Photobioelectrocatalysis of Intact Chloroplasts for Solar Energy

Subscriber access provided by University of Newcastle, Australia

Article

Photobioelectrocatalysis of Intact Chloroplasts for Solar Energy Conversion Kamrul Hasan, Ross D. Milton, Matteo Grattieri, Tao Wang, Megan Stephanz, and Shelley D. Minteer ACS Catal., Just Accepted Manuscript • DOI: 10.1021/acscatal.7b00039 • Publication Date (Web): 13 Feb 2017 Downloaded from http://pubs.acs.org on February 20, 2017

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a free service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are accessible to all readers and citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

ACS Catalysis is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 33

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Catalysis

1

Photobioelectrocatalysis of Intact Chloroplasts for

2

Solar Energy Conversion

3

Kamrul Hasan†, Ross D. Milton†, Matteo Grattieri†, Tao Wang†, Megan Stephanz† and Shelley

4

D. Minteer*†

5

†Departments of Chemistry and Materials Science & Engineering, University of Utah

6 7 8

315 S 1400 E Room 2020, Salt Lake City, Utah 84112, USA * Corresponding author. E-mail: [email protected]

9

Abstract: Recently, interest in photosynthetic energy conversion has substantially increased.

10

Chloroplasts, the photosynthetic organelle inside higher plants and algae, are the ultimate source

11

of carbon-based fuels. However, they are less studied in a photobioelectrochemical cell, because

12

their electrochemical communication at an electrode surface is challenging due to their complex

13

membrane system. Although redox polymers are widely used for mediating bioelectrocatalysis,

14

they have never been explored for wiring chloroplasts to electrodes. Herein, a naphthoquinone-

15

functionalized linear poly(ethylenimine) (NQ-LPEI) redox polymer is used as an electron

16

transfer (ET) mediator as well as the immobilization matrix for chloroplasts. They are

17

immobilized on Toray carbon paper electrodes (TPs) and the photo-excited ET from water

18

oxidation is evaluated showing that intact chloroplasts can undergo direct electron transfer 1 Environment ACS Paragon Plus

ACS Catalysis

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

1

(DET) and mediated electron transfer (MET). Photocurrent generation by DET of chloroplasts

2

results in an oxidative current of 1.5 ± 0.2 µA cm-2. On NQ-LPEI modified electrodes, the

3

oxidative photocurrent increased to 4.7 ± 0.7 µA cm-2 and further improved to 28.7 ± 5.7 µAcm-2

4

in the presence of an additional diffusive mediator, 2,6-dichlorobenzoquinone (DCBQ). The

5

oxidative current produced in the present of light confirms the ability to oxidize water (H2O) at a

6

chloroplast-modified electrode surface.

7

KEYWORDS: Chloroplasts : Photobioelectrocatalysis : Photocurrent : Electron-transfer : Redox

8

Polymer

9

Introduction

10

In the 21st century, one of the most important challenges for human society is the development of

11

sustainable, ecofriendly and economically feasible energy sources1. Presently the majority of

12

energy is delivered from non-renewable energy sources, such as the combustion of fossil fuels,

13

which has a detrimental effect on the ecosystem and eventually on climate change. The current

14

demand for energy is projected to more than double by 2050 compared to the present time due to

15

population growth and economic development1. In contrast, fossil fuel reserves are depleting,

16

which is pressing a demand to invent, develop and implement sustainable energy sources2. A

17

great deal of research is currently centered on the development of sustainable carbon neutral

18

energy sources1, 3. However, only 16% of global energy currently comes from renewable energy

19

resources such as solar energy, wind, rain energy, energy from waves, geothermal heat, etc.

20

Among all renewable energy resources, solar energy is by far the most accessible, reliable and

21

exploitable energy resource4. The amount of solar energy reaching the earth’s surface in 1 hour is

22

greater than the entire annual human society energy consumption5.

2 Environment ACS Paragon Plus

Page 2 of 33

Page 3 of 33

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Catalysis

1

In order to convert solar energy into chemical energy, nature has developed its own energy

2

transducing system, called photosynthesis. All higher plants, algae and some bacteria (i.e., purple

3

bacteria) perform photosynthesis. Since oxygen is released as a byproduct of their photosynthetic

4

activity, they are classified as oxygenic photosynthetic systems. However, photosynthesis in

5

purple bacteria does not generate oxygen as a byproduct and the process is termed anoxygenic

6

photosynthesis6. Currently, the only established technology available to convert sunlight directly

7

into electrical power is silicon-based photovoltaics. However, the efficiency of these systems for

8

sunlight conversion is low (less than 20%)4 and the materials used are exhaustible, highly

9

expensive, and not fully recyclable. Interestingly, the quantum efficiency of the charge

10

separation step in photosynthesis is nearly 100%7. Nevertheless, comparing the efficiency of

11

natural photosynthesis and photovoltaics is a complicated issue, since these two systems operate

12

in different ways and produce dissimilar products. Natural photosynthesis produces biomass or

13

chemical energy whereas photovoltaic devices produce electrical current7.

14

Photobioelectrochemical systems (PBESs), also termed as biological photovoltaics (BPVs)8,

15

bio-photo-electrochemical cell (BPEC)9, and bio-solar cells, are emerging as energy-generating

16

technologies where photosynthetic biomaterials are used to convert solar energy into electrical

17

energy. In PBESs a wide variety of photoactive biological components are used to convert solar

18

energy into electricity8. These biomaterials could be intact photosynthetic organisms or parts of

19

cellular or sub-cellular photosynthetic apparatus10 (i.e. photosystems (PSI11 and PSII12),

20

thylakoid membranes13, cyanobacteria14, algae15 and plants16). Each of these photosynthetic

21

biomaterials has advantageous and disadvantageous properties based on their particular

22

application. For example, purple bacteria are a less preferred candidate in PBESs, since they

23

require an additional fuel (e.g., malate), to run the system17 . Isolated photosystems are


ACS Catalysis

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

1

inherently smaller and yield improved orientation on electrode surfaces, often resulting in

2

enhanced electron transfer (ET) and eventually supporting direct-ET (DET) as a function of

3

decreased ET distances. Engineering of the ET-conduit in isolated photosystems is simpler than

4

that of an intact photosynthetic organism. The competition of electron extraction (from photo-

5

excited water oxidation) in photosystems (PSs) is lower, since there is less interference from

6

other redox enzymes around the electron donor sites. However, the stability of isolated PSs is

7

frequently limited and highly susceptible to photo-inhibition. Moreover, isolated PSs require

8

purification by protein chromatography, which is expensive and time-consuming. Thylakoid

9

membranes are considered a better candidate for use in PBESs due to their improved stability,

10

whereby photosynthetic protein complexes are kept in their native environment. Moreover,

11

thylakoid membranes contain several ET-routes and electrons from photo-excited water

12

oxidation could be transferred to the electrode via different protein complexes such as

13

plastoquinone (PQ), cytochrome b6f (cyt b6f), plastocyanin (PC), and ferredoxin (Fd) (see

14

Scheme 1). Although photosynthetic organisms are considered great candidates in PBES, they

15

require energy for their growth and maintenance. Photosynthesis in eukaryotic organisms, such

16

as in algae, is considered as the most complicated system and photo-excited energy is lost each

17

time electrons are transferred from one carrier to another17a.

