SUPRAMOLECULAR ASSEMBLY OF NANOSTUCTURED MATERIALS FOR
FABRICATION OF FUNCTIONAL INTERFACES
By
Ankush Ashok Gokhale

A DISSERTATION
Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of
Chemical Engineering-Doctor of Philosophy
2014

ABSTRACT
SUPRAMOLECULAR ASSEMBLY OF NANOSTRUCTURED MATERIALS FOR
FABRICATION OF FUNCTIONAL INTERFACES
By
Ankush Ashok Gokhale
In recent years, there has been increased focus towards miniaturization to build
sustainable devices by harnessing the power of synergistic nanoarchitectures. Drawing upon the
strength of the intermolecular forces, supramolecular assembly techniques have enabled the
inclusion of a host of inhomogeneous nanomaterials to give form and structure to macroscopic
objects. Supramolecular techniques are well suited to customize and improvise the assembly of
nanomaterials to allow full translation of their nanoscale properties to macroscopic structures. In
our work, we have primarily focused on developing novel supramolecular multifunctional
architectures for fabrication of nanostructured interfaces.
Firstly, we designed a unique recyclable two-tier carbonaceous biocatalyst consisting of a
polyelectrolyte modified iron-oxide doped graphene support and a biocatalytic component in the
form of cellulase, an enzyme capable of hydrolyzing lignocelluloses to reducing sugars. A
combination of strong and weak polyelectrolytes enabled long-term stabilization of hydrophobic
graphene platelets in aqueous medium and provided an effective means to control the
biocatalytic activity of the immobilized cellulase as a function of pH and temperature. We also
show how the effect of the dissociation of weak polyelectrolytes can overcome the geometric
disadvantage imposed by flat 2D supports. A combination of polyelectrolyte swelling, improved
colloidal stability and enhanced enzyme mobility contributed to preservation of the biocatalytic
activity.

Secondly, we also designed a novel supramolecular conductive interface consisting of
multilayered thin films of functionalized graphene platelets in conjunction with anionic
polyelectrolytes. The interplay of hydrogen bonding and electrostatic forces was studied and its
effect on film thickness and morphology was evaluated. Films developed in this work show
impressive gas shielding capabilities comparable to commercial grade silicon oxide coatings and
almost a five orders of magnitude reduction in sheet resistance.
Lastly, a bioelectronic interface to detect and quantify the presence of anionic nitrate
species was assembled from a surfactant stabilized supramolecular matrix. Nitrate based
fertilizers pose major health hazards when they combine with sources of drinking water. In this
work, we developed a novel 1D nanoarray of polyethylenedioxythiophene based nitrate
biosensor capable of detecting and quantifying minute concentrations of nitrate species. Low
detection limit (0.1 ppm), high signal to noise ratio and improved affinity towards nitrate species
as

compared

to

other

interfering

ions

are

significant

features

of

this

sensor.

Dedicated to my beloved parents for their love, motivation and unconditional support.

iv

ACKNOWLEDGMENTS

First and foremost, I would like to thank the Department of Chemical Engineering at Michigan
State University for placing faith in my abilities as a researcher and inducting me in the graduate
program. I am thankful to the department for its continued support in the form of fellowships,
assistantships, travel awards and easy access to research facilities that created a conductive
environment for scholarship and learning.
My advisor, Prof. Ilsoon Lee has been the most influential guiding force in my PhD years. I
wholeheartedly thank him for his tutelage and wonderful inputs. A special thanks to Dr. Jue Lu
and Ranjith Weerasiri from Technova Corporation for their guidance and feedback on my
projects. I would also like to thank my PhD committee members Prof. Wei Liao, Prof. David
Hodge and Prof. Robert Ofoli for their invaluable suggestions and comments during the PhD
program.
I would also like to thank Prof. Daina Briedis and Prof. Charles Petty in the Chemical
Engineering department for their help, support and advice especially during the first few years of
my PhD. Having served as a teaching assistant for Prof. Briedis for three semesters gave me a
chance to witness and appreciate the effort it takes to produce fifty minutes of quality lecture.
I would like to thank all the past and present members of our research group including Dr. Sumit
Mehrotra, Dr. Shaowen Ji, Dr. Wei Wang, Oishi Sanyal, Jing Yu and Anna Song for their help
and co-operation. I greatly appreciate the undergraduate students Nathan Parker and Andrew
Izbicki for their assistance in my research experiments. I would also like to thank all my officemates in 2522 EB : Dr. Xiangfeng Ma, Dr. Rui Lin, Ying Liu, Bhushan Awate and Yangmu Liu
for their friendship and companionship. It was great to have met and in some cases worked with
some interesting people : Ryan Stoklosa, Dan Williams, Tongjun Liu, Zhenglun Li, Muyang Li,
v

Rohit Nehe, Apoorav Shaligram and Ezhiylmurugan Rangaswamy. I thank them all for the
technical and non-technical discussions we had over the past five years. A special thanks to Lee
Alexander for all the water-cooler conversions we used to have. After my graduation, that is one
thing I am sure to miss.
Last but not the least; I thank my parents for their unconditional love and support. Back in the
day while I was still deciding the right grad school to go to, I distinctly recall my late father’s
words of encouragement to take up the offer to study at Michigan State University. Now on the
verge of completion, I do have a sense of satisfaction of a job well done. To have a part of your
family with you in new place is always a blessing. It was a great relief knowing that I always had
my brother Amit and my sister-in-law Swarupa to turn to if I needed help. Watching my niece
Aditi grow up from a baby into a toddler and now a preschooler has always brought me a sense
of joy.

vi

TABLE OF CONTENTS

LIST OF TABLES..........................................................................................................................ix
LIST OF FIGURES.........................................................................................................................x
Chapter 1: Introduction ...............................................................................................................1
1.1 Overview of Supramolecular chemistry .............................................................................1
1.2 Supramolecular nanostructured assemblies and nanobiocatalysis .......................................3
1.3 Fabrication of nanostructured interfaces using supramolecular techniques .........................6
REFERENCES ..................................................................................................................... 11
Chapter 2: Scope of the dissertation .......................................................................................... 16
2.1 Overview of the dissertation ............................................................................................ 16
2.2 Supramolecular modification of graphene platelets for the biocatalytic saccharification of
biomass ................................................................................................................................. 16
2.3 Conductive graphene-embedded films using a supramolecular layer-by-layer approach... 18
2.4 Nanostructured biosensor for nitrate detection and quantification .................................... 19
REFERENCES ..................................................................................................................... 21
Chapter 3: Supramolecular modification of graphene platelets for biocatalytic saccharification of
biomass ..................................................................................................................................... 23
3.0 Abstract ........................................................................................................................... 23
3. 1. Introduction ................................................................................................................... 23
3.2. Experimental procedures ................................................................................................ 29
3.2.1 Materials ...................................................................................................................... 29
3.2.2 Preparation of polyelectrolyte solutions ........................................................................ 29
3.2.3 Preparation of polyelectrolyte-multilayered graphene scaffolds .................................... 30
3.2.4 Alkaline co-precipitation of maghemite-magnetite nanoparticles (8-10 nm) .................. 30
3.2.5 Maghemite-Magnetite decoration of polyelectrolyte-modified graphene scaffolds ........ 30
3.2.6 Immobilization of cellulase on maghemite-magnetite graphene scaffolds ..................... 31
3.2.7 Characterization of polyelectrolyte coated graphene supports ....................................... 32
3.2.8 Characterization of GNPs and MNP-GNPs ................................................................... 32
3.2.9 Protein assay................................................................................................................. 32
3.2.10 Measurement of cellulase activities ............................................................................. 33
3.3. Results and Discussions .................................................................................................. 35
3.3.1 Maghemite-Magnetite decoration of graphene nanoplatelets ......................................... 36
3.3.2 Chemisorption of polyacrylic acid brushes on MNP-GNP ............................................ 41
3.3 Effect of PAA brushes on activity of immobilized enzymes ............................................. 43
3.3.3.1 Effect of pH on enzyme activity ................................................................................. 45
3.3.3.2 Effect of temperature on cellulase activity ................................................................. 48
3.3.3.3 Mimicking ability of Immobilized enzymes ............................................................... 50

vii

3.3.4 Magnetic separation, recovery and reuse of immobilized enzymes ................................ 51
3.4. Conclusion ..................................................................................................................... 52
REFERENCES ..................................................................................................................... 54
Chapter 4: Conductive oxygen barrier films using supramolecular assembly of graphene
embedded polyelectrolyte multilayers ....................................................................................... 63
4.0 Abstract ........................................................................................................................... 63
4.1 Introduction ..................................................................................................................... 63
4.2 Experimental Procedures ................................................................................................. 67
4.2.1 Materials ...................................................................................................................... 67
4.2.2 Layer-by-layer (LbL) assembly of polyelectrolyte modified GNP ................................. 67
4.2.3 Multilayers Characterizations ....................................................................................... 68
4.2.4 Oxygen Transmission Measurements ............................................................................ 69
4.3 Results and Discussion .................................................................................................... 69
4.3.1 Types of LbL deposited GnP/PEM films ...................................................................... 70
4.3.1.1 Hydrogen bonded films.............................................................................................. 71
4.3.1.2 Electrostatically assembled films ............................................................................... 72
4.3.1.3 Composite film .......................................................................................................... 73
4.3.2 Morphology and Thickness of GNP embedded PEM films ........................................... 73
4.3.3 Oxygen barrier properties of GNP/PEM multilayers ..................................................... 80
4.3.4 Electrical resistance of the GNP/PEM multilayers ........................................................ 84
4.4 Conclusion ...................................................................................................................... 86
REFERENCES ..................................................................................................................... 88
Chapter 5: Amperometric detection and quantification of nitrate ions using a highly sensitive
nanostructured PEDOT biosensor.............................................................................................. 95
5.0 Abstract ........................................................................................................................... 95
5.1 Introduction ..................................................................................................................... 95
5.2 Experimental ................................................................................................................. 100
5.2.1 Materials .................................................................................................................... 100
5.2.2 Preparation of template assisted PEDOT/enzyme nanowire electrode ......................... 100
5.2.3 Characterization of PEDOT/enzyme nanowires .......................................................... 102
5.3 Results and Discussion .................................................................................................. 102
5.3.1 Anodic polymerization of EDOT ................................................................................ 105
5.3.2 Chronoamperometric growth of PEDOT/enzyme nanowires ....................................... 106
5.3.3 Morphology of PEDOT /enzyme nanostructures ......................................................... 108
5.3.4 Amperometric detection of nitrate ions ....................................................................... 111
5.3.5 Effect of other reducible substrates on the nitrate reductase biosensor ......................... 120
5.3.6 Operational stability of the biosensor .......................................................................... 126
5.4 Conclusion .................................................................................................................... 128
REFERENCES ................................................................................................................... 130
Chapter 6: Conclusions and future work .................................................................................. 137
6.1 Conclusions ................................................................................................................... 137
6.2 Future outlook ............................................................................................................... 139
REFERENCES ................................................................................................................... 142
viii

LIST OF TABLES

Table 1.1 : Bond energies of different types of interactions. Reproduced partially with
permission from [14] ...............................................................................................................2
Table 3.1 : ζ-potential (mV) of modified GNP dispersions. The last component in each line
represents the terminal (topmost) layer in the multilayer assembly. Error bars represent the
standard deviation of 3 replicate samples. .............................................................................. 36
Table 3.2 : ζ-potential (mV) of IMC and IMC-PAA as a function of pH. Error bars represent
the standard deviation of 3 replicate samples. ........................................................................ 48
Table 4.1 : Oxygen permeability coefficient of hydrogen bonded films on PET substrate at
STP conditions ...................................................................................................................... 81
Table 4.2 : Oxygen permeability coefficient of electrostatic films on PET substrate at STP
conditions .............................................................................................................................. 82
Table 4.3 : Oxygen permeability coefficient of composite films on PET substrate at STP
conditions .............................................................................................................................. 82
Table 4.4 : Oxygen permeability coefficient of composite films (variable electrostatic layers)
on PET substrate at STP conditions ....................................................................................... 82
Table 5.1 : Estimation of kinetic parameter based on Lineweaver-Burk approximation........ 126
Table 5.2 : Operational parameters for different types of biosensors. ................................... 127

ix

LIST OF FIGURES

Figure 1.1 : Biocatalytic manipulation of supramolecular nanostructures. (reproduced with
permission from: [23]).............................................................................................................4
Figure 1.2 : Site-specific enzyme immobilization for construction of supramolecular
microfluidic chip. (Reproduced with permission from [26]) Note: Sav: biotin-Streptavidin .....5
Figure 1.3 : Supramolecularly assembled amphiphile (PP2b) for separation, immobilization
and biocatalytic utilization of enzymes/proteins. Reproduced from: [27] .................................6
Figure 1.4 : Schematic diagram illustrating the difference between top-down and bottom-up
approaches. Top down approach relies on controlled dismembering of larger blocks to get
nanosized materials. Bottom-up methods utilize controlled assembly of atoms, molecules as
for producing nanoscale interfaces. Reproduced with permission from: [30]............................7
Figure 1.5 : Supramolecular layer-by-layer assembly of polyelectrolytes for formation of thin
films on substrates (Reproduced with permission from [46]). ..................................................9
Figure 1.6 : Entrapment of glucose oxidase (GOx) in a supramolecular matrix of multiwall
carbon nanotubes (MWCNT) and β-cyclodextrin (CD). (Reproduced with permission from
[49]). ..................................................................................................................................... 10
Figure 3.1 : Schematic illustration of the design of the proposed biocatalyst: Assembly of
polyelectrolytes, nanoparticles and cellulase on graphene nanoplatelets. (Figure not drawn to
scale) ..................................................................................................................................... 35
Figure 3.2 : SEM micrographs of (a) GNP, (b) MNP-GNP (low magnification), and (c) MNPGNP (high magnification): Average size of iron oxide nanoparticles around 8-10 nm. (d)
TEM micrograph of GNP. The inset shows high magnification image of a selected portion of
the same platelet. Scale bar for the inset is 100 nm. TEM micrographs of (e) MNP-GNP (low
magnification) and (f) TEM micrograph of MNP-GNP (high magnification): Almost uniform
coating of iron-oxide nanoparticles (8-10 nm) is observed on the surface of the graphene
nanoplatelet. .......................................................................................................................... 39
Figure 3.3 : Effect of external magnetic field on graphene nanoplatelets (a) before iron oxide
nanoparticle decoration: No change observed (b) after iron-oxide nanoparticle decoration:
GNPs driven towards the magnetic source. ............................................................................ 40
Figure 3.4 : Effect of swollen PAA brushes on cellulase mobility. (a) Higher steric hindrance
due to collapsed PAA brushes. (b) Higher cellulase mobility and increased substrate-enzyme
interaction due to swollen PAA brushes. (Figure not drawn to scale) ..................................... 44
Figure 3.5 : Relative activity of free and immobilized enzymes as a function of pH (a)
Avicelase activity (b) CMCase activity. Conditions: 2%w/v avicel/CMC, Hydrolysis time: 90
x

minutes, Temperature 50 0C. Error bar represent standard deviation of three replicate samples.
.............................................................................................................................................. 46
Figure 3.6 : Relative activity of free and immobilized enzymes as a function of temperature (a)
Avicelase activity (b) CMCase activity. Conditions: (a)2 %w/v avicel (b)2%w/v CMC,
Hydrolysis time: 90 minutes, solution pH: 5.1. Error bars represent standard deviation of three
replicate samples. .................................................................................................................. 49
Figure 3.7 : Mimicking ability of immobilized enzymes as a function of (a) Avicelase activity
at variable pH (b) CMCase activity at variable pH (c) Avicelase activity at variable
temperature (d) CMCase activity at variable temperatureConditions: (a) 2%w/v avicel (b)
2%w/v CMC, Hydrolysis time: 90 minutes, Temperature 50 0C. Data was collected at pH 3.0,
3.7, 4.4 ,5.1, 5.8 and 6.5. (c) 2 %w/v avicel (d) 2%w/v CMC, Hydrolysis time: 90 minutes,
solution pH: 5.1. Data collected at 400C, 500C, 600C and 700C. Error bars represent standard
deviation of three replicate samples................................................................................... ........50
Figure 3.8 : Specific activity of immobilized enzymes with and without PAA brushes vs.
number of cycles of hydrolysis. ............................................................................................. 52
Figure 4.1 : Supramolecular assembly of graphene embedded polyelectrolyte multilayers.
(a)Multilayers assembled using hydrogen bonded assembly. m denotes number of bilayers
deposited. (b) Multilayers deposited using electrostatic interactions. n denotes the number of
electrostatic bilayers deposited. (c) Composite film consisting of ‘m’ number of hydrogen
bonded bilayers followed by ‘n’ number of electrostatic bilayers. .......................................... 71
Figure 4.2 : SEM micrographs of 20 bilayer GNP embedded PEM assembly. a, c, e show
sectional view of hydrogen bonded, electrostatic and composite (10H + 10E) multilayers
respectively. Thickness of the coating is 5.5 µm, 0.705 µm, 3.77 µm respectively. b, d, f show
surface morphology of hydrogen bonded, electrostatic and composite films respectively. ...... 75
Figure 4.3 : SEM micrographs of 4 bilayer GNP embedded PEM assembly. a, c, e show
sectional view of hydrogen bonded, electrostatic and composite (2H + 2E) multilayers
respectively. Thickness of the coating is 0.429 µm, 0.167 µm, 0.378 µm respectively. b, d, f
show surface morphology of hydrogen bonded, electrostatic and composite films respectively.
.............................................................................................................................................. 77
Figure 4.4 : SEM micrographs of 10 bilayer GNP embedded PEM assembly. a, c, e show
sectional view of hydrogen bonded, electrostatic and composite (5H + 5E) multilayers
respectively. Thickness of the coating is 3.47 µm, 0.455 µm, 0.893 µm respectively. b, d, f
show surface morphology of hydrogen bonded, electrostatic and composite films respectively.
.............................................................................................................................................. 78
Figure 4.5 : SEM micrographs of composite films with fixed number of hydrogen bonded
multilayers (10 bilayers) and variable number of electrostatic layers a and c show sectional
view of composite films with (10H + 5E) and 10H +20 E) multilayers respectively. Thickness

