and back. (D)

I

pa

measured at different temperature: (a) in the absence of Na

2

SO

4

and (b) in the presence of Na

2

SO

4

, 0.3 mM. Note the LCST shift to the lower temperature in the presence of Na

2

SO

4

. (E) SEM images of the PNIPAM‐GOx film on the PG electrode after treatment with a buffer solution, pH 7.0, at 25 °C (a) and 37 °C (b). Note much higher density and less porosity observed for the hydrophobic collapsed polymer film at 37 °C.

Figure 3.6 (A) Temperature‐switchable bioelectrocatalytic oxidation of glucose (Glc) biocatalyzed by GOx and mediated by soluble ferrocene monocarboxylic acid (Fc,

9

) on the PG electrode surface modified with the PNIPAM (

8

) hydrogel film containing GOx. (B) Cyclic voltammograms obtained on the PNIPAM‐GOx‐modified electrode

in the presence of Fc, 0.5 mM, and Glc, 8 mM: (a) at 25 °C in the absence of Na

2

SO

4

and methanol, (b) at 37 °C in the absence of Na

2

SO

4

and methanol, (c) at 25 °C with 0.3 M Na

2

SO

4

but without methanol, and (d) at 25 °C with 20% methanol but in the absence of Na

2

SO

4

. Potential scan rate, 10 mV s

−1

; pH 7.0.

Figure 3.7 (A) Temperature‐controlled bioelectrocatalytic oxidation of glucose biocatalyzed by soluble PQQ‐GDH and mediated by ferrocene (Fc) units

associated with the thermo‐sensitive polymer covalently bound to a thiolated monolayer self‐assembled on a Au electrode. (B) Bioelectrocatalytic current measured at different temperatures on the Au electrode modified with the PNIPAM‐Fc‐derivative in the presence of PQQ‐GDH, 10 nM, and glucose, 10 mM, in the solution. Applied potential was +350 mV vs Ag/AgCl.

Figure 3.8 (A) Schematics of Pt electrode modification with an interpenetrating‐polymer network (IPN)

composed of PAA and PNIPAM. The electrochemically induced radical polymerization was performed in two steps, first to produce PAA and then to generate PNIPAM, steps A and B, respectively. (B) Cyclic voltammograms obtained with the PAA/PNIPAM‐IPN‐modified Pt electrode in the presence of soluble Fc(MeOH)

2

(

11

) at 20 °C and different pH values: (a) pH 6.5 and (b) pH 2.7. (C) Peak current,

I

p

, dependence on the pH value of the electrolyte solution (measured at 20 °C). (D) Cyclic voltammograms obtained on the PAA/PNIPAM‐IPN‐modified Pt electrode in the presence of soluble Fc(MeOH)

2

(

11

) at pH 6.5 and different temperatures: (a) 20 °C; (b) 45 °C. (E) Peak current,

I

p

, dependence on temperature of the electrolyte solution (measured at pH 6.5).

Figure 3.9 SEM images of the Pt electrode modified with the interpenetrating‐polymer network (IPN)

composed of PAA and PNIPAM. (A) Intact polymer film. (B, C) The electrode surface partially uncovered by scratching the polymer film. Note different magnification in the images shown in (A–C). Insets in (A) and (C) show the polymer film composed of PAA only (the second polymerization step was not performed).

Figure 3.10 (A) Photoisomerization and protonation–deprotonation equilibrium of spiropyran

and merocyanine. Reversible transformations between the four states: spiropyran (SP),

merocyanine

(

MC

), protonated merocyanine (MCH

+

), and

protonated spiropyran

(

SPH

+

). (B) Schematic representation of light‐controlled reorganization of thermo‐responsive polymers functionalized with photoisomerizable spiropyran/merocyanine. Photochemical conversion of the hydrophilic MCH

+

to the hydrophobic SP induces dehydration of the polymer chain, which initiates the phase transition from expanded to folded states. Note that the process is reversible and the polymer can be returned back to the expanded state by the opposite photochemical reaction upon

ultraviolet

(

UV

) irradiation

. Note that in the text merocyanine

is abbreviated as MR instead of MC abbreviation used in this figure.