18

Chloroplasts are the photosynthetic organelle employed within oxygenic photosynthetic

19

organisms, housing the site of photosynthesis - thylakoid membranes. Chloroplasts have self-

20

repair mechanisms against photo-damage and have natural biochemical pathways to scavenge

21

reactive oxygen species (ROS) produced during photosynthesis18, however, they are less studied

22

in photobioelectrochemistry compared to their counterparts for photosynthetic energy

23

conversion. Previously, chloroplasts were shown to have greater photosynthetic activity when


Page 4 of 33

Page 5 of 33

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Catalysis

1

single-walled carbon nanotubes (SWNTs) were passively transported and localized inside their

2

lipid envelope. SWNTs were added for enhanced photo-excited ET (PEET) and also to minimize

3

the production of ROS during photosynthesis19. DET has been demonstrated by the insertion of

4

ultra-sharp nano-electrodes into the chloroplast of an algal cell (Chlamydomonas reinhardtii)20.

5

A photocurrent of 10.2 µA cm-2 was reported with chloroplasts adsorbed on a nano-crystalline

6

TiO2 film on an indium tin oxide (ITO)-coated glass electrode21.

7

The photo-electrochemical communication of chloroplasts in a PBES is presumably difficult,

8

since they are shielded by complex lipid-membrane systems (thickness ≈6-8 nm)22. Outer and

9

inner membranes of chloroplasts are separated by an intermembrane space that has a thickness of

10

10-20 nm23, thus, PEET from the chloroplasts has long been thought to be difficult. To overcome

11

this limitation, exogenous ET-mediators are frequently used to establish their electrochemical

12

communication

13

dichlorophenolindophenol (DCIP)25. The use of these exogenous mediators is not favorable in

14

PBESs due to concerns surrounding their sustainability in a practical application.

on

electrode

surfaces,

such

as

methyl

viologen24

and

2,6-

15

Instead, we hypothesize that surface confined polymeric redox mediators are a better choice

16

for ET. Polymeric mediators provide an immobilization matrix for biocatalysts at electrode

17

surfaces as well as mediate their ET. They do not diffuse into the bulk solution and provide

18

three-dimensional (3D) architectures of biocatalysts on the electrode that results in higher

19

catalytic responses26. Redox polymers for MET of chloroplasts have not yet reported in the

20

literature. Here, we report on the photobioelectrochemical wiring of chloroplasts on Toray

21

carbon paper-based electrodes modified with a naphthoquinone functionalized linear

22

poly(ethylenimine) (NQ-LPEI) redox hydrogel. To confirm the source of photocurrent, diuron is

23

used as a photosynthetic inhibitor.


ACS Catalysis

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

1 2 3 4

Experimental

5

Chemicals: 3-(N-morpholino)propanesulfunic acid (MOPS), magnesium chloride (MgCl2), 3-

6

(3,4-dichlorophenyl)1-1,dimethylurea (diuron), 2,6-dichlorobenzoquione (DCBQ), bovine serum

7

albumin (BSA), sorbitol, 4-(2-hydroxyethyl)piperazine-1-ethanesulfunic acid (HEPES), sodium

8

chloride (NaCl), ethylenediaminetetra acetic acid (EDTA), acetone, ethylene glycol diglycidyl

9

ether (EGDGE) and Percoll®

were purchased from Sigma-Aldrich (St. Louis, MO). All

10

chemicals were either analytical or research grade. Aqueous solutions were prepared by using

11

Milli-Q deionized water (18 MΩ cm-1). The synthesis and preparation of NQ-LPEI with a

12

midpoint potential of E1/2 = -0.34 vs. SCE was prepared as reported earlier27. The NQ-LPEI was

13

dissolved in Milli-Q water at concentration of 10 mg mL-1.

14

Isolation of intact chloroplast: Intact chloroplasts from spinach (Spinacia oleracea) were

15

isolated using the chloroplast isolation kit by Sigma-Aldrich (St. Louis, MO). Spinach was

16

purchased from a local grocery store (Salt Lake City, Utah). The entire isolation process was

17

performed under refrigeration (2-4 °C). Briefly, 30 g of spinach leaves were thoroughly washed

18

with Milli-Q water and then transferred to a blender with 120 mL of 1 M chloroplast isolation

19

buffer (CIB) at pH 7.8 containing 0.1% of (w/v) BSA. The constituents of 1 M CIB per 1 L is as

20

follows: 0.33 M sorbitol (60 g), 0.05 M HEPES (11.92 g), 10 mM NaCl (0.58 g), 2.6 mM EDTA

21

(0.74 g) and 10.6 mM MgCl2 (1 g). Afterwards leaves were processed with 2-4 blender strokes

22

within 5 seconds and the macerate was gradually passed through three-layers of cheesecloth. The


Page 6 of 33

Page 7 of 33

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Catalysis

1

filtrate was evenly distributed into four 50 mL tubes and centrifuged at 200 ×g for 3 minutes to

2

remove all unwanted whole cell and cell-wall debris.

3

Supernatants from each tube were transferred to fresh, chilled 50 mL tubes and centrifuged

4

again at 1000 ×g for 7 minutes and eventually chloroplasts appeared as a green pellet on the

5

bottom. Supernatants were then discarded and the pellet was dislodged by gently tapping the

6

centrifuge tube. The pellet from each tube was re-suspended with 1-2 mL of CIB buffer

7

containing 0.1% of BSA. At this stage, intact chloroplasts were separated from broken

8

chloroplasts by centrifuging at 1700×g for 6 minutes on a Percoll® (40%) layer. 10 mL of 40%

9

Percoll® solution was used for each 6 mL of chloroplast suspension. After the centrifugation,

10

intact chloroplasts were carefully collected from the bottom of the tube and re-suspended in 0.5

11

mL of CIB without BSA.

12

Measurement of chlorophyll concentration: The concentration of chlorophyll inside

13

chloroplasts was estimated according to the procedure of Arnon et al28. Briefly, 10 µL of the

14

freshly-isolated chloroplasts was suspended in 1 mL of 80% acetone and centrifuged at 3000 ×g

15

for 2 minutes. The supernatant was collected and its absorbance was measured at 652 nm. The

16

chlorophyll concentration in the isolated chloroplasts were set at 1 mg mL-1 and stored at -80 °C.

17

All steps were performed under refrigeration (2-4 °C).

18

Working electrode preparation: Toray® carbon-paper electrodes (TPs) (Fuel Cell Earth,

19

Woburn, MA) (1 cm2 geometric surface area) were used as working electrodes. For DET, an

20

aliquot of 30 µL of chloroplasts solution (chlorophyll concentration 1 mg/mL) was spread onto

21

the active surface area of the electrode and allowed to dry at room temperature for ≈20 minutes.