xi

of the coating is 3.56µm and 4.02 µm respectively. b and d show surface morphology of the
composite films (10H +5E) and (10H + 20E) respectively. .................................................... 79
Figure 4.6 : Oxygen transmission rate (OTR) for different types of configurations. (a) 4 bilayer
system. The composite film consists of 2 hydrogen bonded and 2 electrostatic bilayers. (b) 10
bilayer system. Composite film consists of 5 hydrogen bonded and 5 electrostatic bilayers. (c)
20 bilayer system. Composite film consists of 10 hydrogen bonded and 10 electrostatic
bilayers. (d) Composite films consisting of 10 hydrogen bonded (H) followed by varying
number of electrostatic (E) bilayers. ...................................................................................... 81
Figure 4.7 : Sheet resistance for different configurations of LbL assembled graphene
embedded multilayers Sheet resistance at 1 Hz for various samples is as follows:
(a) Bare PET film: 111.1 GΩ, hydrogen bonded film: 108.2 GΩ, electrostatic film: 61.3 GΩ,
composite film: 57.6 GΩ.
(b) Bare PET film: 115.2 GΩ, hydrogen bonded film: 90.9 GΩ, electrostatic film: 54.3 GΩ ,
composite film: 50.2 GΩ.
(c) Bare PET film: 110.2 GΩ, hydrogen bonded film: 40.8 GΩ, electrostatic film: 0.949 GΩ,
composite film: 24.8 MΩ.
(d) Bare PET film: 112.3 GΩ, composite film (10H + 5 E): 31.2 GΩ, (10H + 10 E): 24.8 MΩ,
(10H+20 E): 1.14 MΩ...............................................................................................................85
Figure 5.1 : Electrochemical polymerization of EDOT using a templated approach. (a)
Formation of polymer nanotubes (b) Formation of polymer nanowires ................................ 103
Figure 5.2 : Linear sweep voltammetry (LSV) profile of EDOT polymerization (a) in absence
of enzyme (b) in presence of enzyme. .................................................................................. 106
Figure 5.3 : Chronoamperometric plot of current density as a function of
electropolymerization time. Inclusion of enzyme reduces the creation of active sites for
polymerization evidenced by lower current density during the initial phase of polymerization.
............................................................................................................................................ 107
Figure 5.4 : Microscopy images of PEDOT nanowires at different oxidation potential. SEM
images of nanowires polymerized at (a) 0.85 V (c) 1.1 V (e) 1.4 V. Corresponding TEM
images (b) 0.85 V (d) 1.1 V (f) 1.4 V................................................................................... 110
Figure 5.5 : Digital images of PEDOT in dichloromethane solution. EDOT polymerized at
0.85 V yields clear solution indicative of low degree of polymerization. At 1.1 and 1.4 V,
bluish-black PEDOT is formed. ........................................................................................... 111
Figure 5.6 : Amperometric profile of nitrate biosensor fabricated at polymerization potential of
0.85 V. Detection of nitrate species is possible but quantification information is hard to
establish. ............................................................................................................................. 113

xii

Figure 5.7 : Amperometric profile of nitrate biosensor fabricated at polymerization potential of
1.4 V. Detection of nitrate species is possible but no quantification information is hard to
establish. ............................................................................................................................. 113
Figure 5.8 : TEM micrographs of PEDOT nanowires fabricated at polymerization potential of
1.4 V. (a) length of nanowire close to 6.5 microns (b) length of nanowire close to 9 microns.
Growth is irregular. ............................................................................................................. 114
Figure 5.9 : (a) Amperometric profile of nitrate biosensor fabricated at 1.1 V, 300 sec. in
presence of nitrate species. Inset image shows step-wise increase in current after addition of
200 ppb nitrate after every 50 seconds (b) Linear range of operation of the sensor. .............. 116
Figure 5.10 : Amperometric profile of nitrate biosensor fabricated at polymerization potential
of 0.95 V (300 sec.). The nitrate species are detected but quantitative trend difficult to
establish. ............................................................................................................................. 118
Figure 5.11 : (a) Amperometric profile of nitrate biosensor fabricated at 0.95 V, 600 sec. in
presence of nitrate species. Inset image shows step-wise increase in current after addition of
200 ppb nitrate after every 50 seconds (b) Linear range of operation of the sensor. .............. 119
Figure 5.12 : (a) Amperometric profile of nitrate biosensor fabricated at 1.1 V, 300 sec. in
presence of perchlorate species. Inset image shows step-wise increase in current after addition
of 2.5 ppm perchlorate after every 50 seconds (b) Linear range of operation of the sensor. .. 121
Figure 5.13 : (a) Amperometric profile of nitrate biosensor fabricated at 1.1 V, 300 sec. in
presence of carbonate species. Inset image shows step-wise increase in current after addition
of 25 ppm nitrate after every 50 seconds (b) Linear range of operation of the sensor. .......... 122
Figure 5.14 : Lineweaver-Burk plots for nitrate biosensor (a) Sensor fabricated at 1.1 V, 300
sec. in presence of nitrate species. (b) Sensor fabricated at 0.95 V, 600 sec. in presence of
nitrate species. (c) Sensor fabricated at 1.1 V, 300 sec. in presence of perchlorate species. (d)
Sensor fabricated 1.1V, 300 sec. in presence of carbonate species. ...................................... 124

xiii

Chapter 1: Introduction
1.1 Overview of Supramolecular chemistry
Supramolecular assembly is a branch of chemistry that studies the interplay of
noncovalent interactions and its effect on organization of an ensemble of discrete atoms, ions and
molecules. While molecular chemistry is dominated by covalent forces, supramolecular
chemistry relies on subtle intermolecular forces that exist freely in nature[1]. While initially
confined to host-guest systems, the definition of supramolecular chemistry has now been
expanded to explain a whole range of phenomena. Biological systems starting from natural
molecules such as proteins, oligonuclueotides and lipids to larger complex applications such as
targeted drug release, enzyme-substrate interactions, self-assembly and interactions between
cells, tissues and pathogens (also known as polyvalency) are reported to rely on the
supramolecular forces[2, 3]. Most biological systems are complex crowded biochemical
environments which require information to flow to specific interactive species[4]. Through
supramolecular interactions, molecular information can be easily transferred between the
interacting species with minimum signal dilution due to other interfering species. As a result,
specialized molecular receptors with high degree of specificity to recognize the pattern of flow of
information can be utilized for supramolecular catalysis and selective transport processes[5]. In
recent years, several examples of supramolecular catalysis ranging from biocatalytic
conversions[6, 7] to metal-complex transformations[8-10] have been reported. Similarly
examples of transport processes as a result of supramolecular interactions are also reported[11,
12].

1

The intermolecular interactions as a result of the supramolecular arrangement of the
species are roughly classified into two types depending on the strength of the forces. Isotropic
forces are medium range interactions that give shape, size and structural packing to the material
while anisotropic interactions are long range directional interactions that can exist in charged
species[13]. Supramolecular interactions cover wide spectra of weak noncovalent interactions.
Table 1 lists the different types of interactive forces and provides a comparative analysis of their
respective bond energies.
Table 1.1 Bond energies of different types of interactions. Reproduced partially with permission
from [14]
Type of interaction
Covalent bond
Hydrogen bond
Vander Waal forces
Co-ordination binding
‘fit interaction’*
‘amphiphilic’**
ionic
Hydrophobic effects
π-π stacking forces

Bond energy (kJ/mole)
350
10-65
<5[15]
50-200
10-100
5-50
50-250 (depends on solvent
and ions)
Difficult to access accurately
0-50[15]

Character
Irreversible
Selective, directional
Non-selective, non-directional
Directional
Very selective
Non-selective
Non-selective
Weakly directional

*: precise fitting of surfaces based on geometric limitations (lock-key principle)
**: observed in surfactant micelles
Among the different supramolecular forces, a few have been highlighted here due to their
versatility and unique applications. Hydrogen bonds display a wide window of bond energies.
The formation of the hydrogen bond is a result of interaction between a donor containing an
acidic hydrogen atom and an acceptor containing lone pair of electrons[15]. In some cases,
hydrogen bonds assume the proportion of a quasi-covalent bonds with bond energies rivaling
those of covalent interactions whereas sometimes they can be as weak as Vander Waal

2

forces[16]. These forces are an example of powerful selective interactions that can be assembled
using appropriate solvents and are completely reversible at room temperature. Similarly, though
inherently weak, nature bears testimony to the collective effect of a large number of Vander
Waal forces of interaction. Examples such as a house-lizard adhering to a wall or ceiling as a
result a Vander Waal forces of attraction acting through the fine hair-like structures located on its
footpad abound in nature[13, 17]. π-π stacking forces (aromatic-aromatic) are another example of
secondary interactions. These interactions include face-to-face, edge-to-face and offset,slipped
interactions[18]. Aromatic interactions are hypothesized to be a combination of a variety of
noncovalent forces such as Vander Waal, hydrophobic and electrostatic. The hydrophobic
element in these interactions makes them strong in water whereas the electrostatic component
makes it selective[19]. In recent years, there has been increased focus on using supramolecular
techniques for stabilization and functionalization of a number of nanomaterials. These techniques
can provide robust bottom-up strategies to assemble a broad spectrum of nanostructures on the
host substrates. Multifunctional tailor-made substrates fabricated using this route, have found
applications in catalysis, sensing, and drug delivery.

1.2 Supramolecular nanostructured assemblies and nanobiocatalysis
The synergistic combination of supramolecular nanostructures and biocatalysis provides
a unique opportunity to combine specificity (due to biocatalysis) and ‘smart’ materials (due to
supramolecular assembly techniques). Higher ordered structures can be easily built by tweaking
the biocatalytic component. It has been reported that the supramolecular assembly of amphiphilic
molecules can be manipulated by a biocatalytic process induced by different concentrations of
enzyme resulting in structurally diverse nanoassemblies[20]. In other words, supramolecular
structures of lowest free energy configurations can be obtained by manipulating the biocatalytic
3

process. Figure 1 shows an example of the biocatalytic modification of a supramolecular
assembly process. The left part of the image shows the assembly of peptides through selfassembly. The image on the right shows cleavage of the methyl group as a result of enzymatic
hydrolysis and the resultant self-assembled peptide structure. Another example where
biocatalysis has proven to be of great utility is formation of supramolecular hydrogels. Unlike
chemically cross-linked polymeric hydrogels, supramolecular hydrogels are formed when selfassembly of small molecules result in formation of nanofiber networks[21]. Gao and coworkers[22] used an enzymatic reaction to trigger the self-assembly of a taxol derivative to form
a nanofiber structure.

Figure 1.1 Biocatalytic manipulation of supramolecular nanostructures. (reproduced with
permission from: [23])
Just as biocatalysis can assist in formation of supramolecular assemblies, supramolecular
nanostructures can also aid in facilitating biocatalytic conversions. Freeman et. al.[24] reported
the formation of supramolecular cocaine-aptamer complexes that stimulated the biocatalyzed
oxidation of ethanol. Similarly, the supramolecular assembly of nucleic acid-hemin complex can

4

act as a biocatalyst producing chemiluminescence[25]. Another example of a supramolecular
arrangement to facilitate biocatalytic conversion is provided by Gonzales-Campo et. al.[26] The
authors have shown an interesting site-specific enzyme immobilization method to construct a
supramolecular microfluidic chip for biocatalytic transformations.

Figure 1.2 Site-specific enzyme immobilization for construction of supramolecular microfluidic
chip. (Reproduced with permission from [26]) Note: Sav: biotin-Streptavidin
Similarly, supramolecularly assembled amphiphiles for the formation of biocompatible
membranes for filtration, immobilization and biocatalytic utilization of proteins/enzymes has
been recently reported[27]. Here, the authors have illustrated a good example of fabricating
immobilization supports that can be easily recycled and reused over multiple cycles. Figure 3
shows a schematic diagram for the formation of these supramolecularly assembled membranes.

5

Figure 1.3 Supramolecularly assembled amphiphile (PP2b) for separation, immobilization and
biocatalytic utilization of enzymes/proteins. Reproduced from: [27]
1.3 Fabrication of nanostructured interfaces using supramolecular techniques
The use of supramolecular techniques for fabrication of nanostructured interfaces has
enjoyed tremendous success due to versatility, flexible nature of the noncovalent interfaces and
precise control that these techniques offer. Supramolecular nanostructured interfaces have carved
a place for themselves in almost every application such as thin films, sensors and
semiconductor/electronics industry. Two approaches commonly used for fabrication of
nanostructured interfaces include the ‘top-down’ approach and the ‘bottom-up’ approach. A
schematic diagram shown in Figure 4 shows the difference in the two approaches. Top down
approach which includes techniques such as photolithography or electron lithography has shown
a remarkable ability to commercially produce nanointerfaces with great precision. However, in
the sub-100 nm region this approach faces several challenges [28, 29]. Bottom-up approach on
the other hand utilizes smaller building blocks such as atoms, molecules to produce larger

6

structures. Hence this approach is well-suited to produce nanostructures in nanoscale regimes
where top-down approach fails or faces technological challenges.

Size Reduction
Techniques

Size
Augmentation
Techniques

Figure 1.4 Schematic diagram illustrating the difference between top-down and bottom-up
approaches. Top down approach relies on controlled dismembering of larger blocks to get
nanosized materials. Bottom-up methods utilize controlled assembly of atoms, molecules as for
producing nanoscale interfaces. Reproduced with permission from: [30]
The application of supramolecular chemistry to bottom-up approach presents exciting
opportunities to fabricate multifunctional interfaces. Though it is not possible to introduce all
such applications, keeping in mind the scope of this dissertation a few examples pertaining to
thin film fabrication and development of nanostructured interfaces for biosensor applications are
provided here.

7

Controlled deposition of different types and sizes of nanoparticles in the form of thin
films can find wide range of applications due to difference in fluorescence, absorption and other
properties arising out of contributions from individual components[31]. Aggregation is a
common problem in such systems. To overcome the same, Kao and co-workers[31] recently
developed block co-polymer (BCP) supramolecular approach. The authors used the rigid combblocks in the BCP polymer to orient multi-dimensional (1D chains, 2D lattice and 3D arrays) of
a mixture of nanoparticles by spatial arrangement of these particles in the supramolecular
network. Similarly, Tang and co-workers[32] reported the formation of block-copolymer
assemblies with tunable supramolecular interactions depending on stoichiometry and the number
of hydrogen bonds and studied their effect on film morphology. Several other examples
including

layer-by-layer

assembly (LbL)[33-40],

Langmuir-Blodgett[41,

42],

hot-wall

process[43], molecular beam deposition process[44] involving supramolecular interactions for
effective growth of thin films are available in literature. An interesting review on fabrication of
thin films using supramolecular polymers is presented by Tsukruk[45]. Figure 5 shows the
growth of supramolecular thin films using the sequential LbL approach. More details about this
process will be covered in the subsequent chapters.

8

Figure 1.5 Supramolecular layer-by-layer assembly of polyelectrolytes for formation of thin
films on substrates (Reproduced with permission from [46]).
Just like thin films, the application of supramolecular assemblies for biosensor
technology also holds great promise. Most sensors comprise of two critical elements: the sensing
element for the detection of the signal and a transducer to convert the signal into desired output.
Supramolecular techniques provide excellent opportunities to integrate these two elements. One
of the most common techniques involve guest-host supramolecular interactions for the
immobilization of enzymes, cells, nanoparticles on desired substrates to fabricate stable
biosensors. Villalonga and co-workers[47] used the same strategy to immobilize xanthine
oxidase on a hybrid electrochemical platform consisting of gold nanoparticles and carbon
nanotubes. Miranda and co-workers[48] exploited the electrostatic interactions between cationic
gold nanoparticles and negatively charge β-galactosidase to design a biosensor for detection of
colorimetric bacteria. In another example Alarcon-Angeles and co-workers[49] used a hybrid
supramolecular matrix consisting of carbon nanotubes and electropolymerized β-cyclodextrin
(CD)-glucose oxidase that suited the microenvironment for effective operation of the enzyme.
Figure 6 shows the inclusion of various components in biosensor design.

9

Figure 1.6 Entrapment of glucose oxidase (GOx) in a supramolecular matrix of multiwall carbon
nanotubes (MWCNT) and β-cyclodextrin (CD). (Reproduced with permission from [49]).

10

REFERENCES

11

REFERENCES

1.

Lehn, J.-M., Toward complex matter: Supramolecular chemistry and self-organization.
Proceedings of the National Academy of Sciences, 2002. 99(8): p. 4763-4768.

2.

Uhlenheuer, D.A., K. Petkau, and L. Brunsveld, Combining supramolecular chemistry with
biology. Chemical Society Reviews, 2010. 39(8): p. 2817-2826.

3.

Badjic, J.D., et al., Multivalency and cooperativity in supramolecular chemistry. Accounts
of chemical research, 2005. 38(9): p. 723-732.

4.

Savir, Y. and T. Tlusty. Molecular recognition as an information channel: The role of
conformational changes. in Information Sciences and Systems, 2009. CISS 2009. 43rd
Annual Conference on. 2009. IEEE.

5.

Lehn, J.-M., From supramolecular chemistry towards constitutional dynamic chemistry
and adaptive chemistry. Chem. Soc. Rev., 2007. 36(2): p. 151-160.

6.

Elemans, J.A., et al., Bio‐Inspired Supramolecular Catalysis. Supramolecular Catalysis,
2009: p. 143-164.

7.

Wiester, M.J., P.A. Ulmann, and C.A. Mirkin, Enzyme mimics based upon supramolecular
coordination chemistry. Angewandte Chemie International Edition, 2011. 50(1): p. 114137.

8.

Brown, C.J., et al., High-turnover supramolecular catalysis by a protected ruthenium (II)
complex in aqueous solution. Journal of the American Chemical Society, 2011. 133(31):
p. 11964-11966.

9.

Wang, Z.J., et al., Hydroalkoxylation catalyzed by a gold (I) complex encapsulated in a
supramolecular host. Journal of the American Chemical Society, 2011. 133(19): p. 73587360.

10.

Yoon, H.J., et al., Allosteric supramolecular triple-layer catalysts. Science, 2010.
330(6000): p. 66-69.

11.

Sakai, N., et al., Supramolecular n/p-heterojunction photosystems with antiparallel redox
gradients in electron-and hole-transporting pathways. Journal of the American Chemical
Society, 2010. 132(20): p. 6923-6925.

12

12.
13.

Huettner, S., et al., Tunable charge transport using supramolecular self-assembly of
nanostructured crystalline block copolymers. ACS nano, 2011. 5(5): p. 3506-3515.
Desiraju, G.R., Chemistry beyond the molecule. Nature, 2001. 412(6845): p. 397-400.

14.

Faul, C.F.J. and M. Antonietti, Ionic Self-Assembly: Facile Synthesis of Supramolecular
Materials. Advanced Materials, 2003. 15(9): p. 673-683.

15.

Hoeben, F.J.M., et al., About Supramolecular Assemblies of π-Conjugated Systems.
Chemical Reviews, 2005. 105(4): p. 1491-1546.

16.

Jeffrey, G.A., An introduction to hydrogen bonding. Vol. 12. 1997: Oxford University
Press New York.

17.

Santos, D., et al., Directional adhesion for climbing: theoretical and practical
considerations. Journal of Adhesion Science and Technology, 2007. 21(12-13): p. 13171341.

18.

Roesky, H.W. and M. Andruh, The interplay of coordinative, hydrogen bonding and π–π
stacking interactions in sustaining supramolecular solid-state architectures.: A study
case of bis(4-pyridyl)- and bis(4-pyridyl-N-oxide) tectons. Coordination Chemistry
Reviews, 2003. 236(1–2): p. 91-119.

19.

Waters, M.L., Aromatic interactions in model systems. Current opinion in chemical
biology, 2002. 6(6): p. 736-741.

20.

Hirst, A.R., et al., Biocatalytic induction of supramolecular order. Nat Chem, 2010. 2(12):
p. 1089-1094.

21.

Yang, Z. and B. Xu, Supramolecular hydrogels based on biofunctional nanofibers of selfassembled small molecules. Journal of Materials Chemistry, 2007. 17(23): p. 2385-2393.

22.

Gao, Y., et al., Enzyme-Instructed Molecular Self-assembly Confers Nanofibers and a
Supramolecular Hydrogel of Taxol Derivative. Journal of the American Chemical Society,
2009. 131(38): p. 13576-13577.

23.

Gazit, E., Bioinspired chemistry: Diversity for self-assembly. Nature Chemistry, 2010.
2(12): p. 1010-1011.

24.