Figure 3.11 (A) Faradaic impedance spectra obtained on an SP‐functionalized PNIPAM‐modified Au electrode in the presence of [Fe(CN)6]

3−/4−

, 2 mM, at variable temperature: (a) 20 °C, (b) 26 °C, (c) 28 °C, (d) 30 °C, (e) 32 °C, (f) 34 °C, (g) 36 °C, and (h) 40 °C. (B) Faradaic impedance spectra under the same conditions as in (A) for the MR

+−

‐functionalized PNIPAM‐modified electrode, at: (a) 20 °C, (b) 22 °C, (c) 24 °C, (d) 26 °C, (e) 28 °C, (f) 34 °C, (g) 36 °C, (h) 38 °C, (i) 40 °C, and (j) 44 °C. (C) Interfacial electron‐transfer resistances,

R

et

, derived from the impedance spectra measured at different temperatures for: (a) the SP‐PNIPAM‐modified electrode and (b) the MR

+−

‐PNIPAM‐modified electrode. (D) Cyclic changes in the interfacial electron‐transfer resistance,

R

et

, upon the photochemically induced transition between: (a) the SP‐PNIPAM‐modified electrode and (b) the MR

+−

‐PNIPAM‐modified electrode at

t

 = 36 °C. Data were recorded in an aqueous NaNO

3

background solution (0.2 M, pH 9.5).

Figure 3.12 (A) Electrochemically induced radical polymerization resulting in the formation of an interpenetrating‐polymer PNIPAM network, followed by its modification with photoisomerizable SP and catalytic Pt NPs. Reversible transformation of the PNIPAM network between the swollen and collapsed states upon photoisomerization of SP/MR

+−

species. The collapsed hydrophobic state (in the presence of SP) is electrochemically inactive, while the swollen hydrophilic state (in the presence of MR

+−

) is electrocatalytically active for oxidation of ascorbic acid (

14

) to dehydroascorbic acid (

15

). The reversible photochemical transformations were obtained at 38 °C. (B) The polymer network‐modified electrode was electrochemically inactive at

t

 > 42 °C and electrocatalytically active at

t

 <°25 °C regardless of the SP/MR

+−

states.

Figure 3.13 Linear sweep voltammograms corresponding to the electrocatalytic oxidation of ascorbic acid measured on: (a) SP‐PNIPAM‐Pt‐NPs‐modified electrode and (b) MR

+−

‐PNIPAM‐Pt‐NPs‐modified electrode. Inset: Switchable electrocatalytic current measured for the different isomeric states of the SP/MR

+−

species in the polymer network. Measurements were recorded in an aqueous NaNO

3

background solution (0.2 M, pH 9.5) that included ascorbic acid (2.5 mM) at

t

 = 38 °C. Scan rate, 10 mV s

−1

.

Figure 3.14 (A) Multisignal switchable composite film based on PNIPAM with entrapped graphene or graphene oxide

depending on the potential applied on the Au electrode. The modified electrode was switched between low and high electrochemical activity by four input signals: pH variation, applied potential, temperature changes, and addition of Na

2

SO

4

salt. (B–E) Reversible variation of the anodic peak current,

I

pa

, upon changing applied potential, pH, temperature, and salt concentration. The

I

pa

values were extracted from cyclic voltammograms measured in the presence of 0.5 mM ferrocene dicarboxylic acid (

16

) at potential scan rate of 100 mV s

−1

.

Figure 3.15 Cyclic voltammograms corresponding to the high (a) and low (b) bioelectrocatalytic activity of the modified electrode (PNIPAM with entrapped graphene) in the presence of ferrocene dicarboxylic acid (0.5 mM), GOx (1 mg ml

−1

), and glucose (6 mM). The bioelectrocatalytic activity was controlled by complex combinations of four input signals: pH, potential, temperature, Na

2

SO

4

.

Figure 3.16 Anodic bioelectrocatalytic current,

I

a

, derived from cyclic voltammograms measured in the presence of ferrocene dicarboxylic acid (0.5 mM), GOx (1 mg ml

−1

), and glucose (6 mM) upon various combinations of four input signals (pH, potential, temperature, Na

2

SO

4

). The scheme shows the equivalent logic circuit corresponding to the switchable system. The dash line shows the threshold separating high and low current values defined as logic outputs

1

and

0

, respectively.

Figure 3.17 (A) Reversible transformation of the Cu‐NPs‐PNIPAM composite film between states with high and low conductivity (i.e. electrochemically active and inactive states) at high (40 °C) and low (22 °C) temperatures, respectively. (B) Faradaic impedance spectra (Nyquist plots) obtained with the modified electrode at 22 and 40 °C, demonstrating the high and low

R

et

, respectively. Cyclic temperature change allowed reversible variation of the

R

et

, thus activating and inhibiting electrochemical reactions. The inset shows temperature‐dependent changes of the

R

et

. The impedance spectra were measured in the presence of [Ru(NH

3

)

6

]

2+/3+

, 1 mM, with the bias potential of −0.15 V (vs SCE).