22

For NQ-LPEI modified electrodes, NQ-LPEI (10 mg/mL), chloroplasts (1 mg mL-1), and

23

EGDGE (10% v/v) were mixed together with an amount of 21 µL, 9 µL and 1.13 µL and then 30


ACS Catalysis

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 8 of 33

1

µL of mixed solution was spread onto the electrode and allowed to dry at room temperature for

2

≈20 minutes. of EGDGE was used as a cross-linker between the polymer backbone (LPEI) and

3

the chloroplasts. The ratio of polymer/cross-linker/chloroplast was optimized according to a

4

previously published report44.

5

Confocal images of chloroplasts were obtained with an Olympus FV100 confocal microscope

6

with PLANAPO 60X immersion. To yield ruptured chloroplasts, the chloroplasts were sonicated

7

for 5 minutes at 30% amplitude for 20-second pulse on and 1-second pulse off (Fisher

8

ScientificTM model 505 sonic dismembrator, Pittsburgh, PA, USA) prior to immobilization on

9

the electrode surface.

10

Electrochemical

measurements

and

instruments:

Cyclic

voltammetry

(CV)

and

11

chronoamperometry (CA) were used for electrochemical characterization. All experiments were

12

performed with a VSP Biologic potentiostat controlled by EC-LAB software (Paris, France). A

13

three-electrode setup including a saturated calomel electrode (SCE) as a reference electrode, bare

14

TPs or NQ-LPEI modified TPs as the working electrode and platinum foil as a counter electrode

15

was used. All representative voltammograms presented here were taken from the fourth scan,

16

when system was stable. All potentials in this manuscript are reported vs. SCE. All of the

17

experiments were performed at room temperature (20 ± 2 °C) and all data presented here are

18

based on three independent experimental replicates where the reported uncertainties correspond

19

to one standard deviation. All voltammograms and chronoamperometric traces were presented

20

following the polarographic notation, whereby negative current values represent oxidation

21

reactions.

22

A Dolan-Jenner Fibre-Lite lamp (model 190-1 quartz-halogen illumination system with a

23

optical light guide providing a light intensity of 76 mW cm-2 (a light intensity higher than the


Page 9 of 33

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Catalysis

1

saturated light intensity, i.e., 30 mW cm-2 for photosynthesis)) was used to excite the

2

photosynthetic activity of chloroplasts on TPs. In all experiments, 100 mM MOPS buffer at pH

3

7.0 including 10 mM MgCl2 (MOPS buffer) was used as an electrolyte. In CV experiments, the

4

scan rate was 5 mV s-1 unless stated otherwise. In all CA experiments, the applied potential

5

(Eappl) was +0.3 V vs. SCE, a potential higher than the midpoint potential (E1/2) of any used

6

redox mediator to confirm the generation of potential independent photocurrent. All experiments

7

were conducted under anoxic conditions, achieved by bubbling N2 gas into the electrolyte for

8

≈20 minutes immediately prior to electrochemical analysis.

9 10 11 12 13 14 15 16 17 18 19

Scheme 1. The schematic of photo-excited ET (mediated by chloroplasts) from water oxidation to the electrode. The thylakoid membrane comprises three major photosynthetic protein complexes: PSII, PSI and cytochrome (Cyt) b6f. Upon light absorption, PSII excites the energy level of an electron to a higher energetic state (P680*) and is subsequently relaxed by water oxidation mediated by the oxygen-evolving complex (OEC). The photo-excited electron then passes through a series of electron carriers, such as pheophytin (Phe), quinone-A (QA) and quinone-B (QB) and successively reduces PQ to plastoquinol (PQH2). PQH2 is a strong electron donor and passes electrons to Cyt b6f followed by the PC to the PSI. Upon excitation by light absorption, PSI then transfers electrons to Fd that eventually reduces nicotinamide adenosine dinucleotide phosphate (NADP+) to NADPH via ferredoxin-NADH-reductase (FNR). This ET


ACS Catalysis

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

1 2 3 4 5 6 7 8 9

Page 10 of 33

from photo-excited water oxidation to the production of NADPH is coupled with a proton gradient across the thylakoid membrane that is used to generate ATP (not shown in the scheme). Detailed components of PETC are not shown for simplicity. Black dotted arrows indicate internal-ET. The red arrow identifies the location where diuron inhibits PSII. Purple circles represent the redox center (NQ) of the polymer linked with the backbone (LPEI). Green arrows indicate PEET. All symbols signify the typical sign in photosynthesis and are explained in the text.

Results and Discussion

10

In oxygenic photosynthetic organisms, photosynthesis occurs in thylakoid membranes located

11

within the chloroplast. Electrons from photosynthetic-ET-chain (PETC) of chloroplasts could

12

transfer to the electrode either by DET or MET, as shown in Scheme 1. The terminology of DET

13

and MET is heavily debated among our field. In this specific case, we consider the use of DET to

14

be accurate for experiments conducted without the addition of any exogenous artificial electron

15

transfer mediator. When we have added external diffusional or polymeric mediators to the

16

system, we define it as MET. DET may take place via membrane proteins in the chloroplast

17

outer membrane (left portion of Scheme 1). A lipid soluble ET-mediator (DCBQ) can diffuse

18

through the outer membranes (OM) and inner membranes (IM) of chloroplasts and reduced by

19

the PQ-pool as well as quinones ubiquitously present inside the membranes29. Lipid-insoluble

20

ET-mediators (i.e. NQ-LPEI) can pass through the OM via non-specific porin and can be

21

reduced at the IM either by the un-identified transmembrane protein (TMP) or NADPH. These

22

three possible mechanisms were evaluated electrochemically in this paper to understand the

23

efficiency of each mode of electron transport. However, we first characterized the chloroplast

24

before electrochemical measurements.

25

To evaluate the photosynthetic pigment of the chloroplasts, the UV-visible absorption spectra

26

of the chloroplasts suspended in MOPS buffer was recorded with a spectrophotometer (Figure

27

S1, Supporting information). The primary photosynthetic pigments30, chlorophyll, appeared at


Page 11 of 33

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Catalysis

1

their typical absorption maxima at 682 nm. The secondary pigments, carotenoids, appeared at

2

their regular absorption maxima at 482 and 440 nm. These data are consistent with literature

3

reports for chloroplasts21. The intactness of chloroplasts is important, since their photo-

4

electrochemical function and metabolic activity depends on the structural integrity. To

5

investigate if the internal components of chloroplasts are undamaged during the isolation

6

process, they were imaged using confocal fluorescence microscopy. An aliquot of 200 µL of

7

intact and sonicated chloroplasts solution was investigated as shown in Figure 1. Intact

8

chloroplasts contain thylakoid membranes, stromal compartments and the envelope

9

surroundings. Under microscopy, it is shown that an intact chloroplast gives an opaque, shiny

10

and halo appearance whereas a sonicated chloroplast (deliberately ruptured) gives a granulated

11

and flat appearance that is in close agreement with earlier literature studies31. To calculate the

12

number of intact chloroplasts, images (Figure 1) were threshold-normalized to identify the red

13

fluorescent regions. The areas were encoded to separate objects and then the particles were

14

analyzed with a 101-10000 pixel cutoff. The average size of the intact chloroplasts using this

15

criterion was 1969 pixels and debris was under 100. Comparing the debris/intact area of the

16

chloroplasts sample, more than 90% of material was found to be in intact structures. Therefore,

17

we utilized these isolated, intact chloroplasts for the voltammetric and amperometric studies

18

below.