Freeman, R., et al., Supramolecular cocaine− aptamer complexes activate biocatalytic
cascades. Journal of the American Chemical Society, 2009. 131(14): p. 5028-5029.

25.

Xiao, Y., et al., Lighting Up Biochemiluminescence by the Surface Self-Assembly of DNA–
Hemin Complexes. ChemBioChem, 2004. 5(3): p. 374-379.
13

26.
27.

González-Campo, A., et al., A Supramolecular Approach to Enzyme Immobilization in
Micro-Channels. Small, 2012. 8(22): p. 3531-3537.
Krieg, E., et al., Separation, Immobilization, and Biocatalytic Utilization of Proteins by a
Supramolecular Membrane. PLoS ONE, 2013. 8(5): p. e63188.

28.

Teo, B. and X.H. Sun, From Top-Down to Bottom-Up to Hybrid Nanotechnologies: Road
to Nanodevices. Journal of Cluster Science, 2006. 17(4): p. 529-540.

29.

Balzani, V., A. Credi, and M. Venturi, The Bottom-Up Approach to Molecular-Level
Devices and Machines. Chemistry – A European Journal, 2002. 8(24): p. 5524-5532.

30.

Sanchez, F. and K. Sobolev, Nanotechnology in concrete – A review. Construction and
Building Materials, 2010. 24(11): p. 2060-2071.

31.

Kao, J., et al., Size-Dependent Assemblies of Nanoparticle Mixtures in Thin Films. Journal
of the American Chemical Society, 2013. 135(5): p. 1680-1683.

32.

Tang, C., et al., Thin film morphology of block copolymer blends with tunable
supramolecular interactions for lithographic applications. Macromolecules, 2010. 43(6):
p. 2880-2889.

33.

Crespo-Biel, O., et al., Supramolecular Layer-by-Layer Assembly:  Alternating
Adsorptions of Guest- and Host-Functionalized Molecules and Particles Using Multivalent
Supramolecular Interactions. Journal of the American Chemical Society, 2005. 127(20):
p. 7594-7600.

34.

Decher, G., Fuzzy nanoassemblies: toward layered polymeric multicomposites. Science,
1997. 277(5330): p. 1232-1237.

35.

Decher, G. and J. Hong, Buildup of ultrathin multilayer films by a self‐assembly process:
II. Consecutive adsorption of anionic and cationic bipolar amphiphiles and
polyelectrolytes on charged surfaces. Berichte der Bunsengesellschaft für physikalische
Chemie, 1991. 95(11): p. 1430-1434.

36.

Gokhale, A.A., et al., Conductive oxygen barrier films using supramolecular assembly of
graphene embedded polyelectrolyte multilayers. Journal of colloid and interface science,
2013.

37.

Gokhale, A.A., J. Lu, and I. Lee, Immobilization of cellulase on magnetoresponsive
graphene nano-supports. Journal of Molecular Catalysis B: Enzymatic, 2013. 90(0): p. 7686.

38.

Zheng, H., et al., Two component particle arrays on patterned polyelectrolyte multilayer
templates. Advanced Materials, 2002. 14(8): p. 569.
14

39.

Kidambi, S., I. Lee, and C. Chan, Controlling primary hepatocyte adhesion and spreading
on protein-free polyelectrolyte multilayer films. Journal of the American Chemical
Society, 2004. 126(50): p. 16286-16287.

40.

Hendricks, T.R., et al., Intact pattern transfer of conductive exfoliated graphite
nanoplatelet composite films to polyelectrolyte multilayer platforms. Advanced
Materials, 2008. 20(10).

41.

Ni, Y., R.R. Puthenkovilakom, and Q. Huo, Synthesis and Supramolecular Self-Assembly
Study of a Novel Porphyrin Molecule in Langmuir and Langmuir−Blodgett Films.
Langmuir, 2004. 20(7): p. 2765-2771.

42.

Bjørnholm, T., T. Hassenkam, and N. Reitzel, Supramolecular organization of highly
conducting organic thin films by the Langmuir-Blodgett technique. Journal of Materials
Chemistry, 1999. 9(9): p. 1975-1990.

43.

Khan, R.U.A., et al., Supramolecular Ordered Organic Thin Films for Nonlinear Optical
and Optoelectronic Applications. Advanced Functional Materials, 2006. 16(2): p. 180188.

44.

Esembeson, B., et al., A High‐Optical Quality Supramolecular Assembly for Third‐Order
Integrated Nonlinear Optics. Advanced Materials, 2008. 20(23): p. 4584-4587.

45.

Tsukruk, V.V., Assembly of supramolecular polymers in ultrathin films. Progress in
Polymer Science, 1997. 22(2): p. 247-311.

46.

Wohl, B.M. and J.F.J. Engbersen, Responsive layer-by-layer materials for drug delivery.
Journal of Controlled Release, 2012. 158(1): p. 2-14.

47.

Villalonga, R., et al., Supramolecular Immobilization of Xanthine Oxidase on
Electropolymerized Matrix of Functionalized Hybrid Gold Nanoparticles/Single-Walled
Carbon Nanotubes for the Preparation of Electrochemical Biosensors. ACS Applied
Materials & Interfaces, 2012. 4(8): p. 4312-4319.

48.

Miranda, O.R., et al., Colorimetric bacteria sensing using a supramolecular enzyme–
nanoparticle biosensor. Journal of the American Chemical Society, 2011. 133(25): p.
9650-9653.

49.

Alarcón-Ángeles, G., et al., Enzyme entrapment by β-cyclodextrin electropolymerization
onto a carbon nanotubes-modified screen-printed electrode. Biosensors and
Bioelectronics, 2010. 26(4): p. 1768-1773.

15

Chapter 2: Scope of the dissertation
2.1 Overview of the dissertation
The emergence of nanotechnology tools has made it possible to readily synthesize and
characterize nanostructured materials. The application of supramolecular chemistry has provided
the means to organize these nanostructures in an orderly manner by fine-tuning the
intermolecular forces needed for effective binding of the different nanoscale components[1].
This dissertation is an effort in that direction. By exploring different supramolecular interactions,
novel architectures for three different types of applications are presented here. The first
application deals with bringing about novel structural changes in the design of a supramolecular
carbonaceous enzymatic nanobiocatalyst so that it tunes well with the microenvironment it
operates. The second application deals with development of synergistic supramolecular thin
films with potential applications in electronics/packaging industry. The third project shows the
electropolymerization of 1D polymer/enzyme nanowires from a supramolecularly assembled
surfactant-stabilized matrix to serve as an amperometric sensor for detection and quantification
of anionic species. A brief description of each of these applications is provided here.

2.2 Supramolecular modification of graphene platelets for the biocatalytic saccharification
of biomass
There has been growing interest in cellulosic ethanol as the next generation fuel in recent
years[2, 3]. Depleting fossil fuels, concerns about energy security as well as magnanimous
government incentives in the form of subsidies and tax-breaks have positioned cellulosic ethanol
to be a strong contender as an alternative energy source[4, 5]. With several governments around
the world recommending the use of bioethanol blended fuel to meet the ever increasing energy

16

demands, there has been considerable effort to find a practical and economically feasible solution
to production of ethanol from renewable resources. The enzymatic conversion of lignocelluloses
to reducing sugars by cellulase is the key step in the production of bioethanol. According to
some studies, in a typical hydrolysis process cellulases and β-glucosidases account for as much
as 50% of the total costs[6, 7]. A truly cost efficient process would therefore demand evolving
sustainable routes to ensure recovery of cellulase followed by its reuse. Immobilization of
enzymes on solids supports is one such route.

Functional nanomaterials are promising

candidates for enzyme immobilization with a view to developing efficient industrial biocatalysts
with tailor-made catalytic properties. In the first project (Chapter 3), the preparation of pH
tunable, temperature sensitive magnetoresponsive graphene-based nano-bio carriers for cellulase
immobilization is reported. A simple route to overcome the geometric disadvantage imposed by
most 2D immobilization supports and make them capable of closely mimicking free enzymes
(FE) operating under similar reaction conditions is discussed. The supramolecular assembly of
oppositely charged quenched polyelectrolytes and maghemite-magnetite nanoparticles on 2D
graphene supports followed by immobilization of cellulase shows a marked improvement in the
biocatalytic activity of the graphene supports. The incorporation of magnetic nanoparticles opens
up the possibility of recovery and reuse of the enzyme over multiple cycles. Cellulase
immobilization is achieved by a combination of weak polyelectrolyte brushes and zero-length
spacer molecules. The swelling behavior of annealed polyelectrolyte brushes is a strong function
of the environmental conditions. The degree of polyelectrolyte swelling can be easily tweaked by
manipulating the pH and temperature, providing us an effective tool to control the activity of
immobilized enzymes. Activity of immobilized cellulase is evaluated using both soluble as well
as insoluble substrates like 2% w/v CMC and avicel respectively.

17

2.3 Conductive graphene-embedded films using a supramolecular layer-by-layer approach
In recent years, with a view to move up in the value-chain, there has been a growing
trend towards integrating functionalized nanostructures in packaging films. Packaging films are
projected to constitute nearly 42% of the 100 million ton polymer market[8]. Inclusion of
nanomaterials in during the fabrication of packaging films is expected to add synergy as well
improve the economics of the process. Nano-enabled packaging materials especially food and
beverage packaging materials grew significantly to reach US $4.13 billion in 2008 and are
expected to reach US $7.3 billion in 2014[8]. Besides, food packaging, the use of nano-enabled
barrier films in the electronics industry is also gaining ground. Siemens has been developing
nanotechnology enabled color displays on packaging films for the past few years[9]. Another
area where barrier films play a significant role is in fabrication of OLEDs. Active matrix organic
light emitting devices (AMOLED) currently occupy 2.3% of the mobile phone display
market[10]. But OLEDs can operate on lower battery power as compared to its traditional rivals.
It is estimated that by 2015, AMOLEDs would occupy as much as 40% of the market share[10].
This presents a huge market for gas barrier packaging films.
The supramolecular self-assembly of polyelectrolyte multilayers (PEMs) provides robust
bottom-up strategies to assemble a broad spectrum of nanostructures on the host substrates. The
idea of using suramolecularly assembled polyelectrolytes for graphene modification suggested in
the first project was further expanded to bring other types of applications under its ambit. In the
second project (chapter 4) the formation of graphene nanoplatelet (GNP) embedded
polyelectrolyte films to enhance the oxygen barrier properties of poly (ethylene terephthalate)
(PET) films is discussed. Despite cheaper costs and high mechanical strength, the diffusion of
small gas molecules such as oxygen through PET films remains a matter of great concern. The

18

simple yet robust supramolecular deposition of GNP/polyelectrolyte on PET substrates
significantly increases the tortuous path the oxygen molecule has to travel making it harder to
diffuse through the PET film. With permeability coefficients in the range of 10 -18
cc.cm/cm2.s.Pa, the coatings developed in this study show three orders of magnitude reduction as
compared to the permeability coefficient of the bare PET film, significantly lower than that of
ethylene vinyl alcohol (EVOH) and comparable to silicon oxide thin films used in commercial
gas barrier foils. The use of GNPs in the multilayered films also helped reduce the electrical
sheet resistance to about 1 MΩ which is five orders of magnitude lower than the original PET
substrate opening up promising opportunities for future use in semiconductor and electronics
industry. Making suitable modifications in the deposition process, three configurations of GNP
embedded PEM multilayers namely hydrogen bonded, electrostatic and composite films were
developed and their effect on oxygen barrier property and sheet resistance was monitored.
Oxygen permeability of films was tested in accordance with ASTM D-3985 using a MOCON
2/21 ML instrument whereas electrical sheet resistance was quantified using a Gamry Femtostat
Electrochemical Impedance station.

2.4 Nanostructured biosensor for nitrate detection and quantification
Nanostructured biosensors provide a unique amalgamation of high aspect ratio
nanomaterials and substrate specific biological components. The use of nanostructured
enzymatic biosensors in environmental monitoring is fast emerging as an important tool to
provide valuable inputs on the presence of contaminants in the ecosystem. In the past few
decades, there has been a steady rise in the release of nitrate (NO 3-), a prominent water soluble
contaminant associated with the increasing use of nitrate based fertilizers. The first step in
environmental remediation is assessing the threat by accurate quantification of the contaminant.
19

The use of nanostructured enzymatic biosensors in environmental monitoring is fast emerging as
an important tool to provide valuable inputs on the presence of contaminants in the ecosystem. In
this study, we suggest the use of a highly sensitive, enzymatic biosensor capable of detecting and
quantifying minute concentrations of nitrate. The nitrate biosensor consists of a sensing element
in the form of nitrate reductase, a class of enzyme capable of reducing nitrate to nitrite
immobilized within a conductive polymer matrix responsible for generating a quantifiable
amperometric response. In our work, nanoarrays of co-immobilized nitrate reductase and
poly(3,4-alkylenedioxypyrrole) (PEDOT), a conducting polymer resistant to nuclueophilic attack
at the β-position were grown from a supramolecular surfactant stabilized matrix using a template
assisted electropolymerization route. The performance of the biosensor is a strong function of
electropolymerization conditions and the morphology of the PEDOT nanostructures. The
electropolymerized biosensor displays excellent specificity w.r.t other interfering ions as
evidenced from the initial rate kinetics. With a response time of a few seconds, limit of detection
(LOD) as low as 108 ppb and sensitivity of about 2.6 µA/µM cm2, the one-step
electropolymerized nanostructured nitrate biosensor developed in this study shows improved
performance compared to other electrochemical biosensors. As per our knowledge this is the first
attempt to grow single-step enzyme co-immobilized PEDOT nanostructured biosensor while still
retaining the upright nanowire structure. This combined with easy and fast fabrication technique
(polymerization time is just 300 seconds) opens up exciting opportunities for developing high
accuracy PEDOT based nanobiosensors for on-field testing of nitrate contaminants in the future.

20

REFERENCES

21

REFERENCES

1.

Ling, X.Y., D.N. Reinhoudt, and J. Huskens, From supramolecular chemistry to
nanotechnology: assembly of 3D nanostructures. Pure and Applied Chemistry, 2009.
81(12): p. 2225-2233.

2.

Goldemberg, J., Ethanol for a sustainable energy future. science, 2007. 315(5813): p.
808-810.

3.

Zhang, Y.-H.P., Reviving the carbohydrate economy via multi-product lignocellulose
biorefineries. Journal of industrial microbiology & biotechnology, 2008. 35(5): p. 367375.

4.

Luo, L., E. van der Voet, and G. Huppes, An energy analysis of ethanol from cellulosic
feedstock–Corn stover. Renewable and Sustainable Energy Reviews, 2009. 13(8): p.
2003-2011.

5.

Prasad, S., A. Singh, and H. Joshi, Ethanol as an alternative fuel from agricultural,
industrial and urban residues. Resources, Conservation and Recycling, 2007. 50(1): p. 139.

6.

Galbe, M. and G. Zacchi, A review of the production of ethanol from softwood. Applied
microbiology and biotechnology, 2002. 59(6): p. 618-628.

7.

Tu, M., R.P. Chandra, and J.N. Saddler, Evaluating the distribution of cellulases and the
recycling of free cellulases during the hydrolysis of lignocellulosic substrates.
Biotechnology progress, 2007. 23(2): p. 398-406.

8.

Silvestre, C., D. Duraccio, and S. Cimmino, Food packaging based on polymer
nanomaterials. Progress in Polymer Science, 2011. 36(12): p. 1766-1782.

9.

Yam, K.L., The Wiley encyclopedia of packaging technology2009, Hoboken, N.J.: John
Wiley & Sons.

10.

Laaperi, A., Disruptive factors in the OLED business ecosystem. Information Display,
2009. 9(09): p. 09.

22

Chapter 3: Supramolecular modification of graphene platelets for biocatalytic
saccharification of biomass
(This chapter is reproduced with permission from Journal of Molecular Catalysis B: Enzymatic 90 (2013): 76-86.)

3.0 Abstract
In this study, we show an effective way to immobilize cellulase enzyme on solid nanographene
supports using supramolecular assembly techniques. Using a bottom-up assembly approach we
show the hydrophilic stabilization of graphene supports using a combination of polyelectrolytes.
The inclusion of iron-oxide nanoparticles in the assembly allows facile magnetic separation
allowing recycle and reuse of the immobilized enzyme. Weak polyacrylic acid brushes act as
spacers between the magnetic graphene supports and cellulase. We show the effect of
dissociation of weak polyacid on the enzyme activity. Compared to free enzymes, as much as
80% of the activity is preserved. The resilience of the immobilized enzymes was tested by
repeated magnetic separation, recycle and reuse. As much as 55% of the activity was retained
after four cycles of reuse.