Figure 3.18 Reversible operation of the immune‐sensor based on the PNIPAM polymer covalently attached to an anti‐TnT layer. Cyclic operation of the immune‐sensor was achieved due to temperature‐controlled re‐configuration of PNIPAM between two different states: collapsed globular state at 37 °C and extended coil state at 25 °C.

Chapter 04

Figure 4.1 Light‐controlled switchable electrochemical processes based on photoisomerizable molecules – general concept.

Figure 4.2 Photoisomerizable species frequently used in photoswitchable electrochemical systems: (A) phenoxynaphthacenequinones, (B) diarylethenes, (C) azobenzenes, and (D) spiropyran/merocyanine derivatives. The shown structures are examples only and they can vary in different systems.

Figure 4.3 (A) Absorbance spectra of the photochromic diarylethene derivative in two isomeric states measured in an acetonitrile solution: (a) colorless open‐ring state, (b) colored closed‐ring state. (B) Cyclic voltammograms demonstrating redox features of the diarylethene derivative: (a) The initial scan (pink) was started from the open‐ring state, which does not show any redox activity until ca. 1.1 V potential is reached. The irreversible oxidation of the open‐ring state results in its cyclization through intermediate formation of a double oxidized open‐ring form (O

2+

). Then, the electrochemically produced closed‐ring form can be reversibly reduced–oxidized demonstrating single (C

+

) and double (C

2+

) oxidation states. (b) The cyclic voltammograms demonstrating reversible oxidation of the closed‐ring form in two consecutive steps. The

reversible transition between the closed‐ring and open‐ring states can be achieved by photochemical isomerization applying visible and UV irradiation. The cyclic voltammetry was performed in an acetonitrile solution with 0.1 M NBu

4

ClO

4

, potential scan rate 0.5 V s

−1

.

Figure 4.4 (A) Reversible photochemical isomerization

of the diarylethene spacer separating redox subunits represented by phenol groups or quaternized pyridinium groups. (B) Reversible photo‐induced switching between OFF and ON redox states when the photoisomerizable linker changed between non‐conjugated and conjugated forms, respectively.

Figure 4.5 Viologen molecules

: dimethylviologen (the most frequently used electron transfer mediator with a low redox potential) and caroviologen (an example of a redox molecule with a long conjugated spacer connecting the redox subunits). Note that these molecules are not photoisomerizable.

Figure 4.6 (A) Photoswitchable transformatio

n of the diarylethene functionalized with quaternized pyridinium groups between open and closed structures, (

1a

) and (

1b

), (UV irradiation at

λ

 = 365 nm; Vis illumination at

λ

 > 600 nm). (B) OFF and ON redox states achieved for the open (non‐conjugated) and closed (conjugated) isomeric states. (C) Cyclic voltammograms obtained for the open (a) and closed (b) isomeric states, 1 mM in acetonitrile solution in the presence of 0.1 M NBu

4

BF

4

supporting electrolyte; potentials measured vs SCE.

Figure 4.7 Absorbance spectra of the open (a) and

closed (b) isomeric forms of the photoswitchable viologen measured in an acetonitrile solution. The observed optical changes accompany the reversible redox transformations.

Figure 4.8 (A) Reversible photo‐induced isomerization

of the diarylethene derivative functionalized with phenol groups connected through a photoisomerizable spacer. Then, the closed isomeric form (

2b

) demonstrates reversible redox features corresponding to the quinone/hydroquinone transformations

. Notably, the open‐ring isomer (

2a

) is not electrochemically active. (B) Cyclic voltammograms obtained for the electrochemically inactive (OFF) state (a) and electrochemically active (ON) state (b), 1 mM in acetonitrile (with 2% v/v H

2

O) with 0.1 M NBu

4

BF

4

supporting electrolyte, potential scan rate 200 mV s

−1

, potentials measured vs SCE.

Figure 4.9 An example of a photoswitchable molecular wire

with a long distance separating pyridinium groups. The open‐ring isomeric form corresponds to the redox OFF state, while the closed‐ring form corresponds to the electrochemically active ON state.