19


ACS Catalysis

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

1

Figure 1. Confocal microscopy images of intact (A) and sonicated chloroplasts (B) in solution.

2

Images were taken under an Olympus FV1000 confocal with PLANAPO 60X oil immersion

3

objective.

Page 12 of 33

4

In bioelectrochemical systems (BESs), DET from biomaterials to the electrode surface is

5

preferable, since it simplifies the design and function of an electrochemical cell, minimizes the

6

overpotential loss, and is expected to result in high columbic efficiency32. We investigated

7

whether chloroplasts (which have a thick insulating membrane system) could communicate on

8

bare TPs without the addition of an external mediator. 30 µL of chloroplasts were drop-coated on

9

Toray® carbon paper-based electrodes and allowed to dry for ≈20 min at room temperature.

10

Chloroplast-modified electrodes were excited with a fiber optic light at an intensity of 76 mW

11

cm-2 and MOPS buffer was used as an electron donor. As shown in the voltammograms in Figure

12

2, two pairs of redox peak appear under ‘light off’ conditions and their mid point potentials (E1/2)

13

were calculated to be -0.34 V and -0.19 V. The peaks at -0.34 V and -0.19 V are attributed to QB

14

and Cyt b6f, respectively. Under ‘light on’ conditions, catalytic photo-excited electrons from the

15

oxidation of water are transferred to the electrode at an onset potential of -0.32 V. From a

16

previous study, the formal potential at neutral pH (E0´) for QB and Cyt b6f are estimated to be -

17

0.34 V and -0.16 V33. QB is known to be a mobile ET carrier inside the PETC of PSII34. These

18

data suggest that QB and Cyt b6f are responsible for PEET for chloroplasts. Previously, direct

19

PEET from chloroplasts35 were reported, however, the responsible redox proteins were

20

unidentified21. In control CV experiments (Figure S2, Supporting Information), Toray carbon

21

paper electrodes were illuminated without chloroplasts and are not able to generate any

22

photocurrent.


Page 13 of 33

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Catalysis

1 2

Figure 2. Representative cyclic voltammograms of chloroplast-modified TP electrodes under

3

‘light off’ (black line) and ‘light on’ (red line) conditions. Electrolyte: 100 mM MOPS buffer at

4

pH 7.0 including 10 mM MgCl2. Light intensity: 76 mW cm-2. Scan rate: 5 mV s-1.

5

Chronoamperometric experiments were performed to obtain a quantitative photocurrent

6

generation by applying a potential of 0.3 V vs. SCE (Figure 3), ensuring a large enough

7

overpotential for H2O photo-oxidation. Chloroplast modified-TP electrodes were kept under

8

‘light off’ conditions for 900 s to obtain a stable baseline. The electrodes were then exposed to

9

light for 100 s and three identical runs were recorded. To determine the net photocurrent, the

10

response generated under ‘light off’ conditions was subtracted from the responses obtained under

11

illumination. Illumination does not have any noticeable influence on the current of Toray®

12

carbon paper electrodes in the absence of chloroplasts (Figure S2, Supporting Information),

13

while photocurrent generated on chloroplast-modified electrodes was calculated to be 1.57 ±

14

0.28 µA cm-2 on the first illumination cycle (Figure 3). The photocurrent generation gradually

15

decayed on the second and third cycles to 1.5 ± 0.2 µA cm-2 and 1.4 ± 0.2 µA cm-2, respectively.

16

This gradual decay of photocurrent is most likely attributed to photoinhibition36, a process where


ACS Catalysis

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 14 of 33

1

there is an imbalance between the energy produced in the light dependent reaction of

2

photosynthesis and the energy utilized in the light independent reaction. When PSII is exposed to

3

high light intensities (>> saturated light intensity for photosynthesis ≈ 30 mW cm-2), ROS are

4

generated inside the PETC due to the electron leakage that is consequence of cellular photo-

5

oxidative damage37 .

6 7

Figure 3. Amperometric i-t curves of Toray® carbon electrodes in the absence (black line) and

8

presence of chloroplasts (red line). Electrolyte: 100 mM MOPS buffer at pH 7.0 including 10

9

mM MgCl2. Light intensity: 76 mW cm-2. Applied potential (Eappl) = +0.3 V vs. SCE. First,

10

second and third cycles are shown from left to the right.

11 12

It is critical to evaluate the effect of different light intensities on chloroplast modified-TP

13

electrodes, as shown in Figure 4. The photocurrent of 1.11 ± 0.19 µA cm-2 was obtained at a

14

light intensity of 27 mW cm-2. When the light intensity was increased to 52 mW cm-2 (nearly

15

double), the photocurrent did not increase significantly (1.19 ± 0.16 µA cm-2). This is likely due

16

to the fact that the photosynthetic activity of the chloroplasts is already saturated at 27 mW cm-2

17

and that the greater light intensities result in photoinhibition. When the light intensity was set at


Page 15 of 33

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Catalysis

1

76 mW cm-2, the photocurrent obtained was 1.24 ± 0.17 µA cm-2. It is important to note that the

2

light intensity of 76 mW cm-2 previously yielded the greatest responses (Figure 3), thus, this

3

intensity was used for all of the following studies.

4 5

Figure 4. Effect of different light intensities on photocurrent generated on chloroplasts modified-

6

TP electrodes. Electrolyte: 100 mM MOPS buffer at pH 7.0 including 10 mM MgCl2. Light

7

intensity: 76 mW cm-2. Applied potential (Eappl) = +0.3 V vs. SCE. In control experiments,

8

different light intensities do not show any detectable response on TP electrodes without

9

chloroplasts (data not shown).

10

Although DET is a preferable choice in BESs, electrical current and power generated from a

11

DET-based system is usually lower than that of an MET-based system38, whereby

12

bioelectroatalytic currents generated by sluggish DET can be improved. Instead polymeric

13

mediators are preferred for MET due to their efficient electron shuttling properties, stable

14

adsorption on the electrode surfaces (they do not diffuse into the bulk solution) and formation of

15

an immobilized matrix of biocatalyst on the electrode surface26.

16

mediators have been explored in BESs such as osmium39, ferrocene40, ruthenium41 and

17

viologen42-based redox polymers. Naphthoquinone-based (NQ) redox polymers, having a


A variety of polymeric

ACS Catalysis

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 16 of 33

1

relatively low formal potential (E0´), were demonstrated as an alternative polymeric ET-mediator

2

for high current density, enzymatic bioelectrocatalysis of glucose27. Currently the use of NQ-

3

based polymers for glucose-oxidizing enzymes is rapidly increasing, since their redox potential

4

could be tuned to support the local ET environment43. However, the covalent immobilization

5

chloroplasts alongside a redox polymer has not yet been demonstrated. To improve the PEET

6

mediated by chloroplasts, we have investigated NQ-functionalized linear polyethyleneimine

7

(NQ-LPEI). The step-by-step synthesis of NQ-LPEI has been recently reported and the chemical

8

structure is shown in Figure 527, 44.