3. 1. Introduction
There has been a growing interest in graphene and graphene based nanomaterials ever since
the facile method for its isolation was first reported in 2004.[1] Because of its unique physical
and electronic properties, graphene, a 2D one atom thick allotrope of carbon, has been canvassed
as a potential replacement for silicon in the near future.[2] Besides the obvious applications in
electronics and semiconductor industry, graphene with its high surface area and large aspect
ratio, can provide ample opportunities to anchor a host of biomolecules on its surface. Recently
some groups used graphene oxide (GO) based carriers for loading biomacromolecules such as
23

enzymes and anti-cancer therapeutics followed by their targeted release[3-6]. Unlike its oxide,
the solubility of graphene in water is poor making its dispersion in aqueous medium difficult.
Hence, the complete biological potential has not been realized because of the hydrophobic nature
of pristine graphene. In the past, studies have suggested that the shape and degree of curvature of
the support used for enzyme immobilization can have a significant effect on enzyme activity[7,
8]. Davison et.al. [9] hypothesized that curved nanostructures especially concave structures can
provide ample opportunities for protein binding and help in enzyme stabilization. Soybean
peroxidase immobilization studies carried out by Asuri et. al. [10] on curved single walledcarbon nanotubes (SWNT) and flat 2-D graphite flakes indicated that deactivation constants for
2-D graphite supports tend to be higher as compared to curved SWNT supports. Conformational
changes induced in the structure of the enzyme as it binds to solid supports can inhibit its
bioactivity. Zhang and co-workers[6] reported lower enzymatic activity due to conformational
changes occurring during the immobilization of horseradish peroxidase on flat GO supports.
While shape and size of the supports along with the conformational changes as a result of
immobilization would continue to play an important role in enzymatic activities, in this work we
address both these concerns by making suitable modifications in the biocatalyst architecture.
Graphene nanoplatelets (GNP) are new generation carbon nanomaterials prepared by exfoliation
of natural graphite. Because of their planar morphology, they act as flat 2-D supports when used
for immobilization of biomolecules. Carbon nanostructures have been used in the past for
biomolecule immobilization because of their ability to stabilize biological structures at elevated
temperatures[11-14]. Ji and co-workers[15] showed that the lipase immobilized on multiwalled
carbon nanotubes show improved hydrolytic activity. It has been hypothesized that carbon
nanotubes because of their higher thermal conductivity ensure efficient heat transfer to the

24

immobilized enzyme. In carbon chemistry, graphene is the 2-D analog of carbon nanotubes and
should share similar heat conduction properties. Another type of 2-D nanomaterial is clay
platelets. However, the sorption and immobilization of biomolecules on clay depends on several
factors such as pH and type of cation present in the platelets making it difficult to isolate the
effect of shape of the carrier[16, 17].
Graphene nanoplatelet composed of a few layers of graphene sheets stacked together is one
of the most common types of nanostructured graphene.[18, 19]The surface of these platelets is
composed of sp2 hybridized pure graphene. Platelets with BET surface areas up to 900 m2/gm
can be produced by exfoliation. Such exfoliated cost-effective GNPs with a projected price of
$5/lb can be produced at Michigan State University using a process described elsewhere[20].
High surface area and low cost of production make graphene an ideal candidate for enzyme
immobilization. In the past, surface modification using wet chemistry techniques has been used
to address the issue of hydrophobicity of carbon nanostructures. These methods generally require
using strong inorganic acids and physical processes such as ultrasonication, and high shear
homogenization in presence of surface active reagents[21-24]. However, not all methods are
suited for biological modification. Use of toxic chemicals or harsh process conditions can instead
have a deleterious effect on the biological viability of carbon nanostructures.
Originally conceived by Iler[25] and further developed by Decher and coworkers[26, 27], the
layer-by-layer assembly (LbL) has become an important tool to produce nanoscopically
structured novel materials. Initially used for building polyelectrolyte multilayers, this versatile
technique is now also used to incorporate a plethora of nanoscopic, polymeric and bioactive
components within the film structure[28]. Using extremely simple apparatus such as beakers and
tweezers, the LbL technique allows us to prepare nanoscopic synergistic functional films by
25

combining various components within the same unit. The ability to blend together various
functional materials by creating supramolecular multilayered assemblies makes LbL a very good
example of ‘bottom-up’ approach. Smuleau et. al.[29] and Dotzauer et. al.[30] showed the
immobilization of enzymes and gold nanoparticles respectively on polyelectrolyte films
assembled on membrane filters. Smuleau and co-workers reported that the interaction between
the oppositely charged polyelectrolyte layers and the protein molecules enables a 25 fold
increase in immobilization. Very small conformation changes were noticed in the biomolecule
configuration after the polyelectrolyte modification. Dotzauer and co-workers showed that gold
nanoparticles immobilized on porous supports using the LbL approach can efficiently reduce
almost 99% of the 4-nitrophenol. Wang and Caruso[31] applied the electrostatic LbL assembly
to 3D macroporous membranes for the immobilization of catalase. The polyelectrolyte
multilayers accomplish two outcomes. Firstly they allow attachment of the catalase enzyme on
the membrane. Secondly they also prevent the leakage of the enzymes facilitating improved
biocatalytic performance. Siqueira Jr. et. al.[32] review the various strategies used for
immobilization of biomolecules on LbL and Langmuir-Blodgett nanofilms. Lee and co-workers
at Michigan State University incorporated proteins, dendrimers, nanoparticles, lipids, cells, and
metals onto or into the LbL layers for a variety of applications such as biosensing, biocatalysis,
and biomedical applications[33-37]. The molecular-level design of interfacial bonding at the
interface via self-assembly is the key to fabricating stable and functional interface[38, 39]. This
affects the adhesion and functions of the resulting nanostructured interfaces.
Cellulose, a major component of the lignocellulosic biomass, is hydrolyzed by cellulase
to yield reducing sugars[40]. Cellulase, a class of saccharifying enzymes, has gained attention in
recent years due to the significant role it plays in the production of ethanol from cellulosic

26

substrates. Cellulase is not just one enzyme system According to the widely accepted view,
efficient saccharification of cellulosic biomass is achieved by the cooperative action of a number
of enzymes including endo-glucanase and exo-glucanase[41, 42]. The endo-exo synergism
disrupts the cellulosic matrix to ultimately release reducing sugars[41]. The cost of cellulase
accounts for a significant portion of the hydrolysis costs, which according to some estimates, is
as high as 50%[43, 44]. In recent years, a combination of genetic engineering and cheaper
downstream processes has helped bring about several improvements in enzyme production as
well as cellulase performance[43, 45]. However, further reduction in the cost of cellulase
enzymes by genetically modifying the cellulolytic microorganisms would be challenging[43].
Cellulase recovery and reuse is another potential approach that can be used to offset the high
costs associated with enzyme use. Unfortunately, during a typical hydrolysis reaction, cellulases
tend to get distributed over two heterogeneous phases: the solid substrate and the liquid
supernatant thus making recovery of the enzymes difficult[43, 46].
Immobilization of cellulase on solid supports has been suggested as a viable alternative[47].
Immobilization of enzymes on supports helps to improve the overall operational stability of
enzymes by increasing their resilience towards changes in external factors like temperature and
pH[48-50].

Moreover, it prevents cellulase redistribution over two heterogeneous phases

facilitating easy separation, providing a facile route to recycle and reuse the enzyme over
multiple cycles. In this work, the presence of iron oxide nanoparticles on graphene supports
allows for the magnetic separation of the immobilized cellulase. The recovered cellulase is then
reused over multiple cycles thus reducing the overall cost of using fresh enzymes. With cellulase
poised to capture the second largest segment in the enzyme market [51], improved

27

immobilization strategies with a recovery mechanism to save costs are necessary to ensure its
optimum use.
In our recent graphene studies, we used different cationic and anionic polyelectrolytes like
poly(diallydimethylammonium chloride) (PDAC) and sulfated poly(styrene) (SPS) respectively
to disperse GNPs in aqueous medium [18, 52, 53]. The nanoplatelets develop cationic or anionic
surface charge depending on the type of polyelectrolyte used. Electrostatic assembly of
oppositely charged nanoplatelets is possible using the LbL technique. In this work, we utilize a
similar approach to prepare novel magnetoresponsive biocompatible graphene supports for
cellulase immobilization. These supports will be hence forth called magnetic nanoparticle
incorporated graphene nanoplatelets. (MNP-GNP). Maghemite-magnetite decoration followed
by cellulase immobilization is carried out by using a combination of quenched and annealed
polyelectrolytes. Strong polyelectrolytes mentioned above are used for efficient electrostatic
binding of anionic iron oxide nanoparticles (8-10 nm in size). Polyacrylic acid, a weak
polyelectrolyte, makes the surface of MNP-GNP biocompatible. Cellulase is then attached to the
supports using covalent chemistry. Weak polyacid brushes such as PAA are pH and heat
sensitive[54-56]. The swelling behavior of pH, heat tunable polyacrylic acid brushes gives us an
opportunity to control cellulase mobility. The presence of nanoparticles and polyelectrolytes
between the graphene support and the enzyme provides a ‘cushioning effect’. Increased freedom
of movement, enhanced enzyme mobility and reduced steric hindrance due to the underlying
support help fine-tune the microenvironment of the immobilized enzymes to closely resemble
that of free enzymes[57]. Thus, the limitations imposed by the 2D morphology of graphene
supports can be effectively addressed by making suitable changes in the design of the
immobilized biocatalyst.

28

3.2. Experimental procedures
3.2.1 Materials
Poly(diallydimethylammonium chloride) (PDAC), sulfated poly(styrene) (SPS), and
polyacrylic acid (PAA) sodium salt (35 wt% solution in water) were purchased from Sigma
Aldrich. Average molecular weights of PDAC, SPS and PAA were ~100,000-200,000, 70,000,
and

15,000

respectively.

Carboxymethyl

cellulose

(CMC)

sodium

salt,

2-(N-

morpholino)ethanesulfonic acid (MES) and citric acid monohydrate were purchased from Fisher
Scientific

Co. Avicel PH- 101, 50 μm in size was obtained from Fluka. 1-Ethyl-3-(3′-

dimethylaminopropyl) carbodiimide, Hydrochloride (EDAC, HCl) was purchased from EMD
Chemicals. Accellerase-1000, a commercial cellulase enzyme was a kind gift from Genencor
Division, Danisco US Inc., Rochester, NY, USA. Exfoliated graphene nanoplatelets (GNP)
(BET surface are 327 m2/gm and size ≤ 1µm) were provided by Dr. Drzal’s research group at
MSU. All aqueous solutions were prepared using deionized (DI) water (>18.1 M) supplied by a
Barnstead Nanopure Diamond-UV purification unit equipped with a UV source and final 0.2 m
filter. Unless specified otherwise, all procedures were carried out at room temperature.

3.2.2 Preparation of polyelectrolyte solutions
Aqueous polyelectrolyte solutions were prepared containing either 20 mM PDAC or 2 mg/ml
SPS. Sodium chloride concentration was adjusted to 0.1 M. The concentration of PAA solution
was 1 mg/ml with no addition of salt. The solutions were filtered through 0.22 µm celluloseacetate membrane-filter system from Nalgene to remove particulates. The aqueous
polyelectrolyte solutions were used without pH adjustment.

29

3.2.3 Preparation of polyelectrolyte-multilayered graphene scaffolds
Around 200 mg of GNP were added to 50 ml of SPS solution. The resultant suspension was
stirred for 30 minutes. This was followed by ultrasonication for 90 minutes. The SPS modified
graphenic suspension was then centrifuged at 4000 rpm followed by three washes of deionized
water to remove the excess SPS. To deposit a PDAC layer, the SPS-coated GNP is added to 50
ml of PDAC. A similar procedure as described above is repeated. In order to achieve uniform
surface coverage of polyelectrolytes, we deposited in all four layers of SPS and PDAC (i.e. two
bilayers). Positively charged PDAC layer occupies the uppermost layer in the quad-layer
assembly.

3.2.4 Alkaline co-precipitation of maghemite-magnetite nanoparticles (8-10 nm)
Detailed description of preparation of aqueous suspension of magnetic nanoparticles is
available elsewhere[58, 59]. Briefly speaking, Fe 3+ and Fe 2+ ions are mixed in the molar ratio of
2:1. 5 ml of ammonium hydroxide is added drop-wise. The mixture is heated to 800C under
nitrogen purging. 1 ml of citric acid (2.38 M) is added to the above mixture and the temperature
is raised to 950C. Heating is continued for 90 minutes. The colloidal solution is then separated by
centrifugation (9000 rpm) and washed thrice with a mixture of acetone and water to get a stable
dispersion. The nanoparticles are then dispersed in 30 ml of deionized water and stored at room
temperature.

3.2.5 Maghemite-Magnetite decoration of polyelectrolyte-modified graphene scaffolds
The maghemite-magnetite nanoparticles solution is diluted to 0.53 mg/ml by adding DI
water. Around 50 ml of this solution is combined with polyelectrolyte-coated graphene as
synthesized above and sonicated for 90 minutes. The maghemite-magnetite decorated graphene

30

is then centrifuged and washed with copious quantity of DI water to remove the unattached
nanoparticles. With each successive wash, the color of the supernatant changes from pale
brownish-yellow to that of clear liquid. Finally the precipitate is dried in an oven at 65 C for two
days and stored in a vacuum desiccator for future use.

3.2.6 Immobilization of cellulase on maghemite-magnetite graphene scaffolds
Depending on the substrate used for hydrolysis, a predetermined quantity of maghemitemagnetite decorated graphene is added to 1 mg/ml PAA solution. Ultrasonication for 60 minutes
helps in efficient dispersion of MNP-GNP in PAA. The PAA grafted graphene scaffolds are then
centrifuged and washed twice with copious quantities of DI water. Terminal carboxylic acid
groups of PAA are activated by immersing the PAA grafted MNP-GNP in 0.1 M/0.5 M
MES/NaCl buffer containing 100 mg. of EDAC, HCl. The pH of the buffer solution prior to
addition of EDAC, HCl was adjusted to 5.5. The carboxylic group activation continues for 1
hour on a tube rotator at room temperature. The EDAC activated PAA modified MNP-GNP is
centrifuged at 4000 rpm and washed twice with DI water to remove unattached EDAC, HCl.
Accelerase-1000, a commercial cellulase enzyme containing both endo and exo glucanase is
centrifuged at 2000 rpm. The supernatant solution is separated, diluted to 1.375 mg/ml and used
for further experiments. 2 ml of this diluted enzyme solution is added to EDAC activated
graphene supports as prepared above. Incubation continues for 15 hours on a tube rotator at 40C.
After the incubation time, the immobilized enzymes are centrifuged at 4000 rpm, washed thrice
with DI water. The washings are pooled together for protein estimation. The PAA modified
immobilized enzymes (now called IMC-PAA) are then suspended in appropriate buffer solution
of desired pH. A separate set of immobilized enzymes are prepared by following the exact

31

procedure except without using PAA and EDAC, HCl. These immobilized enzymes will be
henceforth called IMC.

3.2.7 Characterization of polyelectrolyte coated graphene supports
The zeta potential of graphene particles after polyelectrolyte and enzyme modification was
measured by ZetaPALS instrument (Brookhaven Instruments Corp., Holtsville, NY). To
determine the ζ-potential, 100 µl of the sample was added to 1.4 ml 1 mM KCl solution. The
KCl solution was filtered through 0.22 µm cellulose acetate membrane filter prior to use. Phase
analysis light scattering studies typically determine the electrophoretic mobility of charged
particles from measurable parameters such as velocity of the particles and the magnitude of the
applied electric field. This data is then fitted to the Smoluchowski equation by the ZetaPALS
software to obtain the zeta potential value.

3.2.8 Characterization of GNPs and MNP-GNPs
The scanning electron microscopy (SEM) investigation for high-resolution imaging was
carried out using a JEOL 7500F with field emission. Transmission electron microscopy (TEM)
studies were carried out using a JEOL 100CX and JEOL 2200FS. Both SEM and TEM analyses
help us to determine the morphology of graphene before and after nanoparticle decoration. To
determine the extent of nanoparticles doping, MNP-GNP samples were digested in HCl/HNO3
(3:1 volume ratio) mixture and the samples were analyzed using inductively couple plasma
optical emission spectroscopy (ICP-OES).

3.2.9 Protein assay
Protein concentration of the enzyme solution before and after immobilization is determined
using the Bradford method[60]. Protein loaded on MNP-GNPs was estimated by doing a simple
32

mass balance before and after immobilization. The protein loading was found to be 2.5 ± 0.75
mg/gm MNP-GNP. Covalent immobilization helps prevent enzyme leaching. However, a very
small quantity of protein may still desorb from the immobilized enzyme if subjected to repeated
use.

The protein desorbed during the recyclability studies is quantified using the micro-

bicinchoninic acid (mBCA) assay, a technique typically used for measuring small quantities of
protein (0.5-20µg/ml). Since glucose can interfere with mBCA results, two separate sets of
experiments, one for measuring glucose and other for measuring desorbed protein were
performed. For the later set of experiments, no substrate was added.

3.2.10 Measurement of cellulase activities
Cellulase activities are traditionally measured by quantifying the end-products of the
hydrolysis reaction[61]. These end-products are in the form of reducing sugars. For CMC
experiments, 10 mg. MNP-GNPs were used. For avicel based experiments, 5 mg. were used. The
amount of protein immobilized on MNP-GNPs was estimated as described in the previous
section. Free enzyme solution having the same protein content as the immobilized enzymes was
prepared by diluting with appropriate buffer solution. The biocatalytic activity of free and
immobilized enzymes was measured under similar reaction conditions using 2% w/v CMC and
avicel. Hydrolysis was run for 90 minutes. Samples were subjected to 5 minutes of boiling to
stop the reaction. The amount of generated glucose was measured using high-performance liquid
chromatography (HPLC). One activity unit is defined as the amount of cellulase that catalyzes
CMC or avicel to release 1 µmole glucose/min/mg. of protein under specified hydrolysis
conditions. We define two terms here, relative activity and mimicking ability of immobilized
enzymes. Thus, while relative activity normalizes the enzyme activity on the basis of specific
hydrolysis parameters most frequently used in cellulase assay procedures (50 C and pH 4.8),
33

mimicking ability is a measure of the biocatalytic potential of immobilized enzymes when
compared to free enzymes at the same pH and temperature.

..........(1)

..........(2)
For recyclability studies, 50 mg. of MNP-GNP were used. Each hydrolysis cycle was run for 60
minutes at a temperature of 50 C and pH 5.2. The immobilized enzymes were magnetically
separated after every cycle. Four hydrolysis cycles were repeated using the same set of
conditions, each time adding a fresh batch of substrate.

34

3.3. Results and Discussions

Figure 3.1 Schematic illustration of the design of the proposed biocatalyst: Assembly of
polyelectrolytes, nanoparticles and cellulase on graphene nanoplatelets. (Figure not drawn to
scale)
Figure 3.1 illustrates a schematic diagram showing the ‘bottom-up’ assembly of quenched
polyelectrolytes, iron-oxide nanoparticles, annealed polyelectrolyte and finally cellulase. As seen
from the figure, because of the presence of polyelectrolytes and nanoparticles, we anticipate that
most of the cellulase immobilization occurs on the outer surface of modified GNPs as opposed to
between the individual nanoplatelets The proposed design is particularly attractive because it
helps minimize internal diffusion limitation which otherwise is a major impediment in enzyme
biocatalysis.
Based on key steps involved in the preparation of the biocatalyst, we broadly discuss the
theory and results under three headings: Maghemite-magnetite decoration of GNPs,
chemisorption of PAA brushes, and lastly its proposed effect on enzyme activity.