Figure 4.10 Schematic representation of the conductance measurements performed with an STM for a single molecule with the photoswitchable molecular wire. A break junction is formed by pushing a gold probe tip into a gold surface covered with dithiolated dithienylethene molecules, retracting it and recording current as molecules become transiently trapped in the gap.

Figure 4.11 Assembly of the photo/electro‐switchable monolayer

composed of the pyridinium derivative of diarylethene covalently attached to a cysteamine self‐assembled monolayer on a Au electrode surface.

Figure 4.12 Operation of the monolayer‐immobilized pyridinium derivative

of diarylethene as set–reset flip‐flop memory unit. (A) Cyclic voltammograms of the diarylethene modified Au electrode: (a) in the original redox active open‐ring state, (b) after application of +0.35 V for 90 s converting the diarylethene to the redox inactive closed‐ring form, (c) after irradiation at 570 nm for 15 min converting diarylethene back to the open‐ring isomer. The cyclic voltammograms were recorded in phosphate buffer, pH = 7.2, at 100 mV s

−1

. Inset: Peak current values extracted from the cyclic voltammograms upon repeating the steps described in (A). (B) Write‐Read‐Erase

steps based on electrochemical/photochemical isomerization of the diarylethene monolayer. (C) The set–reset flip‐flop memory unit – a general scheme: S and R refer to set and reset signals, Q and Q

next

refer to initial and next state of the memory unit. (D) Schematic operation of the set–reset flip‐flop memory unit. The scheme shows the memory state changes upon applying different combinations of the set–reset signals. (E) Truth table for operation of the set–reset flip‐flop memory unit, where

X

corresponds to either

0

or

1

state.

Figure 4.13 A multielectrode array (A) and

multichannel electrochemical device (B) (Bronjo Medi; https://bronjo.com/multi‐channel‐systems‐mea2100‐systems/) potentially applicable to the Write‐Read‐Erase electrochemical/photochemical systems processing and storing electronic and optical signals. Note that different multielectrode arrays can be applied to operate with the discussed switchable systems. The present device is only an example.

Figure 4.14 Photo/electro‐switchable electron transport

through the diarylethene molecular “wire” from the reconstituted glucose oxidase (GOx) to the electrode conducting support. Note that the open‐ring form of the diarylethene is redox active and supports the electron transfer process, while the closed‐ring isomeric form is redox inactive and thus inhibits the electron transport, thus, switching OFF the bioelectrocatalytic process. The amino‐functionalized FAD synthetic derivative (

5

) is shown in the frame. Glc and

GlcA

are glucose and

gluconic acid

(product of glucose oxidation).

Figure 4.15 (A) General scheme illustrating the switchable operation of the reconstituted GOx controlled by electrical and optical signal

s. (B) Cyclic voltammograms obtained with the modified electrode in the presence of glucose, 80 mM: (a) and (c) recorded with the diarylethene molecular “wire” in the redox active open‐ring form, (b) and (d) recorded with the diarylethene molecular “wire” in the redox inactive closed‐ring form. Data were recorded in a 0.1 M phosphate buffer, pH = 7.4, under Ar. Potential scan rate 5 mV s

−1

. Inset: The cyclic changes of the bioelectrocatalytic current extracted from the cyclic voltammogram at

E

 = 0.2 V. (C) Calibration curve corresponding to the bioelectrocatalytic anodic currents at

E

 = 0.2 V (vs SCE) obtained for the modified electrode in the ON state. Glc and GlcA are glucose and gluconic acid (product of glucose oxidation).

Figure 4.16 Assembling a monolayer composed of a diarylethene derivative (di‐(

N

‐butanoic acid‐1,8‐naphthalimide)‐piperazine dithienylethene) (

6

) at a

Au electrode surface. The photoisomerizable monolayer demonstrates selective complex formation with Ag

+

cations in the open‐ring form. The closed‐ring isomer releases the Ag

+

cations. Similar results were obtained for the signal‐controlled complex formation with Cu

2+

cations.