9

Initially the photosensitivity of NQ-LPEI was studied. 30 µL of NQ-LPEI was drop-coated on

10

the TP electrodes and allowed to dry for ≈ 20 minutes at room temperature. CVs of the polymer-

11

modified electrodes were performed under ‘light off’ and ‘light on’ conditions (Figure S3,

12

Supporting Information). In the absence of light, a pair of redox peak appear at (E1/2 vs. SCE)

13

-0.34 V, which are in close agreement with the theoretical formal redox potential reported earlier

14

for this polymer27. Since the polymer does not show any oxidative photosensitivity under the

15

‘light on’ condition, the mediated oxidative photoelectrochemistry of chloroplasts were

16

subsequently studied with this polymer.

17 18

Figure 5. Chemical structure of NQ-LPEI used in this work.


Page 17 of 33

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Catalysis

1

In the presence of chloroplasts immobilized within the NQ-LPEI redox polymer (on TPs,

2

Figure 6) under ‘light off’ conditions, a typical set of redox peak appears with an E1/2 at -0.34 V.

3

Under illumination, an overall oxidative wave is observed, suggesting that the NQ-LPEI redox

4

polymer can communicate with the chloroplasts through their PETC and catalyze the PEET.

5

Electrons from water oxidation by chloroplasts are eventually transferred to the electrode via the

6

self-exchange characteristics of NQ-LPEI where electrons and protons are transferred between

7

the chloroplast and the electrode through the NQ groups on the functionalized backbone

8

(LPEI)26. Previously, osmium-based redox hydrogels were shown for photo-electrochemical

9

communication of algae via a similar self-exchange-based mechanism15a. Polymeric-mediators

10

were assumed to access the cytosolic membranes of bacterial cells and extract electrons from

11

TMPs, however, the details mechanism is not yet known45.

12 13

Figure 6. Representative cyclic voltammograms of chloroplasts immobilized on NQ-LPEI

14

modified TP electrodes under ‘light off’ (black line) and ‘light on’ (red line) conditions.

15

Electrolyte: 100 mM MOPS buffer at pH 7.0 including 10 mM MgCl2. Light intensity: 76 mW

16

cm-2. Scan rate: 5 mV s-1.


ACS Catalysis

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

1

Following

the

voltammetric

analysis

of

photocurrent

Page 18 of 33

generation

(Figure

6),

2

chronoamperometry were performed on electrodes prepared in an identical fashion (Figure 7) to

3

evaluate their capacitance-free photobioelectrochemical current generation. For all amperometric

4

experiments, an applied potential of +0.3 V was used. The Eappl was more positive than the

5

midpoint potential of the polymer (E1/2 = -0.34 vs. SCE) to ensure significant PEET at the

6

electrodes. Upon illumination of the intact chloroplasts, a photocurrent of 5.70 ± 0.30 µA cm-2

7

was generated on the first cycle that is five times higher than that of the response recorded when

8

chloroplasts were immobilized on TP electrodes without the polymer modification (Figure 3).

9

The photocurrent responses on the second and third cycles are 4.90 ± 0.2 µA cm-2 and 4.3 ± 0.1

10

µA cm-2, respectively. The gradual decreases of photocurrent are attributed to photoinhibition36.

11

The low catalytic activity of NQ-LPEI might be due to its lower formal potential (E1/2 vs. SCE)

12

and limited accessibility through the photosynthetic electron transfer chain of chloroplasts.

13

However, the photocurrent generated in the presence of this polymer (5.7 ± 0.3 µA cm-2, first

14

cycle) is statistically greater compared to a similar condition in the absence of polymer in Figure

15

3 (1.6 ± 0.3 µA cm-2). A series of control experiments were performed to confirm that the

16

photocurrent is produced from intact chloroplasts, as shown in Figure 7. Under illumination,

17

neither the NQ-LPEI redox polymer nor the sonicated chloroplasts on NQ-LPEI modified-TP

18

electrodes are able to generate a significant photocurrent. To deliberately destroy the

19

photosynthetic entities of chloroplasts, they were sonicated for 5 minutes. To confirm that a

20

protein with photosynthetic features is required to generate photocurrent, we investigated bovine

21

serum albumin (BSA, 10 mg/mL) on NQ-LPEI modified-TP electrodes and could not observe

22

any visible response. In control experiments, it is shown that a redox center inside the polymeric-


Page 19 of 33

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Catalysis

1

ET mediators is required for photocurrent generation (Figure S4 and S5, Supporting

2

Information).

3 4

5 6

Figure 7. Amperometric i-t curves of NQ-LPEI modified-TP electrodes at different

7

modification. A) NQ-LPEI without chloroplast; B) NQ-LPEI with sonicated chloroplasts; C)

8

NQ-LPEI with BSA (10 mg/mL); D) NQ-LPEI with intact chloroplasts. Electrolyte: 100 mM

9

MOPS buffer at pH 7.0 including 10 mM MgCl2. Light intensity: 76 mW cm-2. Applied potential

10

(Eappl) = +0.3 V vs. SCE.

11

These data suggest that the photocurrents reported here are generated from intact chloroplasts

12

comprising three distinct entities I) thylakoid membranes that contain pigments and PETC and

13

coupling factors for photophosphorylation; II) stroma that contains necessary soluble proteins

14

and metabolites for light independent reaction in photosynthesis; and III) envelope membranes

15

that enclose the organelle and also contain the metabolite translocators46. Control experiments

16

were performed to show the intactness of chloroplasts on TP electrodes (Figure S6, Supporting

17

Information). Although chloroplasts in NQ-LPEI modified-TP electrodes exhibited five times


ACS Catalysis

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 20 of 33

1

higher photocurrent compared to when chloroplasts were immobilized on directly on TP

2

electrodes, it was interesting to measure their open circuit potential (OCP). The term OCP refers

3

to the potential difference between the working electrode and the reference electrode when no

4

potential or current is applied to the cell. The OCP was measured in a PBES half-cell using a

5

chloroplast anode. Under illumination, chloroplast-modified TP electrodes exhibited a higher

6

potential shift (from +34 mV to -251 mV) when compared to chloroplasts that were immobilized

7

within the NQ-LPEI redox polymer (from -14 mV to -201 mV, Figure 8). A potential loss with

8

the MET-based system is typical. When the systems were kept under ‘light off’ conditions, the

9

OCP of both electrodes closely resembled their initial values. The OCP changes of both

10

electrodes are attributed to the photo-electrochemical reactions of chloroplast on the electrodes.

11

However, the photo-excited-ET mediated by the NQ-LPEI redox polymer system is five times

12

higher than that on TP electrodes (comparing Figure 3 and Figure 7). These data indicated that

13

NQ-LPEI facilitated ET from the chloroplasts to the electrode surface is more efficient.

14

Previously a genetically engineered cyanobacterium was reported for higher OCP compared to

15

its wild counterpart47.