35

3.3.1 Maghemite-Magnetite decoration of graphene nanoplatelets
One of the very first steps in the design of graphene based biocatalyst is to improve its long
term stability in aqueous medium. Stable aqueous dispersion can be obtained by subjecting
graphene to polyelectrolyte modification, as shown in our previous work [18, 52, 53]. Table 3.1
shows the cyclic nature of charge overcompensation as each bilayer of SPS and PDAC is
sequentially adsorbed on the surface of GNPs. Table 1 also shows the effect of adsorption of
magnetic nanoparticles (zeta potential turns negative) and PAA (degree of negative charge
increases further) on the polyelectrolyte modified graphene supports.
Table 3.1 ζ-potential (mV) of modified GNP dispersions. The last component in each line
represents the terminal (topmost) layer in the multilayer assembly. Error bars represent the
standard deviation of 3 replicate samples.
Type of terminal layer
Zeta potential
(mV)
GNP-SPS
-51.50 ± 0.75
GNP-SPS-PDAC
43.25 ± 1.23
GNP-SPS-PDAC-SPS
-49.25 ± 1.22
GNP-(SPS-PDAC)2
46.91 ± 0.95
GNP-(SPS-PDAC)2-MNP
-26.36 ± 2.92
GNP-(SPS-PDAC)2-MNP-PAA
-41.78 ± 3.15
In case of pristine GNPs, the Vander-Waals force of attraction primarily caused by polarized
π-electrons along with the hydrophobic interactions between individual graphene sheets cause
the nanoplatelets to aggregate, rendering them insoluble in water[62]. High molecular weight
polyions (in this case SPS and PDAC) possess a remarkable ability to cover substrate
irregularities through multipoint attachment[63]. As compared to small molecules and charged
colloids, they provide a more consistent and uniform coating[64]. Depending on the functional
groups, different polyelectrolytes can disperse GNPs upto varying degree of stability[18]. The
ability of negatively charged SPS, the first polyelectrolyte in the quad-layer assembly, to
solubilize GNPs can be explained by the concept of ‘polymer wrapping’. The concept of
36

‘polymer-wrapping’ followed by LbL assembly has been used in the past to add functionality to
different types of carbon nanostructures including carbon nanotubes[65, 66]. SPS is an
amphiphile. While its backbone is highly hydrophobic, the sulfonate groups impart hydrophilic
and anionic character to the molecule. Limited scale interaction calculations reveal that the
robust adsorption of SPS on GNPs is primarily because of edge-to-face interactions between the
graphene surface and aromatic rings of SPS[67]. Thus, while the hydrophobic areas of the GNPs
are screened by the hydrophobic part of SPS, the anionic sulfonate groups project themselves in
the aqueous medium and stabilize the suspension through mutual repulsion. The polyelectrolyte
multilayer formation is a thermodynamic driven process in which both electrostatic as well as
hydrophobic interactions play an important role. The incoming polyelectrolyte solution also
produces a change in the ionic environment as well as the hydration shell around both the
polyelectrolytes[68]. A simple free energy model developed by Kotov[68] to explain the
polyelectrolyte adsorption process accounts for all individual contributions mentioned above.
The presence of two bilayers of oppositely charged polyelectrolytes serves two purposes.
Firstly, it gives rise to tightly coiled intertwined polyelectrolyte conformations around the GNPs.
The polyelectrolytes stabilize the GNPs promoting their effective dispersion in aqueous medium.
It has been suggested that multilayers of strong polyelectrolytes (in this case SPS and PDAC)
can disperse colloidal particles more effectively as opposed to weak polyelectrolytes because of
their stability over wide range of conditions[69]. Secondly, they also serve as precursor films for
the subsequent adsorption of iron oxide nanoparticles. It is a widely accepted view that the
presence of precursor films help in eliminating substrate effects generating surfaces with almost
uniform configurations[63]. In our case, we observed that precursor film composed of two
bilayers was sufficient to enable reasonably uniform adsorption of iron-oxide nanoparticles. The

37

incorporation of iron oxide nanoparticles on the surface of graphene nanoplatelets makes these
supports magnetic. Thus, rather than using the cumbersome energy intensive process of
centrifugation, facile separation of these supports can be achieved by using small bar/disc
magnets. Cellulase immobilization on magnetic supports offers attractive opportunities to lower
the enzyme requirement of the process by recovery and reuse over multiple cycles.
Alkaline co-precipitation of maghemite-magnetite nanoparticles is a long established method
for preparation of monodisperse iron oxide nanoparticles. According to Massart,[59]
surfactantless iron oxide colloids may aggregate due to neutralization of charges when used
between pH 5 to pH 9. α-hydroxy acids such as tartaric, oleic and gluconic acid have been used
in the past to stabilize the colloids[70]. In our case, we used citric acid, another α-hydroxy acid
known for bearing tridentate carboxylate ligands. The presence of three negatively charged
acidic groups not only helps in stabilizing the magnetic nanoparticles over a wide range of pH
but also provides sufficient interaction sites when the citric acid capped iron oxide nanoparticles
are adsorbed on the positively charged PDAC layer. The nanoparticle decoration as estimated by
ICP-OES is about 47 mg iron/gm of MNP-GNP. It has been shown that the isoelectric point of
citric acid coated magnetic nanoparticles is lesser than pH 3 [71] which is lower than the lowerbound of the pH range we use for our enzyme activity tests. Thus, the chances of iron oxide
nanoparticles reversing their surface charge and getting desorbed from the surface of graphene
are minimal. The SEM and TEM micrographs (Figure 3.2) show the adsorption of iron oxide
nanoparticles roughly of size 8-10 nm on the surface of polyelectrolyte-coated GNPs.

38

Figure 3.2 SEM micrographs of (a) GNP, (b) MNP-GNP (low magnification), and (c) MNPGNP (high magnification): Average size of iron oxide nanoparticles around 8-10 nm. (d) TEM
micrograph of GNP. The inset shows high magnification image of a selected portion of the same
platelet. Scale bar for the inset is 100 nm. TEM micrographs of (e) MNP-GNP (low
magnification) and (f) TEM micrograph of MNP-GNP (high magnification): Almost uniform
coating of iron-oxide nanoparticles (8-10 nm) is observed on the surface of the graphene
nanoplatelet.
39

The size of the iron-oxide nanoparticles has a significant bearing on the superparamagnetic
property with the saturation magnetization decreasing with increasing particle size[72]. Besides
our current strategy of using polyelectrolyte precursor films to fixate nanoparticles, another
promising approach suggested in literature is embedding metal precursors in the graphene
matrix[19]. These precursors act as ‘seeds’ for the subsequent growth of nanoparticles. The
‘seeding’ approach has been shown to produce dense uniform films of nanoparticles. However,
with higher loading of nanoparticles, there are greater chances of aggregation[73]. Also, the
nanoparticles grown from precursors show a high degree of polydispersity. Though methods
have been suggested to monitor the growth of nanoparticles and ensure the monodispersity,[19,
73] they usually require careful process control, or additional steps such as developing a method
to embed the precursor in the support matrix followed by its chemical reduction. The layer-bylayer method, on the other hand, involves separate preparation of nanoparticles of desired size to
suit our needs. The nanoparticles can then be adsorbed on the surface of graphene by merely
contacting the supports with the nanoparticle solution. The incorporation of iron-oxide
nanoparticles on GNPs provides these supports with a new functionality. The magnetoresponsive
behavior of GNPs after nanoparticle decoration is evident from Figure 3.3.

Figure 3.3 Effect of external magnetic field on graphene nanoplatelets (a) before iron oxide
nanoparticle decoration: No change observed (b) after iron-oxide nanoparticle decoration: GNPs
driven towards the magnetic source.
40

3.3.2 Chemisorption of polyacrylic acid brushes on MNP-GNP
Maghemite-magnetite decorated graphene supports offer an attractive opportunity to
immobilize a number of bioactive components. Biological molecules have often shown subdued
activity when subjected to direct immobilization on supports[74]. So in this case, we use
polyacrylic acid, a well-known biocompatible material, to serve as a bridge between the cellulase
enzyme and the MNP-GNPs. Many studies have shown that PAA can be used as an effective
biomaterial in several applications such as muco-adhesion, drug delivery, etc.[75, 76]. PAA also
serves as a good dispersing agent. Going back to Table 1, we observe that the use of PAA helps
in increasing the magnitude of negative zeta potential, thus improving stability of the dispersion.
PAA has multiple hydrophilic carboxylic groups, some of which are used up when PAA interacts
with the iron oxide layer of MNP-GNPs while the rest can be utilized for cellulase
immobilization. Sugama et. al. [77] has proposed two possible mechanisms for chemisorption of
PAA on metal oxide surfaces. One of the proposed mechanisms suggests that the acid base
interaction between the –COOH groups of PAA and the polar groups –OH of the oxide allow for
strong chemisorption of PAA on the metal oxide surface. The other mechanism involves
formation of a salt bridge between the carboxylic groups of PAA and the free metal ions of the
metal oxide. The formation of a hydrophilic PAA layer on the surface of MNP-GNP helps to
improve its dispersibility in aqueous medium. As mentioned earlier, the remaining –COOH
groups are then utilized for cellulase binding. EDAC- HCl, a commonly used zero-length spacer
molecule, helps to covalently bind the amine groups of cellulase to the –COOH groups of PAA.
Besides being a biocompatible material capable of anchoring proteins through covalent
linkage, PAA is also a weak polyelectrolyte. Unlike strong polyelectrolytes, which regardless of
the pH, dissociate completely when in solution, weak polyelectrolytes are heavily influenced by

41

salt content and acidity/alkalinity of the contacting solution[78]. In order to eliminate one of the
interfering factors, no salt was added while preparing the PAA solution. When the PAA is
chemisorbed on the surface of MNP-GNP, it gives rise to coiled conformations with some of the
segments stretching out into the solution because of electrostatic repulsion. Studies have shown
that the benign nature of the ‘brush-like’ polyelectrolyte assembly helps in preserving most of
the activity of the enzymes or other bioactive molecules immobilized within the assembly[79,
80].
There are many examples in literature that focus on ‘smart’ polyelectrolyte-protein
bioconjugates[81-84]. Wang and Caruso[85] used PAA to infiltrate into protein loaded silica
mesoporous spheres. PAA binds to most proteins electrostatically and covalently and thus acts as
a ‘bridging agent’.In some cases, even a small change in the environmental condition can trigger
a large response in the structure and properties of the bioconjugate. Imanishi and Ito[82] used
glucose oxidase immobilized weak PAA brushes to build a ‘chemical valve’ that would permit
the transport of insulin under low pH condition.
An important criterion in case of weak polyelectrolyte brushes is the pKa value. Based on
values available in literature, the pKa value of polyacrylic acid brushes occurs between 4.24.8[86-88]. An important fact worth noting at this juncture is that the pH optimum of cellulase is
also in that range, the relevance of which should become clear later. It has been wellestablished[56] that when the pH < pKa, the protonation of weak polyelectrolyte brushes leads to
collapse of the chain segments. When the pH > pKa, more and more –COOH groups become
charged producing swollen brushes with chain segments extending all the way into the
contacting buffer solution. This pH-induced swelling of the PAA brushes can have a significant
bearing on the enzyme activity. Similar examples of pH-responding polymer brushes for

42

biological applications can be found in literature[89-91]. While Zhou et. al. [91] discuss
reversible immobilization of a protein Cytochrome c made possible because of the pH induced
protonation-deprotonation of PAA brushes, Tam and co-workers[89] demonstrated the
successful use of a modified Poly(4-vinyl pyridine), (P4VP) polybase brush for bioelectronic
applications. The electrochemical activity is enhanced at low pH when the brushes are in a
swollen state. A collapsed P4VP brush at higher pH restricts the flow of electrons between the
redox species and the conductive support thus reducing the electrochemical activity. Likewise,
neutron reflectometry (NR) experiments have also confirmed that certain polymer brushes such
PAA brushes show a remarkable swelling behavior when subjected to moderate heat
treatment[92].

3.3 Effect of PAA brushes on activity of immobilized enzymes
Figure 3.4 illustrates how environmental conditions such as pH and temperature affect the
PAA brushes.

43

Figure 3.4 Effect of swollen PAA brushes on cellulase mobility. (a) Higher steric hindrance due
to collapsed PAA brushes. (b) Higher cellulase mobility and increased substrate-enzyme
interaction due to swollen PAA brushes. (Figure not drawn to scale)
Before examining how these conditions typically affect the immobilized enzyme activity, it
is important to note that cellulases consist of a complex mixture of various enzymes and this
includes endoglucanase and exoglucanse. Endo- and exo glucanases usually produce a
synergistic effect making it difficult to demarcate the contribution of each enzyme. However,
some studies claim that certain substrates because of their internal structure can help isolate endo
and exoglucanse activities from each other[93]. Carboxymethyl cellulose (CMC), a soluble form
of cellulose, is a model substrate used to evaluate the cleavage of internal glycosidic bonds, an
action which is typical endoglucanase. Henceforth, activity determined using CMC as substrate
would be referred to as CMCase activity. Exoglucanase activity, a measure of the efficiency with
which the terminal glycosidic bonds are cleaved, can be evaluated using avicel, a crystalline
form of cellulose. For the enzyme recovery experiments, we continued to use avicel since it
44

resembles the real biomass samples more closely as compared to CMC. Henceforth, activity
determined by using avicel as the substrate would be referred to as avicelase activity.

3.3.3.1 Effect of pH on enzyme activity
Data obtained from hydrolysis of avicel and CMC by three sets of enzymes (FE, IMC-PAA
and IMC) is shown in Figures 3.5. Relative enzyme activity was evaluated at six different values
of pH starting with a pH of 3.0.. Subsequent points are 0.7 pH units apart. These pH values were
selected keeping in mind the pKa value of the PAA brushes (4.2-4.8). pH 3.0 and 6.5 are far off
from the pKa value of PAA. They form the two extremities of the data set. Our data (not
included here) showed that the trend exhibited by the enzymes at these two data-points continues
likewise beyond these points. Data obtained at pH 3.7, 4.4 5.1 and 5.8 is critical in understanding
the effect of PAA brushes on immobilized cellulase activity. pH 3.7 is slightly below the lower
bound of the pKa value. At this pH, the brushes are protonated and are in a collapsed state. pH
4.4 is in the transition range as the brushes undergo a transformation from a collapsed state to
swollen state. pH 5.1 is slightly higher than the upper bound of the pKa value. At this pH, the
brushes are swollen and extend all the way into the buffer solution. At a pH of 5.8, other
concerns such as higher agglomeration due to lower zeta potential and lower enzymatic activity,
start dominating. Two different types of buffer solution were used. 0.05 M citrate buffer was
used in experiments where the pH was adjusted to 3 and 3.7 and 4.4. 0.05 M MES buffer
solution was used for adjusting the pH to 5.1, 5.8 and 6.5.

45

(a)

(b)

Figure 3.5 Relative activity of free and immobilized enzymes as a function of pH (a) Avicelase
activity (b) CMCase activity. Conditions: 2%w/v avicel/CMC, Hydrolysis time: 90 minutes,
Temperature 50 0C. Error bar represent standard deviation of three replicate samples.
Refer Figure 3.5(a). The free enzyme activity is about 20% at a pH of 3.0 reaches 75% at a
pH of 3.7. At a pH 4.4 and 5.1 it is almost 100% and finally tapers off to 65% and 35% at pH of
5.8 and 6.5 respectively. IMC follows a similar trend, though the activity is much lower at the
same pH. Enzyme activity reaches an optimum value of around 57% around pH 3.7 and stays in
a similar range upto pH 5.1. On the other hand, in case of IMC-PAA, there is a steep rise from
22% to almost 80% as the pH is raised from 3.7 to 5.1. The endo-glucanase data obtained by
substituting avicel with CMC (Figure 5(b)) mirrors the same trend. IMC-PAA activity increases
from 52% to 78% in the pH range 3.7-5.1. On the other hand, the avicelase and CMCase
activitites for IMC remain low (upto 62% at the most) in the pH interval 3.7-5.1.
We attribute the sudden spike in enzyme activity from pH 3.7 to 5.1 as fallout of the swelling
behavior exhibited by the PAA brushes. At lower pH, the brushes are in collapsed state pulling
the enzymes closer to the graphene support. The steric hindrance caused by the proximity of the
enzyme to the underlying support could explain the reduced enzyme activity. As the pH is raised
above the pKa value, the brushes elongate and push the enzymes away from the graphene
support. Greater interaction between the catalytic sites of the enzyme and the substrate enhances
46

the activity. This proposed theory is further supported by the IMC data. Without the PAA
brushes, both avicelase as well as CMCase activities show lower activity in the pH range 3.7-5.1.
We carried out a zeta potential analysis (please refer Table 3.2) of both IMC-PAA and IMC
over the entire pH range starting 3.0 to 6.5. Zeta potential being an indication of the stability of
colloidal particles can provide valuable insight into the behavior of enzyme modified particles at
different pH conditions. Table 3.2 clearly shows how the PAA modification impacts the charge
structure of the immobilized enzymes. The difference in charge structure can influence the
microenvironment of the immobilized enzymes producing variations in the enzyme activity. For
IMC, the particles display a positive charge at lower pH (3.0 and 3.7). The zeta potential is
negative at other pH values. This implies that the isoelectric point for IMC lies somewhere
between 3.7 and 4.4. On the other hand, IMC-PAA shows a negative zeta potential over the
corresponding pH range. An interesting observation can be made from this analysis. The zeta
potential data for IMC shows that the magnitude of surface charge for this set of immobilized
enzymes is low. In other words, IMC enzymes are not very stable and must be agglomerating
during the hydrolysis process thus reducing their effectiveness. On the other hand, IMC-PAA
becomes much more stable as the pH is raised from 3.0 to 5.1. At 4.4 and 5.1, it shows identical
surface charge of almost similar magnitude within the margin of error. Thereafter at pH 5.8 and
6.5, the magnitude of zeta potential reduces, thereby reducing its stability and hence its
effectiveness.

47

Table 3.2 ζ-potential (mV) of IMC and IMC-PAA as a function of pH. Error bars represent the
standard deviation of 3 replicate samples.
pH
IMC
IMC-PAA
3.0
7.23 ± 0.63
-19.28 ± 1.38
3.7
7.51 ± 0.79
-25.61 ± 2.27
4.4
-11.85 ± 0.54
-31.72 ± 2.32
5.1
-13.18 ± 0.57
-29.13 ± 1.25
5.8
-10.29 ± 0.95
-18.97 ± 0.98
6.5
-10.78 ± 0.69
-17.94 ± 1.23
Depending on the source from which it is derived and the ingredients that make up the
enzyme broth, the pH optimum for cellulase ranges from 4.2-5.0 with most studies using the
value 4.8 for experimental purposes.

As mentioned before, the choice of the weak

polyelectrolyte and the conditions under which it is used in the immobilization process are very
important. PAA has a pKa value which almost coincides with the optimum pH for free enzymes.
Consider poly(methacrylic acid), another example of a weak polyelectrolyte. AFM studies haves
confirmed that densely grafted poly(methacrylic) acid brushes show a remarkable swelling
behavior at a pH of about 9[94]. Swelling starts at around 7.0 and continues till 10.5. The
biocatalytic potential of cellulase at this pH is highly doubtful rendering the whole exercise of
using pH tunable annealed brushes ineffective.

3.3.3.2 Effect of temperature on cellulase activity

48

(a)

(b)

Figure 3.6 Relative activity of free and immobilized enzymes as a function of temperature (a)
Avicelase activity (b) CMCase activity. Conditions: (a)2 %w/v avicel (b)2%w/v CMC,
Hydrolysis time: 90 minutes, solution pH: 5.1. Error bars represent standard deviation of three
replicate samples.
Data for three systems: FE, IMC-PAA and IMC is presented in Figure 3.6(a) and (b). Our
results show that immobilized enzymes with PAA brushes closely follow the trend shown by free
enzymes. It has been shown previously that PAA adopt a swollen conformation at higher
temperatures[95, 96]. In another example involving formation of a mixed hydrogel of poly(vinyl)
alcohol (PVA) and PAA, Lee and co-workers[97] have argued that at higher temperature the
dissociation of hydrogen bonding between the –OH group of (PVA) and –COOH of PAA
produces excess carboxylate ions which can lead to swollen configuration. We hypothesize that
in our case as the temperature is raised from 400 C through 700C, the PAA brushes continue to
undergo incremental swelling as a result of interactions between the –OH groups of the ironoxide and –COOH groups of PAA. We believe that just like pH variation, increase in
temperature also facilitates formation of swollen brushes ensuring greater mobility and better
substrate-enzyme interaction. Thus, IMC-PAA can closely mimic the free enzymes. At 600C,
avicelase activity for IMC-PAA reaches its optimum value. Likewise, the CMCase activity for
IMC-PAA reaches its optimum of 78% at 500C. On the other hand, IMC show subdued avicelase
49

as well as CMCase activity (57% or less) over the entire temperature range. Thus the swollen
conformation of PAA brushes provided a favorable micro-environment for cellulase. After
reaching a maximum at 600C for avicel and 500C for CMC, cellulase activity showed a decline.
However this decrease in cellulase activity also corresponds to decrease in activity of free
enzymes as seen from Figure 3.6 and could be attributed because of partial deactivation of the
active centers at higher temperatures.
3.3.3.3 Mimicking ability of Immobilized enzymes

(a)

(b)

Figure 3.7 Mimicking ability of immobilized enzymes as a function of (a) Avicelase activity at
variable pH (b) CMCase activity at variable pH (c) Avicelase activity at variable temperature (d)
CMCase activity at variable temperature. Conditions: (a) 2%w/v avicel (b) 2%w/v CMC,
Hydrolysis time: 90 minutes, Temperature 50 0C. Data was collected at pH 3.0, 3.7, 4.4 ,5.1, 5.8
and 6.5. (c) 2 %w/v avicel (d) 2%w/v CMC, Hydrolysis time: 90 minutes, solution pH: 5.1.
Data collected at 400C, 500C, 600C and 700C. Error bars represent standard deviation of three
replicate samples.