Figure 4.17 (A) High‐resolution XPS spectrum at a normal photoelectron emission angle (0°) corresponding to the complex formed by the diarylethene monolayer with Ag

+

cations. The blue line corresponds to the experimental spectrum and the purple line indicates a deconvolution curve. (B) Cyclic voltammograms obtained with the modified electrode: (a) in the closed‐ring form, which does not form the complex with Ag

+

cations, (b) in the open‐ring form, which produces the complex with

Ag

+

cations (note the peaks corresponding to the redox process of Ag

+

cations associated with the monolayer). The measurements were performed in an aqueous solution of 0.1 M NaNO

3

; potential scan rate 100 mV s

−1

. Inset: Amperometric responses, at

E

 = 0.26 V (vs SCE) obtained upon the cyclic photoisomerization of the modified electrode. After each isomerization step the modified electrode was treated with Ag

+

cations, followed by rinsing to remove the unbound Ag

+

cations. (C) Contact angle changes measured for a droplet of 1 mM Ag

+

solution placed on the monolayer‐modified Au electrode upon cyclic photoisomerization of the monolayer between the open‐ring state (a) and closed‐ring state (b). Note that the open‐ring state, which produces the complex with Ag

+

cations, demonstrates smaller contact angles reflecting more hydrophilic surface.

Figure 4.18 (A) Fluorescent images of the diarylethene‐modified surface obtained with a confocal microscope for different isomeric states of the photoswitchable molecules: (a) the initial non‐fluorescent closed‐ring form, (b) four fluorescent molecules observed after visible irradiation converting the closed‐ring isomer to the open‐ring form, (c) return back to the inactive form upon UV irradiation, (d) activating four fluorescent molecules again with the visible light. (B) Reversible photoisimerization of the studied diarylethene derivative. (C) The time trace of the photoresponse of a single molecule.

Figure 4.19 (A) The reversible photoisomerization process for

trans

‐ (

7a

) and

ana

‐isomeric (

7b

) forms of the phenoxynaphthacenequinone

derivative. (B) Immobilization of the photoisomerazable quinone at a Au electrode surface, including the following steps: (i) self‐assembling of a cystamine monolayer, (ii) carbodiimide coupling of the quinone carboxylic groups with the amino groups of the cystamine monolayer, (iii) formation of a densely packed monolayer by incorporation of the long alkyl thiol into pinholes of the primary monolayer. (C) Cyclic voltammograms obtained for the quinone‐modified electrode in the presence of different isomeric states of the immobilized quinone: (a)

trans

‐ and (b)

ana

‐isomeric forms. The experiments were performed in 0.01 M phosphate buffer including 0.1 M Na

2

SO

4

, pH 7.0, under Ar, scan rate 50 mV s

−1

. Inset: Amperometric responses of the modified electrode extracted from the cyclic voltammograms and measured at

E

 = −650 mV (vs SCE) upon reversible photochemical switching of the monolayer between the

trans

‐quinone and

ana

‐quinone forms.

Figure 4.20 (A) The pH‐dependent re

dox potential

E

° of the immobilized

trans

‐isomer of phenoxynaphthacenequinone. (B) The scheme of the pH‐gated electrochemical reduction of benzyl‐viologen (BV

2+

) mediated by

trans

‐phenoxynaphthacenequinone: The redox potential of the quinone at pH = 8 is negative enough to mediate the electrochemical reduction of BV

2+

; however, the redox potential of the quinone at pH = 4 is not negative enough for reduction of BV

2+

. Note that direct (non‐mediated) reduction of BV

2+

is not possible because of the barrier produced by the densely packed long alkyl thiol on the electrode surface.

Figure 4.21 (A) Photochemically switchable bioelectrocatalytic reduction of nitrate (NO

3

−

). The process is biocatalyzed

by soluble nitrate reductase (NR; 1 U mL

−1

) and the electron transport is mediated by a soluble benzyl‐viologen (BV

2+

; 1 mM) (

8

) and monolayer immobilized phenoxynaphthacenequinone rigidified with C

14

H

29

SH. The structure of benzyl‐viologen is shown in the frame. (B) Cyclic voltammograms corresponding to the bioelectrocatalytic system in the ON state (a) and OFF state (b), when the mediating quinone is in the

trans

‐form and

ana

‐form, respectively. The ON and OFF states are reversibly controlled by the UV and Vis light signals. The experiments were performed in 0.01 M phosphate buffer, pH = 7.5, in the presence of 0.1 M Na

2

SO

4

and 50 mM KNO

3

; potential scan rate 5 mV s

−1

. Inset: Cyclic variation of the electrocatalytic current upon reversible photochemical transformation of the quinone between

trans

‐form (ON state) and

ana

‐form (OFF state).

Figure 4.22 (A) Molecular structures (calculated) of two isomeric forms (

ana

‐ and

trans

‐) of phenoxynaphthacenequinone bridging

single‐walled carbon nanotubes (SWCNTs). (B) Current‐voltage characteristics and ON/OFF ratio of the nanoelectrodes connected through two isomeric forms of the quinone.