16


Page 21 of 33

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Catalysis

1

Figure 8. OCP of chloroplast modified TP electrodes and on NQ-LPEI-TP electrodes at ‘light

2

off’ and ‘light on’ condition. Electrolyte: 100 mM MOPS buffer at pH 7.0 including 10 mM

3

MgCl2. Light intensity: 76 mW cm-2. Reference electrode: SCE.

4

To enhance the photo-dissociated-ET, we investigated two different ET-mediators together.

5

NQ-LPEI was used as an immobilized redox polymer and DCBQ was used as a soluble

6

exogenous mediator. Recently, DCBQ was reported as a superior quinone-based mediator for

7

photosynthetic electron extraction from a green alga48. Chloroplast-modified NQ-LPEI-TP

8

electrodes were prepared and 50 µM of DCBQ was dissolved in the electrolyte. Under ‘light off’

9

conditions, a pair of redox peaks with a E1/2 of -0.34 V appears for NQ-LPEI and another pair of

10

redox peaks appears at E1/2 = +0.07 V for DCBQ (Figure 9). These observed E1/2 values are in

11

close resemblance with their theoretical redox potential (E0´) reported earlier (E1/2)44, 48b. Under

12

‘light on’ conditions, the anodic currents increase significantly and the cathodic currents

13

concomitantly decrease for both of these ET-mediators. The oxidative current associated with

14

DCBQ-mediated ET is much higher than that obtained from the use of only the NQ-LPEI redox

15

polymer. It was expected since the accessibility of a non-lipid soluble ET-mediator to the PETC

16

of chloroplast is lower than that of a lipid soluble mediator (Scheme 1).

17


ACS Catalysis

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 22 of 33

1

Figure 9. Representative cyclic voltammograms of chloroplasts immobilized on NQ-LPEI

2

modified TP electrodes in the presence of 50 µM DCBQ at ‘light off’ (black line) and ‘light on’

3

(red line) conditions. Electrolyte: 100 mM MOPS buffer at pH 7.0 including 10 mM MgCl2.

4

Light intensity: 76 mW cm-2. Scan rate: 5 mV s-1.

5

Quinone molecules are known as PSII electron acceptors and compete with PQ molecules for

6

electrons generated from photo-excited water oxidation. PQs function as a mobile ET-carrier and

7

functionally link PSII with PSI. DCBQ could displace the rate-limiting step in photosynthetic-

8

ETC, QA QB, and thus efficiently oxidize PQ-pool34. The addition of DCBQ into electrolyte

9

was reported for enhanced the oxidation of QA29 and eventually improved photocurrent

10

generation.

11

To quantity photocurrent generation at the double-mediator system, amperometry was

12

performed (Figure 10) in an identical experimental setup to that used in Figure 9. Under ‘light

13

on’ conditions, NQ-LPEI modified TP electrodes in the absence of chloroplasts could not

14

generate any noticeable photocurrent. The immobilization of the chloroplasts on NQ-LPEI

15

modified-TP electrodes yields a maximum photocurrent of 29 ± 6 µA cm-2 (the value was

16

calculated as average from three cycles). This generated photocurrent is five times greater than

17

those obtained with NQ-LPEI-TP electrodes in the absence of DCBQ (Figure 7). To the best of

18

our knowledge, this is the largest photocurrent obtained with isolated chloroplasts in a PBES. It

19

is likely that DCBQ has improved accessibility through the photosynthetic housing of

20

chloroplasts due to their lipophilic structure and they are not chemically-anchored to a polymeric

21

backbone. An additive effect between the two mediators may also be present, where DCBQ

22

could act as an intermediary electron mediator to the NQ-LPEI redox polymer immobilized on

23

the electrode surface. A double mediator system was studied with cyanobacteria49 as well as in


Page 23 of 33

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Catalysis

1

yeast cell50 for greater electrochemical communication. Enhanced photocurrent generation in the

2

presence of double mediator was observed also in the present results, as reported in figure 11.

3

Three independent experiments were performed for each MET system and their standard

4

deviations are shown. The causes of different amount of photocurrent generation at different

5

mediating systems were explained in the respective sections.

6 7

Figure 10. Amperometric i-t curves of NQ-LPEI modified TP electrodes in the absence (black

8

line) and presence of chloroplasts (red line) in addition with 50 µM DCBQ. Electrolyte: 100 mM

9

MOPS buffer at pH 7.0 including 10 mM MgCl2. Light intensity: 76 mW cm-2. Applied potential

10

(Eappl) = +0.3 V vs. SCE.

11


ACS Catalysis

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 24 of 33

1

Figure 11. Comparison of photocurrent generation mediated by chloroplasts at different

2

mediating systems. Amperometric current of TP (black), NQ-LPEI modified TP (red) and NQ-

3

LPEI modified TP electrodes in the presence of 50 µM of DCBQ (green) dissolved in the

4

electrolyte: 100 mM MOPS buffer at pH 7.0 including 10 mM MgCl2. Light intensity: 76 mW

5

cm-2,.Applied potential (Eappl) = +0.3 V vs. SCE.

6

In order to investigate the source of photocurrent, we used the most commonly studied

7

photosynthetic inhibitor (diuron). Diuron is a PSII inhibitor that irreversibly blocks ET between

8

QA and QB and consequently prevents electron flow to the PQ-pool (Scheme 1)51. Chloroplasts

9

were immobilized on NQ-LPEI modified TP electrodes in the presence of 50 µM of DCBQ

10

(Figure 12). A stable baseline was obtained by keeping the electrode under ‘light off’ conditions

11

for 900 S and subsequent illumination for 200 s results in a photocurrent response of 28.7 ± 5.7

12

µA cm-2. At this point, 0.2 mM of diuron was added to the electrolyte (at 1100 s) that drastically

13

quenched more than 90% of the initial photocurrent, reaching 2.3 ± 0.2 µA cm-2. At 1300 s, the

14

light was turned off and the photocurrent returned to the baseline, however, subsequently

15

illumination resulted in a photocurrent that could barely reach less than 10% of the initial

16

response.

17

photobioelectrochemical activity of chloroplasts by diuron. Diuron was previously reported for

18

photosynthetic inhibition on spinach thylakoid membranes52.

This

data

clearly

shows

the

irreversible


photosynthetic

inhibition

of

Page 25 of 33

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Catalysis

1 2

Figure 12. The effect of diuron on the photocurrent generation from chloroplasts immobilized on

3

NQ-LPEI modified TP electrodes in the presence of 50 µM DCBQ. Diuron: 0.2 mM, Electrolyte:

4

100 mM MOPS buffer at pH 7.0 including 10 mM MgCl2. Light intensity: 76 mW cm-2. Applied

5

potential (Eappl) = +0.3 V vs. SCE.

6

Conclusion

7

In conclusion, this is the first time that the photobioelectrochemical wiring of isolated

8

chloroplasts (from spinach) by a redox polymer (NQ-LPEI) has been reported. The photocurrent

9

obtained via DET from chloroplasts was 1.5 ± 0.2 µA cm-2, which improved to 4.7 ± 0.7 µA cm-2

10

on NQ-LPEI modified Toray carbon electrodes and further improved to 28.7 ± 5.7 µA cm-2

11

following the addition of a diffusible exogenous mediator (50 µM DCBQ). The origin of the

12

photocurrent was determined (by the use of diuron) to arise from photo-excited water oxidation.