50

Figure 3.7 (cont'd)

(c)

(d)

The mimicking ability of immobilized enzymes (IMC-PAA and IMC) is shown in Figure 3.7.
The superiority of IMC-PAA over IMC is evident in this figure. In Figure 3.7(a) and (b) we see a
marked improvement in the mimicking ability of IMC-PAA as the pH is raised to 5.1. We
observe higher than expected enzymatic activities for both IMC-PAA and IMC at extreme pH
conditions of 3.0 and 6.5. This can be attributed to lower free enzyme activity at those pH values.
In Figure 3.7 (c) and (d), we observe that IMC-PAA closely mimic free enzymes at a
temperature of 600C almost to the extent of 90-95%. Both avicelase and CMCase activities for
IMC-PAA consistently register 70% or more mimicking ability over the entire temperature
range.

3.3.4 Magnetic separation, recovery and reuse of immobilized enzymes

51

Figure 3.8. Specific activity of immobilized enzymes with and without PAA brushes vs. number
of cycles of hydrolysis.
Immobilized enzymes were subjected to four cycles of hydrolysis to test their efficiency after
repeated use. Data provided in Figure 3.8 shows that immobilized enzyme activity decreases
after every hydrolysis cycle. The decrease in enzymatic activity is not uncommon and is
routinely observed in most immobilized enzyme systems that are used multiple times. The
decrease could be because of protein desorption as well as denaturation. However, our results
(not shown here) indicated that in case of IMC-PAA, using covalent chemistry helped minimize
protein loss due to desorption. Compared to IMC, IMC-PAA displays higher resilience to
activity loss. As much as 55% of the original activity is retained after four cycles of use.

3.4. Conclusion
In summary, we demonstrated a polyelectrolyte based, ‘bottom-up’ approach for preparation
of maghemite-magnetite decorated graphene supports followed by successful immobilization of
cellulase using a combination of annealed polyacid brushes and zero-length spacer molecules.
The method proposed here is simple and can be easily repeated with other proteins/enzyme
systems. The cellulase hydrolysis results from the initial-rate experiments confirm considerably
high endo and exo-glucanase activities for soluble as well as insoluble substrates respectively.
Immobilized enzyme activity was studied as a function of both pH and temperature. Annealed
52

polyacid brushes being pH and heat sensitive, play a very crucial role in biocatalysis. The lower
enzymatic activity reported for most flat 2-D carriers can be overcome with the use of swollen
PAA brushes.The immobilized enzymes with PAA brushes closely mimic free enzymes (as
much as 90-95%) at 600C and a pH of 5.1. This is, by far, the highest cellulase activity reported
for two-dimensional flat carriers.
Three factors namely protein loading, activity of the loaded enzyme and more importantly,
the activity of loaded enzyme after several cycles of recycle should be taken into consideration
while evaluating the overall operational efficiency of immobilized enzymes. The proposed
immobilized enzyme design addresses the three key issues raised above. Firstly, a fair amount of
protein loading up to 2.5 ± 0.91 mg/gm was observed when PAA modified magnetoresponsive
graphene supports were used for immobilization. There is still some scope to increase the protein
loading by (a) using the ‘grafting-from’ approach to grow PAA brushes which is suggested as a
more efficient way to ensure dense coverage of PAA. However, this comes at the cost of
increased pKa value. Its effect on enzyme activity remains to be seen. (b) Immobilizing cellulase
at a pH close to its isoelectric point (pI). However, one should note that since cellulase is a
combination of several enzymes, an exact value for pI may not be available. Secondly, the pH
and heat tunable flexible polyacid brushes provide better mobility to the enzyme enhancing its
interaction with the substrate under consideration. Lastly, the covalent binding helps minimize
enzyme leeching, thus allowing multiple cycles of reuse. As much as 55% of the specific activity
of immobilized enzymes is retained after four cycles of use.

53

REFERENCES

54

REFERENCES

1.

Novoselov, K.S., et al., Two-dimensional atomic crystals. Proceedings of the National
Academy of Sciences of the United States of America, 2005. 102(30): p. 10451-10453.

2.

Van Noorden, R., Moving towards a graphene world. Nature, 2006. 442(7100): p. 228229.

3.

Zhang, L.M., et al., Functional Graphene Oxide as a Nanocarrier for Controlled Loading
and Targeted Delivery of Mixed Anticancer Drugs. Small, 2010. 6(4): p. 537-544.

4.

Liu, Z., et al., PEGylated nanographene oxide for delivery of water-insoluble cancer
drugs. Journal of the American Chemical Society, 2008. 130(33): p. 10876-+.

5.

Sun, X.M., et al., Nano-Graphene Oxide for Cellular Imaging and Drug Delivery. Nano
Research, 2008. 1(3): p. 203-212.

6.

Zhang, J.L., et al., Graphene Oxide as a Matrix for Enzyme Immobilization. Langmuir,
2010. 26(9): p. 6083-6085.

7.

Kim, J., J.W. Grate, and P. Wang, Nanostructures for enzyme stabilization. Chemical
Engineering Science, 2006. 61(3): p. 1017-1026.

8.

Kim, J., H.F. Jia, and P. Wang, Challenges in biocatalysis for enzyme-based biofuel cells.
Biotechnology Advances, 2006. 24(3): p. 296-308.

9.

Davison, B.H., et al., Enzyme stabilization by covalent binding in nanoporous sol-gel
glass for nonaqueous biocatalysis. Biotechnology and Bioengineering, 2001. 74(3): p.
249-255.

10.

Asuri, P., et al., Increasing protein stability through control of the nanoscale
environment. Langmuir, 2006. 22(13): p. 5833-5836.

11.

Asuri, P., et al., Water-soluble carbon nanotube-enzyme conjugates as functional
biocatalytic formulations. Biotechnology and Bioengineering, 2006. 95(5): p. 804-811.

12.

Pavlidis, I.V., et al., Development of effective nanobiocatalytic systems through the
immobilization of hydrolases on functionalized carbon-based nanomaterials. Bioresource
Technology, 2012. 115: p. 164-171.

13.

Feng, W. and P.J. Ji, Enzymes immobilized on carbon nanotubes. Biotechnology
Advances, 2011. 29(6): p. 889-895.
Lu, J., R.R. Weerasiri, and I. Lee, Carbon nanotubes tuned foam structures as novel
nanostructured biocarriers for lignocellulose hydrolysis. Biotechnology Letters, 2013.
35(2): p. 181-188.

14.

55

15.

Ji, P.J., et al., Lipase Covalently Attached to Multiwalled Carbon Nanotubes as an
Efficient Catalyst in Organic Solvent. Aiche Journal, 2010. 56(11): p. 3005-3011.

16.

Staunton, S. and H. Quiquampoix, Adsorption and Conformation of Bovine SerumAlbumin on Montmorillonite - Modification of the Balance between Hydrophobic and
Electrostatic Interactions by Protein Methylation and Ph Variation. Journal of Colloid
and Interface Science, 1994. 166(1): p. 89-94.

17.

Sinegani, A.A.S., G. Emtiazi, and H. Shariatmadari, Sorption and immobilization of
cellulase on silicate clay minerals. Journal of Colloid and Interface Science, 2005.
290(1): p. 39-44.

18.

Lu, J., et al., Stable Aqueous Suspension and Self-Assembly of Graphite Nanoplatelets
Coated with Various Polyelectrolytes. Journal of Nanomaterials, 2010. 18: p. 64-86.

19.

Lee, I., et al., Nanometal-decorated exfoliated graphite nanoplatelet based glucose
biosensors with high sensitivity and fast response. Acs Nano, 2008. 2(9): p. 1825-1832.

20.

Drzal, L.T. and H. Fukushima, Expanded graphite and products produced therefrom, , in
U.S. Patent Application 11/361, 2552006.

21.

Rao, C.N.R., A. Govindaraj, and B.C. Satishkumar, Functionalised carbon nanotubes
from solutions. Chemical Communications, 1996(13): p. 1525-1526.

22.

Zhao, W., C.H. Song, and P.E. Pehrsson, Water-soluble and optically pH-sensitive
single-walled carbon nanotubes from surface modification. Journal of the American
Chemical Society, 2002. 124(42): p. 12418-12419.

23.

Niyogi, S., et al., Ultrasonic dispersions of single-walled carbon nanotubes. Journal of
Physical Chemistry B, 2003. 107(34): p. 8799-8804.

24.

Badaire, S., et al., In situ measurements of nanotube dimensions in suspensions by
depolarized dynamic light scattering. Langmuir, 2004. 20(24): p. 10367-10370.

25.

Iler, R.K., Multilayers of Colloidal Particles. Journal of Colloid and Interface Science,
1966. 21(6): p. 569-&.

26.

Decher, G. and J.D. Hong, Buildup of Ultrathin Multilayer Films by a Self-Assembly
Process .1. Consecutive Adsorption of Anionic and Cationic Bipolar Amphiphiles on
Charged Surfaces. Makromolekulare Chemie-Macromolecular Symposia, 1991. 46: p.
321-327.

27.

Decher, G., and J. D. Hong, Buildup of ultrathin multilayer films by a self ‐assembly
process: II. Consecutive adsorption of anionic and cationic bipolar amphiphiles and

56

polyelectrolytes on charged surfaces. Berichte der Bunsengesellschaft für physikalische
Chemie,1991. 95(11): 1430-1434.
28.

Ariga, K., J.P. Hill, and Q.M. Ji, Layer-by-layer assembly as a versatile bottom-up
nanofabrication technique for exploratory research and realistic application. Physical
Chemistry Chemical Physics, 2007. 9(19): p. 2319-2340.

29.

Smuleac, V., D.A. Butterfield, and D. Bhattacharyya, Layer-by-layer-assembled
microfiltration membranes for biomolecule immobilization and enzymatic catalysis.
Langmuir, 2006. 22(24): p. 10118-10124.

30.

Dotzauer, D.M., et al., Catalytic membranes prepared using layer-by-layer adsorption of
polyelectrolyte/metal nanoparticle films in porous supports. Nano Letters, 2006. 6(10): p.
2268-2272.

31.

Wang, Y.J. and F. Caruso, Macroporous zeolitic membrane bioreactors. Advanced
Functional Materials, 2004. 14(10): p. 1012-1018.

32.

Siqueira, J.R., et al., Immobilization of biomolecules on nanostructured films for
biosensing. Biosensors & Bioelectronics, 2010. 25(6): p. 1254-1263.

33.

Kidambi, S., I. Lee, and C. Chan, Primary neuron/astrocyte co-culture on polyelectrolyte
multilayer films: A template for studying astrocyte-mediated oxidative stress in neurons.
Advanced Functional Materials, 2008. 18(2): p. 294-301.

34.

Hassler, B.L., et al., Renewable dehydrogenase-based interfaces for bioelectronic
applications. Langmuir, 2007. 23(13): p. 7127-7133.

35.

Hendricks, T.R., et al., Intact pattern transfer of conductive exfoliated graphite
nanoplatelet composite films to polyelectrolyte multilayer platforms. Advanced Materials,
2008. 20(10): p. 2008-2012.

36.

Kidambi, S., et al., Cell adhesion on polyelectrolyte multilayer coated
polydimethylsiloxane surfaces with varying topographies. Tissue Engineering, 2007.
13(8): p. 2105-2117.

37.

Sambanthamoorthy, K., et al., Identification of a novel benzimidazole that inhibits
bacterial biofilm formation in a broad-spectrum manner. Antimicrobial agents and
chemotherapy, 2011. 55(9): p. 4369-4378.

38.

Lee, I. and R.P. Wool, Polymer adhesion vs substrate receptor group density.
Macromolecules, 2000. 33(7): p. 2680-2687.

39.

Lee, I. and R.P. Wool, Controlling amine receptor group density on aluminum oxide
surfaces by mixed silane self assembly. Thin Solid Films, 2000. 379(1-2): p. 94-100.

57

40.

Lee, I., W. Wang, and S. Ji. DEVICE AND METHOD FOR PRETREATMENT OF
BIOMASS 2011.

41.

Medve, J., et al., Hydrolysis of microcrystalline cellulose by cellobiohydrolase I and
endoglucanase II from Trichoderma reesei: Adsorption, sugar production pattern, and
synergism of the enzymes. Biotechnology and Bioengineering, 1998. 59(5): p. 621-634.

42.

Woodward, J., Synergism in Cellulase Systems. Bioresource Technology, 1991. 36(1): p.
67-75.

43.

Tu, M.B., R.P. Chandra, and J.N. Saddler, Evaluating the distribution of cellulases and
the recycling of free cellulases during the hydrolysis of lignocellulosic substrates.
Biotechnology Progress, 2007. 23(2): p. 398-406.

44.

Galbe, M. and G. Zacchi, A review of the production of ethanol from softwood. Applied
Microbiology and Biotechnology, 2002. 59(6): p. 618-628.

45.

Tu, M.B., R.P. Chandra, and J.N. Saddler, Recycling cellulases during the hydrolysis of
steam exploded and ethanol pretreated lodgepole pine. Biotechnology Progress, 2007.
23(5): p. 1130-1137.

46.

Lee, D., A.H.C. Yu, and J.N. Saddler, Evaluation of Cellulase Recycling Strategies for
the Hydrolysis of Lignocellulosic Substrates. Biotechnology and Bioengineering, 1995.
45(4): p. 328-336.

47.

Taherzadeh, M.J. and K. Karimi, Enzyme-Based Hydrolysis Processes for Ethanol from
Lignocellulosic Materials: A Review. Bioresources, 2007. 2(4): p. 707-738.

48.

Tran, D.N. and K.J. Balkus, Perspective of Recent Progress in Immobilization of
Enzymes. Acs Catalysis, 2011. 1(8): p. 956-968.

49.

Sheldon, R.A., Enzyme immobilization: The quest for optimum performance. Advanced
Synthesis & Catalysis, 2007. 349(8-9): p. 1289-1307.

50.

Gokhale, A.A. and I. Lee, Cellulase Immobilized Nanostructured Supports for Efficient
Saccharification of Cellulosic Substrates. Topics in Catalysis, 2012: p. 1-16.

51.

van Beilen, J.B. and Z. Li, Enzyme technology: an overview. Current Opinion in
Biotechnology, 2002. 13(4): p. 338-344.

52.

Worden, R.M., et al., Simple fabrication of a highly sensitive glucose biosensor using
enzymes immobilized in exfoliated graphite nanoplatelets Nafion membrane. Chemistry
of Materials, 2007. 19(25): p. 6240-6246.

58

53.

Lee, I., et al., Intact pattern transfer of conductive exfoliated graphite nanoplatelet
composite films to polyelectrolyte multilayer platforms. Advanced Materials, 2008.
20(10): p. 2008-+.

54.

Ruhe, J., et al., Polyelectrolyte brushes. Polyelectrolytes with Defined Molecular
Architecture I, 2004. 165: p. 79-150.

55.

Biesalski, M., D. Johannsmann, and J. Ruhe, Synthesis and swelling behavior of a weak
polyacid brush. Journal of Chemical Physics, 2002. 117(10): p. 4988-4994.

56.

Dong, R., M. Lindau, and C.K. Ober, Dissociation Behavior of Weak Polyelectrolyte
Brushes on a Planar Surface. Langmuir, 2009. 25(8): p. 4774-4779.

57.

Wan, L.S., et al., Enzyme immobilization on electrospun polymer nanofibers: An
overview. Journal of Molecular Catalysis B-Enzymatic, 2009. 56(4): p. 189-195.

58.

Liu, Z.L., et al., Synthesis and characterization of ultrafine well-dispersed magnetic
nanoparticles. Journal of Magnetism and Magnetic Materials, 2004. 283(2-3): p. 258262.

59.

Massart, R., Preparation of Aqueous Magnetic Liquids in Alkaline and Acidic Media.
Ieee Transactions on Magnetics, 1981. 17(2): p. 1247-1248.

60.

Bradford, M.M., Rapid and Sensitive Method for Quantitation of Microgram Quantities
of Protein Utilizing Principle of Protein-Dye Binding. Analytical Biochemistry, 1976.
72(1-2): p. 248-254.

61.

Ghose, T.K., Measurement of Cellulase Activities. Pure and Applied Chemistry, 1987.
59(2): p. 257-268.

62.

Kotov, N.A., et al., Aqueous dispersions of single-wall and multiwall carbon nanotubes
with designed amphiphilic polycations. Journal of the American Chemical Society, 2005.
127(10): p. 3463-3472.

63.

Kotov, N.A., J.W. Ostrander, and A.A. Mamedov, Two modes of linear layer-by-layer
growth of nanoparticle-polylectrolyte multilayers and different interactions in the layerby-layer deposition. Journal of the American Chemical Society, 2001. 123(6): p. 11011110.

64.

Decher, G., Fuzzy nanoassemblies: Toward layered polymeric multicomposites. Science,
1997. 277(5330): p. 1232-1237.

65.

Correa-Duarte, M.A., et al., Alignment of carbon nanotubes under low magnetic fields
through attachment of magnetic nanoparticles. Journal of Physical Chemistry B, 2005.
109(41): p. 19060-19063.

59

66.

67.

Liz-Marzan, L.M. and M.A. Correa-Duarte, Carbon nanotubes as templates for onedimensional nanoparticle assemblies. Journal of Materials Chemistry, 2006. 16(1): p. 2225.
Nguyen, S.T., et al., Stable aqueous dispersions of graphitic nanoplatelets via the
reduction of exfoliated graphite oxide in the presence of poly(sodium 4-styrenesulfonate).
Journal of Materials Chemistry, 2006. 16(2): p. 155-158.

68.

Kotov, N.A., Layer-by-layer self-assembly: The contribution of hydrophobic interactions.
Nanostructured Materials, 1999. 12(5-8): p. 789-796.

69.

Kato, N., et al., Thin multilayer films of weak polyelectrolytes on colloid particles.
Macromolecules, 2002. 35(26): p. 9780-9787.

70.

Corot, C., et al., Recent advances in iron oxide nanocrystal technology for medical
imaging. Advanced Drug Delivery Reviews, 2006. 58(14): p. 1471-1504.

71.

Nigam, S., K.C. Barick, and D. Bahadur, Development of citrate-stabilized Fe(3)O(4)
nanoparticles: Conjugation and release of doxorubicin for therapeutic applications.
Journal of Magnetism and Magnetic Materials, 2011. 323(2): p. 237-243.

72.

Mukadam, M.D., et al., Particle size-dependent magnetic properties of gamma-Fe2O3
nanoparticles. Journal of Magnetism and Magnetic Materials, 2004. 272: p. 1401-1403.

73.

Xing, Y.C., Synthesis and electrochemical characterization of uniformly-dispersed high
loading Pt nanoparticles on sonochemically-treated carbon nanotubes. Journal of
Physical Chemistry B, 2004. 108(50): p. 19255-19259.

74.

Chimanage, P., et al., Properties of Cellulase Immobilized on Agarose-Gel with Spacer.
Biotechnology and Bioengineering, 1986. 28(12): p. 1876-1878.

75.

Park, H. and J.R. Robinson, Mechanisms of Mucoadhesion of Poly(Acrylic Acid)
Hydrogels. Pharmaceutical Research, 1987. 4(6): p. 457-464.

76.