Figure 4.23 General scheme demonstrating three effects possibly observed for azobenzene‐functionalized systems upon

cis

‐/

trans

‐photoisomerization

: (i) switching between redox active and inactive states (usually

cis

‐isomer is more redox active), (ii) alternating penetration of external soluble redox species through the monolayer to an electrode surface, and (iii) demonstrating different distances from the electrode surface (measured by STM). To be specific for illustrating purposes, the scheme shows a two‐component mixed thiolated monolayer self‐assembled on a Au electrode; however, in reality the shown effects have been observed for different azobenzene derivatives included in various thin films/monolayers.

Figure 4.24 (A) A Langmuir–Blodgett thin film

deposited on an ITO electrode and reversibly photoisomerized between

trans

‐ (

9a

) and

cis

‐isomeric (

9b

) forms. (B) Cyclic voltammograms obtained for the modified electrode in the presence of the

cis

‐isomer (a) and

trans

‐isomer (b). The experiment was started with the

trans

‐isomer and the

cis

‐isomer

was produced by UV irradiation with a xenon lamp or laser 325 nm) for 1 min. The solution was composed of Britton–Robinson buffer, pH 7.0, with 0.2 M KClO

4

; potential scan rate 20 mV s

−1

.

Figure 4.25 (A) Anodic peaks of the cyclic voltammograms obtained for different areas of the electrode irradiated

with UV light for 10 s. The ITO electrode was modified according to Figure 4.24A. The numbers on the peaks correspond to the numbers of irradiated electrode areas (from 1 to 6) using a He‐Cd laser (beam diameter ca. 1 mm). The inset shows a linear dependence of the number of the photoisomerized azobenzene molecules on the number of the irradiated areas. (B) Chronoamperometric response of the photochemically produced

cis

‐azobenzene derivative with the bias potential of −0.4 V (vs Ag/AgCl). The integrated area under the current response curve corresponds to the charge (Q) of the responding

cis

‐azobenzene derivative molecules. (C) The charge dependence on the irradiation intensity.

Figure 4.26 (A) Covalent immobilization of ferrocene s

pecies bound to the electrode through a photoisomerizable azobenzene linker. First, the ITO electrode was silanized to provide reactive amino groups for further reaction steps. Then, a ferrocene derivative containing an azobenzene group in a side radical (abbreviated as Fc‐azo‐COOH (

10

), see the structure shown in the frame) was covalently bound to the aminated electrode surface using standard carbodiimide reaction in the presence of

dicyclohexyl‐carbodiimide

(

DCC

) and

4‐dimethylaminopyridine

(

DMAP

). (B) Cyclic voltammograms obtained with the modified electrode after different irradiation treatment: (a) the initial form of the electrode containing the

trans

‐state of the azobenzene linker, (b) after UV irradiation for 1 h converting the linker to the

cis

‐state, (c) after visible light (Vis; ambient room light) illumination for 1 day returning (not completely) the electrode to its original state. Note that the redox response corresponds to the ferrocene groups, but not to azobenzene. The experiments were performed using 0.1 M NaCl aqueous supporting electrolyte; potential scan rate 20 mV s

−1

.

Figure 4.27 Atomic force microscopy (AFM)

images of the original bare ITO electrode (A) and ferrocene‐azobenzene‐functionalized electrode (see the structure in Figure 4.26A) (B). (C) Optical absorbance spectra obtained for the transparent modified electrode after different time‐periods of the UV irradiation (color‐coded spectra correspond to 30, 60, 90, and 120 min of the UV irradiation). After that the electrode was illuminated with visible light for 1 day. (D) The differential spectra (absorbance difference) after different time‐periods of the irradiation (extracted from the spectra shown in part C).

Figure 4.28 (A) Schematic illustration of reversible photo‐induced restructuring of the ferrocene‐azobenzene‐functionalized electrode and transport of

the Cl

−

counter‐ions upon the ferrocene redox transformations. (B) and (C) peak potential variation upon changing the potential scan rate in cyclic voltammetry experiments performed with

trans

‐ and

cis

‐forms of the linker, respectively.

E

pa

and

E

pc

correspond to the peak anodic and peak cathodic, respectively; the difference between

E

pa

and

E

pc

corresponds to Δ

E

p

– peak‐to‐peak separation. These dependences were used to derive the corresponding electron‐transfer rate constants (

k

et

) shown in the figures.