13

Although the photobioelectrochemical stability of isolated chloroplasts is still a challenge, it will

14

be an area of exploration for future work, as well as the incorporation of carbon nanomaterials to

15

increase current density.

16 17

ASSOCIATED CONTENT


ACS Catalysis

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

1 2 3 4

Page 26 of 33

Supporting Information Available: UV-visible spectrum, control experiments with Toray electrode (with and without NQ-LPEI modification) in the absence of chloroplast, confocal microscopy images, amperometric curves for system stability. This material is available free of charge via the Internet at http://pubs.acs.org.

5 6

ACKNOWLEDGMENT

7

Financial support from Fulcrum Biosciences and the Army Research Office are greatly

8

acknowledged.

9 10

Notes

11

The authors declare no competing financial interest.


Page 27 of 33

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

1 2 3 4 5 6 7

ACS Catalysis

References.

(1) Ort, D. R.; Merchant, S. S.; Alric, J.; Barkan, A.; Blankenship, R. E.; Bock, R.; Croce, R.; Hanson, M. R.; Hibberd, J. M.; Long, S. P., Proc. Natl. Acad. Sci. 2015, 28, 8529-8536. (2) New, M.; Liverman, D.; Schroder, H.; Anderson, K., Philos. Trans. R. Soc. A. 2011, 369, 619.

8

(3)(a) Ravi, S. K.; Tan, S. C., Energ. Environ. Sci. 2015, 8, 2551-2573; (b) Kalyanasundaram,

9

K.; Graetzel, M., Curr. Opin. Biotechnol. 2010, 21, 298-310; (c) Hambourger, M.; Moore, G. F.;

10

Kramer, D. M.; Gust, D.; Moore, A. L.; Moore, T. A., Chem. Soc. Rev. 2009, 38, 25-35; (d)

11

Rosenbaum, M.; He, Z.; Angenent, L. T., Curr. Opin Biotechnol. 2010, 21, 259-264; (e) Kruse,

12

O.; Rupprecht, J.; Mussgnug, J. H.; Dismukes, G. C.; Hankamer, B., Photochem. Photobiol. Sci.

13

2005, 4, 957-970.

14

(4) Voloshin, R. A.; Rodionova, M. V.; Zharmukhamedov, S. K.; Hou, H. J.; Shen, J.-

15

R.Allakhverdiev, S. I., In Applied Photosynthesis - New Progress, Najafpour, M. M., Ed. InTech:

16

Rijeka, Croatia, 2016; p 161.

17

(5) Lewis, N. S.; Nocera, D. G., Proc. Natl. Acad. Sci. U.S.A. 2006, 103, 15729-15735.

18

(6) Sekar, N.; Ramasamy, R. P., J. Photochem. Photobiol. C., 2015, 22, 19-33.

19

(7) Blankenship, R. E.; Tiede, D. M.; Barber, J.; Brudvig, G. W.; Fleming, G.; Ghirardi, M.;

20

Gunner, M. R.; Junge, W.; Kramer, D. M.; Melis, A.; Moore, T. A.; Moser, C. C.; Nocera, D. G.;

21

Nozik, A. J.; Ort, D. R.; Parson, W. W.; Prince, R. C.; Sayre, R. T., Science 2011, 332, 805-809.

22 23

(8) McCormick, A. J.; Bombelli, P.; Bradley, R. W.; Thorne, R.; Wenzel, T.; Howe, C. J., Energ. Environ. Sci. 2015, 8, 1092-1109. 27 Environment ACS Paragon Plus

ACS Catalysis

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

1 2 3 4

(9) Pinhassi, R. I.; Kallmann, D.; Saper, G.; Dotan, H.; Linkov, A.; Kay, A.; Liveanu, V.; Schuster, G.; Adir, N.; Rothschild, A., Nat. Commun. 2016, 7, 12552. (10) Badura, A.; Kothe, T.; Schuhmann, W.; Rögner, M., Energ. Environ. Sci. 2011, 4, 32633274.

5

(11) (a) Yehezkeli, O.; Wilner, O. I.; Tel-Vered, R.; Roizman-Sade, D.; Nechushtai, R.;

6

Willner, I., J. Phys. Chem. B 2010, 114, 14383-14388; (b) Mershin, A.; Matsumoto, K.; Kaiser,

7

L.; Yu, D.; Vaughn, M.; Nazeeruddin, M. K.; Bruce, B. D.; Graetzel, M.; Zhang, S., Sci. Rep.

8

2012, 2, 1-7.

9 10

(12) Yehezkeli, O.; Tel-Vered, R.; Wasserman, J.; Trifonov, A.; Michaeli, D.; Nechushtai, R.; Willner, I., Nat. Commun. 2012, 3, 742.

11

(13) (a) Kirchhofer, N. D.; Rasmussen, M. A.; Dahlquist, F. W.; Minteer, S. D.; Bazan, G. C.,

12

Energ. Environ. Sci. 2015, 8, 2698-2706; (b) Hamidi, H.; Hasan, K.; Emek, S. C.; Dilgin, Y.;

13

Åkerlund, H.-E.; Albertsson, P.-Å.; Leech, D.; Gorton, L., ChemSusChem 2015, 8, 990-993; (c)

14

Calkins, J. O.; Umasankar, Y.; O'Neill, H.; Ramasamy, R. P., Energ. Environ. Sci. 2013, 6,

15

1891-1900.

16

(14) (a) Hasan, K.; Yildiz, H. B.; Sperling, E.; Conghaile, P. Ó.; Packer, M. A.; Leech, D.;

17

Hägerhäll, C.; Gorton, L., Phys. Chem. Chem. Phys. 2014, 16, 24676-24680; (b) Sarma, M. K.;

18

Kaushik, S.; Goswami, P., Biomass and Bioenergy. 2016, 90, 187-201.

19 20

(15) (a) Hasan, K.; Çevik, E.; Sperling, E.; Packer, M. A.; Leech, D.; Gorton, L., Adv. Energ. Mater. 2015, 5, 1501100; (b) Rosenbaum, M.; Schröder, U., Electroanalysis 2010, 22, 844-855.

21

(16) Stavrinidou, E.; Gabrielsson, R.; Gomez, E.; Crispin, X.; Nilsson, O.; Simon, D. T.;

22

Page 28 of 33

Berggren, M., Sci. Adv. 2015, 1, e1501136.


Page 29 of 33

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Catalysis

1

(17) (a) Bradley, R. W.; Bombelli, P.; Rowden, S. J. L.; Howe, C. J., Biochem. Soc. Trans.

2

2012, 40, 1302-1307; (b) Hasan, K.; Reddy, K. V. R.; Eßmann, V.; Górecki, K.; Conghaile, P.

3

Ó.; Schuhmann, W.; Leech, D.; Hägerhäll, C.; Gorton, L., Electroanalysis 2015, 27, 118-127.

4

(18) Edelman, M.; Mattoo, A. K., Photosyn. Res. 2008, 98, 609-620.