Ma, Y.H., et al., Magnetically targeted thrombolysis with recombinant tissue
plasminogen activator bound to polyacrylic acid-coated nanoparticles. Biomaterials,
2009. 30(19): p. 3343-3351.

77.

Sugama, T., L.E. Kukacka, and N. Carciello, Nature of Interfacial Interaction
Mechanisms between Polyacrylic-Acid Macromolecules and Oxide Metal-Surfaces.
Journal of Materials Science, 1984. 19(12): p. 4045-4056.

78.

Zhang, H.N. and J. Ruhe, Polyelectrolyte complexes and multilayers at solid surface via
polymer brushes. Abstracts of Papers of the American Chemical Society, 2003. 225: p.
U621-U621.

60

79.

Ballauff, M., T. Neumann, and B. Haupt, High activity of enzymes immobilized in
colloidal nanoreactors. Macromolecular Bioscience, 2004. 4(1): p. 13-16.

80.

Caruso, F. and C. Schuler, Enzyme multilayers on colloid particles: Assembly, stability,
and enzymatic activity. Langmuir, 2000. 16(24): p. 9595-9603.

81.

Galaev, I.Y. and B. Mattiasson, 'Smart' polymers and what they could do in
biotechnology and medicine. Trends in Biotechnology, 1999. 17(8): p. 335-340.

82.

Imanishi, Y. and Y. Ito, Glucose-sensitive insulin-releasing molecular systems. Pure and
Applied Chemistry, 1995. 67(12): p. 2015-2021.

83.

Uhlmann, P., et al., In-situ investigation of the adsorption of globular model proteins on
stimuli-responsive binary polyelectrolyte brushes. Langmuir, 2007. 23(1): p. 57-64.

84.

Cole, M.A., et al., Stimuli-responsive interfaces and systems for the control of
protein–surface and cell–surface interactions. Biomaterials, 2009. 30(9): p. 18271850.

85.

Wang, Y.J. and F. Caruso, Nanoporous protein particles through templating mesoporous
silica spheres. Advanced Materials, 2006. 18(6): p. 795-+.

86.

Choi, H.K., M.K. Chun, and C.S. Cho, Mucoadhesive drug carrier based on interpolymer
complex of poly(vinyl pyrrolidone) and poly(acrylic acid) prepared by template
polymerization. Journal of Controlled Release, 2002. 81(3): p. 327-334.

87.

Ding, B., et al., Layer-by-layer structured films of TiO2 nanoparticles and poly(acrylic
acid) on electrospun nanofibres. Nanotechnology, 2004. 15(8): p. 913-917.

88.

Leaist, D.G., Coupled Diffusion of Weakly Ionized Poly-Electrolytes - Polyacrylic Acids
in Water. Journal of Solution Chemistry, 1989. 18(5): p. 421-435.

89.

Tam, T.K., et al., Polymer brush-modified electrode with switchable and tunable redox
activity for bioelectronic applications. Journal of Physical Chemistry C, 2008. 112(22):
p. 8438-8445.

90.

Gil, E.S. and S.M. Hudson, Stimuli-reponsive polymers and their bioconjugates. Progress
in Polymer Science, 2004. 29(12): p. 1173-1222.

91.

Zhou, J., et al., Reversible Immobilization and Direct Electron Transfer of Cytochrome c
on a pH-Sensitive Polymer Interface. Chemistry – A European Journal, 2007. 13(10): p.
2847-2853.

92.

Czeslik, C., O. Hollmann, and T. Gutberlet, Structure and protein binding capacity of a
planar PAA brush. Langmuir, 2007. 23(3): p. 1347-1353.

61

93.

Qin, W.S., et al., Cellulase activities in biomass conversion: measurement methods and
comparison. Critical Reviews in Biotechnology, 2010. 30(4): p. 302-309.

94.

Parnell, A.J., et al., Synthesis, characterization and swelling behaviour of
poly(methacrylic acid) brushes synthesized using atom transfer radical polymerization.
Polymer, 2009. 50(4): p. 1005-1014.

95.

Chibowski, S., M. Wiœniewska, and E. Opala Mazur, The effect of temperature on the
adsorption and conformation of polyacrylic acid macromolecules at the
ZrO2–polymer solution interface. Powder Technology, 2004. 141(1–2): p. 12-19.

96.

Wiśniewska, M., S. Chibowski, and T. Urban, Adsorption and thermodynamic
properties of the alumina–polyacrylic acid solution system. Journal of Colloid and
Interface Science, 2009. 334(2): p. 146-152.

97.

Lee, Y.M., S.H. Kim, and C.S. Cho, Synthesis and swelling characteristics of pH and
thermoresponsive interpenetrating polymer network hydrogel composed of poly(vinyl
alcohol) and poly(acrylic acid). Journal of Applied Polymer Science, 1996. 62(2): p. 301311.

62

Chapter 4: Conductive oxygen barrier films using supramolecular assembly
of graphene embedded polyelectrolyte multilayers
(This chapter is reproduced with permission from Journal of colloid and interface science, 409(2013), 219-226.)

4.0 Abstract
In this work, we show the development of synergistic graphene embedded polyelectrolyte
multilayer assembly. Three types of films were developed by exploiting the change in the
macromolecular structure of two polyelectrolytes namely branched polyethylenimine and
polyacrylic acid as a result of pH triggered dissociation. The films differ in thickness and
morphology and clear evidence of these differences are presented here. We also examined the
effect of these films on two properties namely oxygen permeability and sheet resistance.
Depending on the fabrication conditions, these films can provide an impressive three orders of
magnitude improvement in oxygen barrier property and close to five orders of magnitude
reduction in sheet resistance.

4.1 Introduction
In recent years there is a growing trend towards using flexible plastic as packaging
material due to its low cost, easy formability, light weight and reasonably good durability. Glass
has been traditionally used in the food packaging industry and fabrication of display panels on
account of its transparency, heat tolerance and gas shielding capability[1]. However today,
flexible plastics are making rapid inroads in replacing glass in most of these applications. In
order to truly compete as an alternative packaging material, flexible plastic needs to fulfill an
important requirement of preserving the product from environmental degradation. Dust,
oxidation and moisture are principal forces in nature that can have a deleterious effect on
63

perishable food products and sensitive electronic components[2]. Unfortunately, flexible plastics
despite their numerous advantages suffer from poor gas barrier properties[3]. Efforts have been
directed to overcome this problem by depositing inorganic coatings and thin films of polymer
blends to improve the gas shielding capability. Among the inorganic materials, aluminum and
silicon oxide coatings are excellent candidates to limit the permeation of gas molecules.
However, inorganic coatings especially pure SiO2 thin films require high temperature for
effective deposition[4]. Most flexible plastics have low glass transition temperatures (T g)
imposing a limitation on the use of inorganic coatings.
Flexible plastics are positioned to make key contributions to the electronics industry in
the near future. Devices such as organic light emitting diodes (OLED) expected to corner a 40%
share of the mobile phone display market by 2015 present exciting growth prospects for the
plastics industry[5]. With the world rapidly moving towards miniaturization, researchers have
sought to make electronic components more compact by designing new prototypes that promote
improved electrical conductivity and gas barrier properties. This would be particularly useful in
the fabrication of certain semiconductor materials such as the dielectric constant oxides that are
frequently used in the construction of large capacitors. These devices are liable to fail in the
presence of non-conducting oxide coatings[6, 7]. Other devices with similar conductive gas
barrier requirements include solar cells grown on flexible supports and flat panel displays[8-10].
Silicon oxide though available ubiquitously, acts as an electrical insulator[11]. So it cannot be
directly applied for the above applications. In recent years, there has been a growing interest in
the use of nanomaterials to supplement the gas barrier properties of thin polymer films. A
number of theoretical models have been used to examine the critical role played by the aspect
ratio of nano/colloidal materials dispersed in polymer matrix in improving the gas barrier

64

properties[12-14]. Upon encountering a nanofiller, gas molecules are forced to adopt a tortuous
pathway producing a significant lag in the transmission. Therefore, high aspect ratio
nanomaterials such as clay platelets[15-17], mica sheets[18, 19], cellulose[20, 21] and graphene
oxide[22] have been used to build gas resistant thin films. Inspite of their high gas/moisture
barrier properties, these nanomaterials provide little advantage in the preparation of conductive
films. On the other hand, graphene with its sp 2 hybridized 2D structure has unique physical and
electronic properties[23]. Thin graphene layers are impermeable to most gases and could serve as
excellent nanofillers in the polymer matrix[24]. High charge mobility and ballistic transport at
room temperature endow graphene with superior electrical conductivity making it a promising
candidate for applications such as electrochemical sensing, field effect transistors (FET) and
supercapacitors[25].
Layer-by-layer (LbL) assembly is a versatile technique that can be used to add multiple
functionalities to a host of substrates. The success of the LbL technique lies in its ubiquity and
simple operation. The extension of Iler’s work[26] to the polyelectrolyte multilayer assembly by
Decher and co-workers[27, 28] has given rise to a large number of nanoscopic functional
materials. The non-covalent functionalization of nanomaterials using polyelectrolytes followed
by sequential assembly is a form of LbL that has successfully demonstrated its utility. A number
of interactions such as electrostatic, hydrophobic, hydrogen bonding, are ascribed to the
formation of multilayered polyelectrolyte LbL assembly.[29] Multifunctional tailor-made
substrates fabricated using this route have found several applications such as in catalysis[30],[31,
32], membrane science[33, 34], biosensors[35-38] and drug delivery[39-42]. Ionic strength and
pH of the solution can influence the adsorption process producing films of distinct morphology
and properties[43-46].

65

With an estimated theoretical elastic modulus of single-layered graphene projected to be
1 TPa, and surface conductivity about 50 × 106 S/cm, GNPs enjoy superior mechanical and
electrical properties[47]. In recent years, methods such as thermal expansion, dielectric heating
and acid intercalation have facilitated exfoliation of graphitic flakes enabling the production of
high surface area single layered graphene nanoplatelets[48]. In this study we report the LbL
assembly of GNP embedded polyelectrolyte multilayers. Unlike inorganic coatings prepared
under high temperature conditions, the GNP/PEM LbL assembly can be processed at room
temperature. Similarly, compared to the recently developed fabrication methods for free standing
graphite papers[49] and graphene nanoplatelet-high density polyethylene (GNP-HDPE)
nanocomposite[50], the LbL process can be accomplished under milder process conditions using
lower energy input. The GNP embedded polyelectrolyte films serve two purposes. Firstly, they
serve as barriers to gas permeation. Secondly, graphene with its unique electronic configuration
is positioned to deliver improved electrical conductance. The combination of gas barrier
properties and electrical conductance make these films attractive candidates for applications in
the semiconductor and electronics industry. Graphene dispersion in polyelectrolytes followed by
its LbL assembly has been a subject of extensive examination[51, 52]. Both strong and weak
polyelectrolytes have been utilized for this purpose with varying degrees of success. While
strong polyelectrolytes are fairly stable over a wide range of pH, weak polyelectrolytes are
known to exhibit a remarkable variation in the charge distribution when subjected to a pH
change[53]. This can affect the film thickness, the mechanism of interaction, the adsorption
kinetics and the quality of adhesion of the films to the base substrate. Unlike other studies that
utilized graphene oxide followed by its reduction[54, 55], our process did not use any oxidation
or reduction steps.

Instead we chose two weak polyelectrolytes namely branched

66

polyethylenimine (BPEI) and polyacrylic acid (PAA) for LbL deposition. BPEI is rich in amino
pendant groups giving it a cationic character. The abundant availability of primary, secondary
and tertiary amino residues provides ample opportunities to use BPEI to complement the anionic
carboxylic group rich PAA in a multilayered assembly. Using different process conditions three
sets of GNP embedded LbL assemblies were developed. They comprised of hydrogen bonding,
electrostatic and composite interactions (combination of hydrogen bonding and electrostatic
interactions).

4.2 Experimental Procedures
4.2.1 Materials
Polyacrylic acid (PAA) sodium salt (35 wt% solution in water) and branched polyethylenimine
(BPEI) average Mw ~25,000 were purchased from Sigma Aldrich. Poly (ethylene terephthalate)
(PET) films (Mylar A: thickness approximately 76.2 µm) produced by Dupont-Teijin were
obtained from Tekra (New Berlin, WI). Exfoliated graphene nanoplatelets (GNP) (BET surface
area: 300 m2/gm and size 1µm, and average thickness of 2-4 nm) were purchased from XG
Sciences, Lansing, MI. All aqueous solutions were prepared using deionized (DI) water (>18.1
M) supplied by a Barnstead Nanopure Diamond-UV purification unit equipped with a UV
source and final 0.2 m filter. Unless specified otherwise, all procedures were carried out at
room temperature.

4.2.2 Layer-by-layer (LbL) assembly of polyelectrolyte modified GNP
For the hydrogen bonded assembly, the concentration of PAA and BPEI was adjusted to 2 mg/ml
and 1 mg/ml respectively. GNP platelets were dispersed in BPEI solution (now called BPEIGNP ) using a Fisher Scientific Ultrasonic probe (16 W) by sonicating for 1 hour. GNP-BPEI

67

solution was stirred overnight before use. The pH of the PAA and BPEI-GNP solution was
adjusted to 4.0 and 7.5 respectively. For the deposition of the electrostatic layers, GNP were
dispersed in 2 mg/ml BPEI solution using ultrasonication followed by magnetic stirring for at
least 8 hours. The excess polyelectrolyte is removed by passing the BPEI modified graphene
solution through a 0.22 µm cellulose-acetate membrane-filter systems from Nalgene. The BPEI
modified graphene platelets collected on the filter membrane were washed with copious quantity
of DI water and subjected to another cycle of filtration. They are finally suspended in DI water
and the pH is adjusted to 3.5-3.8.

The PAA solution used for electrostatic deposition

(concentration: 2mg/ml) was adjusted to a pH of 6.0. For the preparation of composite films, the
desired number of hydrogen bonded bilayers was initially deposited. This was followed by the
required number of electrostatic layers.
The LbL assembly was carried out using a customized Carl-Zeiss slide-stainer. Before
depositing the multilayers, the surface of the polyester films was cleaned using a Harrick plasma
cleaner (Harrick Scientific Corporation, Broading Ossining, NY). The films were subjected to air
plasma for a period of 15 minutes. Air plasma helps in surface activation by producing
hydrophilic moieties on the surface of the films. The immersion time was fixed to 15 minutes
followed by a DI water rinse cycle of 2 minutes. After the deposition of each layer, the films
were allowed to dry naturally for 5 minutes. After the LbL process was complete, the films were
heated in an oven maintained at 650C for 2 hours to promote thermal crosslinking.

4.2.3 Multilayers Characterizations
Multilayer films of GNP films on PET surface were characterized using scanning
electron microscopy (SEM, JEOL 6610LV). The samples were cross-sectioned to reveal the
thickness of the deposited films using a single edged razor blade that was cleaned with ethanol
68

prior to use. The samples were coated with a thin layer of osmium for enhanced conductivity
before SEM measurements. The surface resistance of multilayer films was measured with a
Gamry Instruments Femtostat Station with frequency ranging from 1-10000 Hz.

4.2.4 Oxygen Transmission Measurements
Oxygen barrier properties of films were tested in accordance using a MOCON 2/21 ML
instrument (Mocon Inc., Minneapolis, MN, USA). The samples were tested at 0% relative
humidity (RH) at a temperature of 230C.

4.3 Results and Discussion
Hydrophilic polymers with numerous polar groups typically exhibit low specific free
volume[56]. The polar groups can undergo hydrogen bonding producing a compact polymer
structure that can prevent the permeation of gas molecules. The LbL assembly of BPEI and PAA
is a pH driven process. These polymers form interpenetrating hydrogen bonded network when
the terminal groups of the polymer chains exist in uncharged state[57]. The presence of
impermeable material in the polymer matrix can further add to the gas barrier properties.
Materials with high aspect ratio are generally preferred as fillers because of their ability to
significantly increase the lag time for gas permeation through the polymer. Commercially
available GNPs used in this study were prepared by using a unique non-oxidizing technology
that enabled the platelets to retain their sp2 configuration[58]. With an individual platelet
thickness averaging 2 nm, and particle diameter of about 1-2 microns, the graphene nanoplatelets
used in this study exhibit a significantly high aspect ratio increasing the chances of providing
strong resistance to gas permeation. The elongated sheet-like geometry of graphene combined
with high aspect ratio is also expected to achieve low percolation threshold value due to its

69

ability to form a conducting network as compared to spherical or elliptical fillers[59]. However,
the platelets have a tendency of stacking up reducing the effectiveness of using graphene as a
high aspect ratio material. However ultrasonication of graphene followed by continuous
overnight stirring helps in improved dispersion of the nanoplatelets. The ultrasonication disrupts
the stacking of the graphene platelets exposing individual platelets to BPEI functionalization.
BPEI adsorbs on the graphene platelets due to hydrophobic interactions[52]. In case of
negatively charged substrates, BPEI can assume a flat ‘pancake-like’ structure as the cationic
polymer segments in BPEI compensate the surface charges[60]. However in this case, the
interaction between the neutral GNP and BPEI results in an optimized structure with bulky
globular BPEI adsorbing on the surface of GNP. The polyelectrolyte modification minimizes
graphene-graphene interaction improving the exfoliation of the platelets within the BPEI matrix.
In our study we tested three different LbL configurations of graphene embedded polyelectrolyte
multilayers. The effect of these configurations on oxygen barrier properties and electrical sheet
resistance has been shown in the subsequent sections. Mylar PET film about 76 µm thick was
used as a base substrate in all cases.

4.3.1 Types of LbL deposited GnP/PEM films
The schematic diagram shown in Figure 4.1 shows the three types of LbL assembly namely
hydrogen bonded, electrostatic and composite films. This section highlights the role played by
adsorption conditions to produce GNP/PEM films with specific type of interactions.

70

Figure 4.1. Supramolecular assembly of graphene embedded polyelectrolyte multilayers.
(a)Multilayers assembled using hydrogen bonded assembly. m denotes number of bilayers
deposited. (b) Multilayers deposited using electrostatic interactions. n denotes the number of
electrostatic bilayers deposited. (c) Composite film consisting of ‘m’ number of hydrogen
bonded bilayers followed by ‘n’ number of electrostatic bilayers.
4.3.1.1 Hydrogen bonded films
Figure 4.1a shows a schematic representation of the hydrogen bonded multilayers. m indicates
the number of bilayers of hydrogen bonded graphene films deposited on the PET substrate. In
this configuration, a multilayer assembly of BPEI-GNP and PAA was assembled on PET. The
pH of the BPEI-GNP solution was adjusted to 7.5 whereas PAA was adjusted to 4.0. The native
pH of BPEI-graphene is between 9.5-10.2. We observed that the BPEI-graphene solution when
used in the LbL assembly without pH adjustment resulted in non-uniform coatings. We believe
that the large difference between the pH values of BPEI-graphene and PAA solution can induce
precipitation of certain charged groups in both the polyelectrolyte solutions leading to poor
quality of LbL deposited film. The pKa value for the primary and secondary amines in BPEI is
71

approximately 4.5 and 6.7 respectively[61]. At a pH 7.5, most of the amine residues in BPEI
should exist in uncharged state. The pKa value of PAA is close to 4.8[62]. Hence, at a pH 4.0,
most of the terminal groups in PAA exist in uncharged state (carboxylic acid groups –COOH)
-

rather than as carboxylate ion -COO . The presence of neutral terminal groups for both PAA and
BPEI results in the deposition of highly coiled, loopy polyelectrolyte network. The interdiffusion
of the polyelectrolyte segments followed by thermal crosslinking for 2 hours at 65 0C gives rise
to compact film architecture.