Figure 4.29 Study of an electrode modified with a self‐assembled thiol monolayer composed of two components: long‐alkyl thiol functionalized

with azobenzene (

11a, 11b

) and alkyl thiol functionalized with azobenzene and ferrocene (

12

). The monolayer was prepared with the ratio

11a

:

12

 = 99 : 1. (A) Cyclic voltammogram obtained with the modified electrode in the presence of 0.5 mM K

4

[Fe(CN)

6

] – the azobenzene unit was in the

cis

‐form. (B) The same as (A), but the azobenzene unit was in the

trans

‐form obtained by illumination with visible light (

λ

 > 400 nm) for 1 h. The schemes below the cyclic voltammograms explain the electrochemical process, which is dependent on the azobenzene isomeric state. The experiments were performed in an aqueous solution containing 0.2 M NaClO

4

under N

2

at 5 °C; potential scan rate 100 mV s

−1

.

Figure 4.30 (A) Schematic drawing of the electrochemical cell (top) and mass‐transport of probing redox molecules through the photoresponsive nanocomposite membrane assembled on an ITO electrode (bottom). The channel on the left shows diffusion of the redox species through smaller pores with azobenzene ligands in their

trans

‐configuration

; the channel on the right shows diffusion through larger pores with azobenzene ligands in their

cis

‐configuration. The red and green ovals correspond to the ferrocene redox species in their reduced and oxidized states, respectively; the yellow ovals and yellow elongated oval correspond to the azobenzene

cis

‐ and

trans

‐isomers, respectively. WE, CE and RE are working, counter and reference electrodes, respectively. (B) Chronoamperometric measurements (

I

vs

t

) performed with two ferrocene derivatives (FDM and FDMDG; see Figure 4.31C for their structures) penetrating through the photoresponsive nanocomposite film under alternate exposure to UV (360 nm) and visible light (435 nm). Last cycle uses room light, 400–700 nm, with a lower light intensity, thus resulting in slower current changes due to slower photoisomerization process. Inset shows the light‐induced optical absorbance changes at 356 nm characteristic of the

trans

‐azobenzene isomer measured with the same membrane immersed in an aqueous solution. The time scale of the UV/Vis irradiation corresponds to that of the first cycle in the chronoamperometric measurements.

Figure 4.31 (A) Schematic drawing of the reversible c

hange in size of the azobenzene‐modified pores in response to the light signals. (B) Cross‐sectional TEM image of the photoresponsive nanoporous film. Inset is a 2D GISAXS pattern shown in reverse grayscale. The lower half of the plane is in the shadow of the silicon substrate, and the scattering spots are attenuated. (C) Structures of the used ferrocene derivatives.

Figure 4.32 β‐Aminocyclodextrin

(

16

) monolayer assembly and its inclusion complex formation with azobenzene‐functionalized viologen (

17a/b

). The association constant,

K

a

depends on the isomeric state of azobenzene unit, which is photochemically controlled.

Figure 4.33 (A) Cyclic voltammograms obtained with the cyclodextrin‐modified electrode in the presence of the azobenzene‐functionalized viologen

being in different isomeric states (

cis

‐ or

trans

‐). The experiments were performed in 0.01 phosphate buffer, pH 10.8; scan rate 100 mV s

−1

. Inset: Cyclic cathodic response measured at 0.6 V (vs SCE) upon reversible photoisomerization of the azobenzene unit. The higher and lower currents correspond to the

trans

‐ and

cis

‐isomeric states, respectively, produced by UV and Vis irradiation (see the scheme in Figure 4.32). (B) Time‐dependent quartz crystal microbalance (QCM) measurements performed in the presence of the

cis

‐ and

trans

‐isomers. The smaller response in the presence of the

cis

‐isomer corresponds to the smaller association of this isomer with the recognition cyclodextrin units. The inset photo shows a QCM sensing unit used for the measurements. The inset plot shows reversible changes of the QCM frequency upon photoisomerization process.

Figure 4.34 Schematic illustration showing assemblin

g of the “molecular machine” moving the redox species along the molecular wire to different distances from the electrode surface. EDC is 1‐ethyl‐3‐(3‐dimethylaminopropyl)carbodiimide and Fc is ferrocene.