5

(19) Giraldo, J. P.; Landry, M. P.; Faltermeier, S. M.; McNicholas, T. P.; Iverson, N. M.;

6

Boghossian, A. A.; Reuel, N. F.; Hilmer, A. J.; Sen, F.; Brew, J. A., Nat. Mater. 2014, 13, 400-

7

408.

8 9 10 11 12 13

(20) Ryu, W.; Bai, S.-J.; Park, J. S.; Huang, Z.; Moseley, J.; Fabian, T.; Fasching, R. J.; Grossman, A. R.; Prinz, F. B., Nano Lett. 2010, 10, 1137-1143. (21) Amao, Y.; Tadokoro, A.; Nakamura, M.; Shuto, N.; Kuroki, A., Res. Chem. Intermed. 2014, 40, 3257-3265. (22) Kim, E.; Archibald, J., In The Chloroplast, Sandelius, A. S.; Aronsson, H., Eds. Springer: Berlin, Germany, 2009; p 1.

14

(23) Block, M. A.; Dorne, A.-J.; Joyard, J.; Douce, R., J. Biol. Chem. 1983, 258, 13281-13286.

15

(24) Okano, M.; Iida, T.; Shinohara, H.; Kobayashi, H.; Mitamura, T., Agric. Biol. Chem.

16

1984, 48, 1977-1983.

17

(25) Bhardwaj, R.; Pan, R. L.; Gross, E. L., Nature. 1981, 289, 396-398.

18

(26) Heller, A., Curr. Opin. Chem. Biol. 2006, 10, 664-672.

19

(27) Milton, R. D.; Hickey, D. P.; Abdellaoui, S.; Lim, K.; Wu, F.; Tan, B.; Minteer, S. D.,

20

Chem. Sci. 2015, 6, 4867-4875.


ACS Catalysis

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

1

(28) Arnon, D. I., Plant Physiol. 1949, 24, 1-15.

2

(29) Bukhov, N. G.; Sridharan, G.; Egorova, E. A.; Carpentier, R., BBA-Bioenergetics, 2003,

3 4 5

1604, 115-123. (30) Lichtenthaler, H. K.; Buschmann, C., In Current Protocols in Food Analytical Chemistry, Wrolstad, R. E., Ed. John Wiley & Sons, Inc.: NJ, USA, 2001; pp A2.1.1-A2.1.3.

6

(31) Lilley, R.; Fitzgerald, M.; Rienits, K.; Walker, D., New Phytol. 1975, 75, 1-10.

7

(32) Logan, B. E.; Hamelers, B.; Rozendal, R.; Schröder, U.; Keller, J.; Freguia, S.; Aelterman,

8 9

P.; Verstraete, W.; Rabaey, K., Environ. Sci. Technol. 2006, 40, 5181-5192. (33) McConnell, I.; Li, G.; Brudvig, G. W., Chem. Biol. 2010, 17, 434-447.

10

(34) Srivastava, A.; Strasser, R. J., J. Photochem. Photobiol. B. 1995, 31, 163-169.

11

(35) (a) Ochiai, H.; Shibata, H.; Fujishima, A.; Honda, K., Agric. Biol. Chem. 1979, 43, 881-

12

883; (b) Bulychev, A.; Andrianov, V.; Kurella, G.; Litvin, F., BBA-Bioenergetics. 1976, 430,

13

336-351.

14

(36) Roach, T.; Krieger-Liszkay, A. K., Curr. Prot. Pep. Sci. 2014, 15, 351-362.

15

(37) Melis, A., Trend. Plant Sci. 1999, 4, 130-135.

16

(38) Schröder, U., Phys. Chem. Chem. Phys. 2007, 9, 2619-2629.

17

(39) Heller, A.; Feldman, B., In Applications of Electrochemistry in Medicine, Schlesinger, M.,

18

Ed. Springer US: 2013; Vol. 56, pp 121-187.


Page 30 of 33

Page 31 of 33

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

1 2

ACS Catalysis

(40) Hickey, D. P., In Enzyme Stabilization and Immobilization: Methods and Protocols, Minteer, S. D., Ed. Springer New York: New York, NY, 2017; p 181.

3

(41) Reuillard, B.; Le Goff, A.; Cosnier, S., Anal. Chem. 2014, 86, 4409-4415.

4

(42) Plumeré, N.; Rüdiger, O.; Oughli, A. A.; Williams, R.; Vivekananthan, J.; Pöller, S.;

5 6 7 8 9 10 11 12 13

Schuhmann, W.; Lubitz, W., Nat. Chem. 2014, 6, 822-827. (43) (a) Zhu, Z.; Kin Tam, T.; Sun, F.; You, C.; Percival Zhang, Y. H., Nat. Commun. 2014, 5, 3026; (b) Giroud, F.; Milton, R. D.; Tan, B.-X.; Minteer, S. D., ACS Catal. 2015, 5, 1240-1244. (44) Milton, R., In Methods in molecular biology (Clifton, NJ), Minteer, S.D., Eds., Springer, New Jersery, 2017, Vol. 1504, p 193. (45) Coman, V.; Gustavsson, T.; Finkelsteinas, A.; Von Wachenfeldt, C.; Hägerhäl, C.; Gorton, L., J. Amer. Chem. Soc. 2009, 131, 16171-16176. (46) Walker, D. A.; Cerovic, Z. G.; Robinson, S. P., Methods in enzymology 1987, 148, 145157.

14

(47) Sekar, N.; Jain, R.; Yan, Y.; Ramasamy, R. P., Biotechnol. Bioeng. 2016, 113, 675-679.

15

(48) (a) Longatte, G.; Rappaport, F.; Wollman, F.-A.; Guille-Collignon, M.; Lemaître, F.,

16

Photochem. Photobiol. Sci. 2016, 15, 969-979; (b) Longatte, G.; Fu, H.-Y.; Buriez, O.; Labbé,

17

E.; Wollman, F.-A.; Amatore, C.; Rappaport, F.; Guille-Collignon, M.; Lemaître, F., Biophys.

18

Chem. 2015, 205, 1-8.

19 20

(49) Hasan, K.; Grippo, V.; Sperling, E.; Packer, M. A.; Leech, D.; Gorton, L., ChemElectroChem 2016, 4, 412-417.


ACS Catalysis

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

1 2

(50) Heiskanen, A.; Coman, V.; Kostesha, N.; Sabourin, D.; Haslett, N.; Baronian, K.; Gorton, L.; Dufva, M.; Emnéus, J., Anal.Bioanal.Chem. 2013, 405, 3847-3858.

3

(51) Hsu, B.-D.; Lee, J.-Y.; Pan, R.-L., Biochem. Biophys. Res. Commun. 1986, 141, 682-688.

4

(52) Rasmussen, M.; Wingersky, A.; Minteer, S. D., Electrochim. Acta 2014, 140, 304-308.

5 6 7 8


Page 32 of 33

Page 33 of 33

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Catalysis

table of contents entry 282x198mm (300 x 300 DPI)

ACS Paragon Plus Environment

Photobioelectrocatalysis of Intact Chloroplasts for Solar Energy

Recommend Documents