Figure 4.35 (A) Schematic illustration showing different positioning of the ferrocene redox species for different isomeric states of the azobenzene units in the molecular wires. The

trans

‐isomer allows shorter distance between the ferrocene and electrode, thus providing faster electron transfer. (B) Chronoamperometri

c traces measured for

cys

‐ and

trans

‐states of the azobenzene units. Faster current decay measured chronoamperometrically corresponds to the faster electron transfer process. Inset: Cyclic changes of the electron transfer rate constant upon reversible photoisomerization of the system between the

cys

‐ and

trans

‐states.

Figure 4.36 Reversible photochemical isomerization followed by reversible protonation/deprotonation of the open‐ring isomer

: structures of the neutral spiropyran (SP), zwitterionic merocyanine (MR), and positively charged protonated merocyanine (MRH

+

).

Figure 4.37 Schematic illustration of the photochemically controlled electrochemical reduction of O

2

biocatalyzed

by cytochrome oxidase (COx) and mediated by cytochrome

c

(Cyt). The active (ON) state of the electrode corresponds to the neutral SP species and the inactive (OFF) state of the electrode corresponds to the positively charged protonated MRH

+

species on the Au electrode surface; (red) and (ox) indicate the reduced and oxidized species, respectively.

Figure 4.38 (A) Cyclic voltammograms corresponding to the redox reactions of Cyt measured w

ith the photoswitchable modified electrode shown schematically in Figure 4.37: (a) the electrode in the active (ON) state in the presence of SP form at the electrode surface obtained after 5 min of electrode illumination,

λ

 > 475 nm (Vis light); (b) the electrode in the inactive (OFF) state in the presence of MRH

+

form at the electrode surface obtained after 2 min irradiation, 360 < 

λ

< 400 nm (UV light). The solution included 0.1 mM Cyt with 0.1 M Na

2

SO

4

and 0.01 M phosphate buffer, pH 7.0; potential scan rate 50 mV s

−1

. (Note that in this experiment COx was not included in the solution). Inset: Peak current (

I

pc

) variation upon cyclic irradiation of the electrode with Vis (ON state) and UV (OFF state) light. (B) Frequency changes of a quartz crystal microbalance (QCM) modified with a mixed monolayer similar to one shown in Figure 4.37 upon cyclic irradiation with Vis and UV light. The inset photo shows a QCM sensing unit used for the measurements.

Figure 4.39 Cyclic voltammograms corresponding to the

bioelectrocatalytic reduction of O

2

measured with the modified electrode shown schematically in Figure 4.37: (a) the electrode in the active (ON) state in the presence of SP form at the electrode surface obtained after 5 min of electrode illumination,

λ

 > 475 nm (Vis light); (b) the electrode in the inactive (OFF) state in the presence of MRH

+

form at the electrode surface obtained after 2 min irradiation, 360 < 

λ

 < 400 nm (UV light). The solution included 0.1 mM Cyt, 1 μM COx, and O

2

in equilibrium with air. The background electrolyte was 0.1 M Na

2

SO

4

and 0.01 M phosphate buffer, pH 7.0; potential scan rate 2 mV s

−1

. Inset: Electrocatalytic current (

I

cat

) variation measured at

E

 = −0.1 V upon cyclic irradiation of the electrode with Vis (ON state) and UV (OFF state) light.

Figure 4.40 Schematic illustration of the photochemically controlled electrochemical oxidation of lactate (Lac) to pyruvate (Pyr) biocatalyzed by lactate dehydrogenase (LDH) and mediated by cytochrome

c

(Cyt). The active (ON) state of the electrode corresponds to the neutral SP species and the inactive (OFF) state of the electrode corresponds to the positively charged protonated MRH

+

species on the Au electrode surface; (red) and (ox) indicate the reduced and oxidized species, respectively.

Figure 4.41 Schematic illustration of the photochemically switchable

bioelectrocatalytic glucose oxidation at the electrode surface functionalized with the SP/MRH

+

photoisomerizable species. The GOx functionalized with a ferocene mediator was electrostatically attracted to the positively charged MRH

+

species resulting in higher enzyme concentration near the electrode surface, thus increasing the reaction rate.

Figure 4.42 Cyclic voltammograms obtained with the modified electrode (see Figure 4.41 for the scheme) in the presence of the ferrocene‐functionalized GOx, 1.5 mg mL

−1

: (a) with the SP electrode and in the absence of glucose, (b) with the SP electrode and glucose, 5 × 10

−2

M, and (c) with the MRH

+

electrode and glucose, 5 × 10

−2

M. Inset: Reversible variation of the electrocatalytic current measured at

E