4.3.1.2 Electrostatically assembled films
Another type of LbL configuration was tested by retaining the same polyelectrolytes as before,
except for two changes. The schematic representation of the same is shown in Figure 4.1b. n
denotes the number of electrostatic bilayers deposited.

The deposition of BPEI-graphene

solution during the LbL process represents a unique situation where both BPEI and
polyelectrolyte modified graphene are competing for adsorption on the negatively charged PAA
chains. Lu and co-workers[52] addressed this issue by subjecting the polyelectrolyte-graphene
solution to vacuum filtration. By filtering the polyelectrolyte-graphene solution through 0.22 µm
filter, Lu and co-workers removed the excess unbound polyelectrolyte minimizing the ill-effects
of competitive adsorption. The removal of non-conducting polyelectrolyte solution should also
help in improved interconnection between individual graphene platelets. To counter the dense
nature of the films produced in the earlier configuration, we implemented another change that
targeted the interactions between the cationic and anionic components of the LbL assembly. The
BPEI functionalized graphene after vacuum filtration was dispersed in DI water and the pH was
adjusted to 3.5-3.8. This pH value is lower than the pKa value of the primary amines in BPEI
(usually around 4.5). At this pH condition, the BPEI modified graphene platelets are positively
72

charged. The BPEI chains that are functionalized on graphene platelets occur in protonated state
(-NH3+). Meanwhile, the pH of PAA solution was adjusted to 6.0. This value is higher than the
pKa value of PAA. Thus at this pH, PAA chains mostly exist in the ionized form. Though ionic
crosslinks are possible, the overall LbL assembly is governed by electrostatics.

4.3.1.3 Composite film
The synergistic combination of the above two systems is expected to deliver a result that
includes contributions due to both hydrogen bonding and electrostatics, as shown in Figure 4.1c.
Using the same pH conditions as described before, a composite film consisting of the desired
number of hydrogen bonded layers followed by electrostatic layers were deposited. The
composite film is depicted as mH+ nE where H and E stand for hydrogen bonded and
electrostatic films respectively.

4.3.2 Morphology and Thickness of GNP embedded PEM films
The three types of LbL configurations listed above have distinct impact on the
morphology and the thickness of the deposited films. Previous research has established that
hydrogen bonded and electrostatically assembled PEMs can exhibit differences in morphology
such as thickness and surface roughness[63, 64]. Some of the hydrogen bonded multilayers are
reported to exhibit an exponential growth pattern[65, 66]. After a certain number of bilayers are
deposited these films may show linear increase in thickness[67]. Most electrostatic assemblies
grow linearly[68] enabling easy estimation of thickness of individual bilayer. However for
exponential or hybrid (exponential+linear) growth regimes, the task is harder and requires more
detailed analysis. In our case, the presence of graphene in the multilayer assembly can add
complexity to the deposition process and influence the growth of these films. Though

73

establishing the growth pattern of GNP embedded multilayers is not within the scope of this
work, the morphology and thickness information about these films can provide useful insight
into understanding the gas barrier and sheet resistance properties.
Figure 4.2 shows SEM micrographs of the three LbL configurations for a 20 bilayer
sample. The cross-sectional images (Figures 4.2 a,c,e) give an idea of the thickness of the
multilayers whereas Figures 4.2 (b,d,f) establish the surface morphology of these films. For the
20 bilayer sample under consideration, the thickness is highest for the hydrogen bonded
assembly (about 5.5 µm). Electrostatic assembly results in thinner film (See Figures 4.2c). With
a thickness of around 705 nm, the electrostatic film is substantially thinner than the hydrogen
bonded film. The composite film (Figure 4.2e) has a thickness of 3.77 µm (lower than hydrogen
bonded film but higher than electrostatically assembled multilayers). The effect of the interactive
forces associated with these configurations is evident from Figures 4.2 b,d,f. Figure 4.2b shows
surface morphology of 20 bilayer hydrogen bonded assembly. The surface is relatively smooth
and free of graphene agglomerations. On the other hand, Figure 4.2d shows the agglomerations
of GNPs as a result of the electrostatic assembly. We suspect that the formation of aggregates is
due to poor adhesion of BPEI functionalized graphene on the PET substrate. Hydrogen bonded
films generally have good adhesive properties due to the presence of polar groups. A lower share
of hydrogen bonding interactions in case of electrostatic films can reduce

74

Figure 4.2 SEM micrographs of 20 bilayer GNP embedded PEM assembly. a, c, e show sectional
view of hydrogen bonded, electrostatic and composite (10H + 10E) multilayers respectively.
Thickness of the coating is 5.5 µm, 0.705 µm, 3.77 µm respectively. b, d, f show surface
morphology of hydrogen bonded, electrostatic and composite films respectively.
75

the adhesive capacity of the multilayers to hold the GNPs together. Figure 2f shows the surface
morphology of a composite film. As expected, the surface morphology is a synergistic
combination of the two assemblies shown previously. The surface has a certain texture
resembling the hydrogen bonded films along with some degree of graphene agglomeration
associated with electrostatic films. The adhesion provided by the underlying hydrogen bonded
films facilitates good surface coverage by the electrostatic films. The improved interconnection
between individual GNPs in the electrostatic layers promotes reduction in sheet resistance. The
other cases (4, 10, 15 and 30 bilayers samples) show similar thickness trend (See Figures.
4.3,4.4,4.5)

76

Figure 4.3 SEM micrographs of 4 bilayer GNP embedded PEM assembly. a, c, e show sectional
view of hydrogen bonded, electrostatic and composite (2H + 2E) multilayers respectively.
Thickness of the coating is 0.429 µm, 0.167 µm, 0.378 µm respectively. b, d, f show surface
morphology of hydrogen bonded, electrostatic and composite films respectively.

77

Figure 4.4 SEM micrographs of 10 bilayer GNP embedded PEM assembly. a, c, e show sectional
view of hydrogen bonded, electrostatic and composite (5H + 5E) multilayers respectively.
Thickness of the coating is 3.47 µm, 0.455 µm, 0.893 µm respectively. b, d, f show surface
morphology of hydrogen bonded, electrostatic and composite films respectively.

78

Figure 4.5 SEM micrographs of composite films with fixed number of hydrogen bonded
multilayers (10 bilayers) and variable number of electrostatic layers a and c show sectional view
of composite films with (10H + 5E) and 10H +20 E) multilayers respectively. Thickness of the
coating is 3.56µm and 4.02 µm respectively. b and d show surface morphology of the composite
films (10H +5E) and (10H + 20E) respectively.

79

4.3.3 Oxygen barrier properties of GNP/PEM multilayers
Figures 4.6 a-d help us compare the effect of hydrogen bonding, electrostatic and
composite interactions on the barrier properties of PET films. A bare PET film has an oxygen
transmission rate (OTR) of 30.3 cc O2/m2.day atm. Deposition of just 4 bilayers of hydrogen
bonded BPEI-graphene/PAA reduces the OTR by 90.07% (Figure 4.6a). Measurements were
also done after depositing 10 and 20 bilayers (See Figures 4.6b and 4.6c). The OTR decreases as
the number of bilayers increase. A 20 bilayer film (Fig. 4.6c) shows an impressive 98.6%
reduction in the OTR value. The electrostatically assembled GNP/PEM films present a
completely different result. Lack of hydrogen bonding can cause little or no significant reduction
in the specific free volume greatly diminishing the ability of the graphene embedded film to
function as effective oxygen barrier. A 20 bilayer electrostatic film exhibits a mere 14%
reduction in OTR (Figure 4.6c). Compared to electrostatic films, composite films display
improved gas barrier properties (Figures 4 a-d). These films show a decreasing trend in OTR as
the number of bilayers increased. When the deposition of hydrogen bonded layers reached 10,
the OTR reduced by 98%. Increasing the number of electrostatic layers alone on hydrogen
bonded films has little effect on the OTR values. This can be inferred from Figure 4.6d where
increasing the number of electrostatic layers (from 5 to 20) on fixed number of hydrogen bonded
films results in a very small decrease in the OTR.

80

Figure 4.6 Oxygen transmission rate (OTR) for different types of configurations. (a) 4 bilayer
system. The composite film consists of 2 hydrogen bonded and 2 electrostatic bilayers. (b) 10
bilayer system. Composite film consists of 5 hydrogen bonded and 5 electrostatic bilayers. (c) 20
bilayer system. Composite film consists of 10 hydrogen bonded and 10 electrostatic bilayers. (d)
Composite films consisting of 10 hydrogen bonded (H) followed by varying number of
electrostatic (E) bilayers.
Table 4.1 Oxygen permeability coefficient of hydrogen bonded films on PET substrate at STP
conditions
Number of
Thickness
Permeability coefficient
bilayers
of the
(10-18 cc. cm/cm2.s.Pa)
coating µm
Film (Pf)
Total (Pt)
4 bilayers
0.439
3.02
244.5
10 bilayers
3.47
4.55
53.45
20 bilayers
5.5
4.82
37.68

81

Table 4.2 Oxygen permeability coefficient of electrostatic films on PET substrate at STP
conditions
Number of
Thickness
Permeability coefficient
bilayers
of the
(10-18 cc. cm/cm2.s.Pa)
coating µm
Film (Pf)
Total (Pt)
4 bilayers
0.167
76.7
2146
10 bilayers
0.455
25.2
2205
20 bilayers
0.705
24.2
2094

Table 4.3. Oxygen permeability coefficient of composite films on PET substrate at STP
conditions
Number of
Thickness
Permeability coefficient
bilayers
of the
(10-18 cc. cm/cm2.s.Pa)
coating µm
Film (Pf)
Total (Pt)
4 bilayers
0.379
16.6
1003.4
10 bilayers
0.893
10.1
374.03
20 bilayers
3.77
7.12
76.79
Table 4.4. Oxygen permeability coefficient of composite films (variable electrostatic layers) on
PET substrate at STP conditions
Number of
Thickness of
Permeability coefficient
bilayers
the coating
(10-18 cc. cm/cm2.s.Pa)
µm
Film (Pf)
Total (Pt)
10 H+ 5E
3.56
7.36
83.43
10 H+ 10E
3.77
7.12
76.70
10H + 20E
4.02
6.78
69.26

The SEM data from the previous section also provides critical information about film
thickness that can be used for the estimation of the oxygen permeability coefficient. Unlike the
OTR, the permeability coefficient takes into account the contribution of the thickness of the film
and can provide useful information to compare different systems. For the calculation of film
permeability coefficient, its contribution to the total permeability coefficient (inclusive of both
the film as well as the PET substrate) was decoupled using a method described elsewhere[69].
Tables 4.1-4.4 show oxygen permeability coefficients for different LbL configurations at STP
conditions. With thickness ranging from nanometers to few microns, the GNP/PEM films
82

developed in this study show superior oxygen barrier properties compared to a 76.2 µm thick
PET film. The graphene embedded PEM films show low permeability coefficients almost three
orders of magnitude lower than the uncoated PET substrate (2433×10 -18 cc.cm/cm2.s.Pa). Since
hydrogen bonded systems display the best oxygen barrier properties, a 20 bilayer hydrogen
bonded BPEI/PAA system (without GNP) was assembled under the same pH conditions and its
permeability coefficient was estimated to be 3.05×10-18 cc.cm/cm2.s.Pa . However, the same
result can be achieved by using just 4 bilayers of hydrogen bonded GNP/PEM multilayers
assembled under similar deposition conditions. Among polymers, ethylene vinyl alcohol
(EVOH) finds great application in the packaging industry on account of its low oxygen
permeability. EVOH films with ethylene concentration ranging from 25-45 mol% are reported to
exhibit enhanced gas barrier properties[70]. However, the GNP embedded hydrogen bonded and
composite PEM films developed in this study have oxygen permeability coefficients that are at
least 10 times lower than the permeability coefficient of a 15 µm thick EVOH film containing 44
mol% ethylene and about 1.4-3 times lower than the similarly sized EVOH film with 32 mol%
ethylene[70]. The GNP/PEM films also show vastly superior oxygen barrier properties as
compared to other graphene based nanocomposites reported in literature[49, 71]. With
permeability coefficient in the range of 10 -18 cc.cm/cm2.s.Pa, these coatings (hydrogen bonded
and composite films) are comparable to SiO2 thin films deposited on commercial gas barrier foils
using plasma enhanced chemical vapor deposition (PECVD) technique[72]. Some important
observations can be made from the data presented in Tables 4.1-4.4. Table 4.1 shows the effect
of increasing number of hydrogen bonded GNP multilayers on oxygen permeability. While the
total permeability coefficient (Pt) continues to register a drop, the permeability coefficient of the
deposited film (Pf) increases slightly. This could be because the increase in the thickness of the

83

coating negates the corresponding small drop in OTR. The data shown in Table 4.2 reinforces
the poor barrier properties of electrostatic films. With increase in the number of bilayers, the data
shows no clear trend partly because of poor adhesion of the GNP multilayers in absence of
uncharged polar groups. Table 4.3 shows a healthy decrease in P f and Pt. The increasing number
of hydrogen bonded multilayers facilitates a decrease in the permeability coefficient. Table 4.4
shows the effect of increasing number of electrostatic films deposited on fixed number of
hydrogen bonded films. Both Pt and Pf do not register a significant decrease which is consistent
with our hypothesis that electrostatic films do not contribute to barrier properties.

4.3.4 Electrical resistance of the GNP/PEM multilayers
The electrical sheet resistance was also tested as a function of the number of bilayers. As
seen from Figure 4.7a, at 1 Hz, the electrical sheet resistance of the bare PET film is about 111.1
GΩ. The high value of resistance indicates that the film is highly resistive. The deposition of
hydrogen bonded BPEI-graphene/PAA films reduces the resistance. The PET substrate shows a
successive reduction in the resistance value for 4, 10 and 20 bilayers. For 20 bilayers, the value
reduces to about 142 GΩ. Though there is a substantial reduction in the electrical resistance of
the PET film, the values still end up in GΩ. We suspect that the high values of sheet resistance
are a result of the dense network of hydrogen bonded polyelectrolytes surrounding the graphene
platelets. The polymer wrapping of GNPs is known to reduce their conductivity.

84

Figure 4.7. Sheet resistance for different configurations of LbL assembled graphene embedded
multilayers. Sheet resistance at 1 Hz for various samples is as follows:
(a) Bare PET film: 111.1 GΩ, hydrogen bonded film: 108.2 GΩ, electrostatic film: 61.3 GΩ,
composite film: 57.6 GΩ.
(b) Bare PET film: 115.2 GΩ, hydrogen bonded film: 90.9 GΩ, electrostatic film: 54.3 GΩ ,
composite film: 50.2 GΩ.
(c) Bare PET film: 110.2 GΩ, hydrogen bonded film: 40.8 GΩ, electrostatic film: 0.949 GΩ,
composite film: 24.8 MΩ.
(d) Bare PET film: 112.3 GΩ, composite film (10H + 5 E): 31.2 GΩ, (10H + 10 E): 24.8 MΩ,
(10H+20 E): 1.14 MΩ

Further, the loopy, coiled conformation of the polyelectrolytes prevents the
interconnection between the individual graphene units limiting the electrical conductance of the
film.

As inferred from the SEM images, unlike the dense hydrogen bonded films,

electrostatically assembled BPEI modified graphene/PAA films are substantially thinner. The
combined effect of elimination of excess polyelectrolyte and thin film structure promotes further
85

reduction in the electrical sheet resistance. As seen from Fig. 4.7c, a 20 bilayer electrostatic film
exhibits a sheet resistance of 0.95 GΩ. However, we suspect that further reduction in electrical
resistance is hard to achieve due to poor adhesion of the multilayers to the substrate PET film.
For the composite films, the electrical resistance drops further. Figure 7d shows substantial
reduction in the sheet resistance as the number of electrostatic layers is increased. The sheet
resistance for a composite film consisting of 10 hydrogen bonded bilayers followed by 20
electrostatic bilayers drops to about 1.14 MΩ (Figure 4.7d). This represents a five orders of
magnitude reduction in sheet resistance. We believe that the composite film shows improved
performance due to the synergistic effect of hydrogen bonded and electrostatic multilayers.
Hydrogen bonded films have good adhesive properties due to presence of polar groups. The
presence of hydrogen bonded films provides the adhesion to the electrostatic layers.

4.4 Conclusion
In summary, we showed the effect of three configuration types of graphene embedded
polyelectrolyte systems. Hydrogen bonded graphene films showed good oxygen barrier property.
Electrostatic films given their thin morphology and absence of excessive non-conducting
polyelectrolyte exhibit lower sheet resistance. A composite film comprising of graphene
embedded hydrogen bonded and electrostatic coatings satisfied both these requirements. The
composite film showed oxygen permeability coefficient that is comparable to SiO 2 coatings with
electrical sheet resistance five orders of magnitude lower than the base substrate. Besides the
configurations described here, there is certainly more scope for improvement especially with
respect to the electrostatic layers. The deposition process for the electrostatic layers comprising
of BPEI modified graphene/PAA multilayers needs to be monitored for precipitation of charged
groups. Being a pH sensitive system makes the solutions prone to sedimentation. This problem
86

can be overcome by sonicating the BPEI modified graphene frequently (every 2 hours) and
adjusting the pH of the solutions in case of change. A more robust reproducible design can be
obtained by casting films made from polystyrene sulfonate (PSS) functionalized graphene and a
conjugate polycation such as poly(diallydimethylammonium chloride) (PDAC) which are less
sensitive to pH changes. These modifications may entail making suitable changes in the
hydrogen bonded layers. Another parameter that significantly alters the morphology and
potentially the conductivity of the coatings is the annealing temperature. The glass transition
temperature (Tg) of PET is reported to be about 700C[73]. This has restricted the annealing
temperature in our experiments to 650C. Higher temperatures generally contribute to increased
crosslinking (improves gas barrier properties) and evaporation of moisture from the multilayers
(facilitates improved conductivity). Using a polymer substrate with increased temperature
tolerance can help achieve these objectives.

87

REFERENCES

88

REFERENCES

1.

Ebina, T. and F. Mizukami, Flexible transparent clay films with heat-resistant and high
gas-barrier properties. Advanced Materials, 2007. 19(18): p. 2450-+.

2.

Charton, C., et al., Development of high barrier films on flexible polymer substrates. Thin
Solid Films, 2006. 502(1-2): p. 99-103.

3.

Shirakura, A., et al., Diamond-like carbon films for PET bottles and medical
applications. Thin Solid Films, 2006. 494(1-2): p. 84-91.

4.

Teshima, K., et al., Gas barrier properties of silicon oxide films prepared by plasmaenhanced CVD using tetramethoxysilane. Vacuum, 2002. 66(3-4): p. 353-357.

5.

Laaperi, A., Disruptive factors in the OLED business ecosystem. Information Display,
2009. 9(09): p. 09.

6.

Grill, A., et al., Base Electrodes for High Dielectric-Constant Oxide Materials in Silicon
Technology. Journal of Materials Research, 1992. 7(12): p. 3260-3265.

7.

Grill, A., C. Jahnes, and C. Cabral, Layered TaSiN as an oxidation resistant electrically
conductive barrier. Journal of Materials Research, 1999. 14(4): p. 1604-1609.

8.

Dennler, G., et al., Flexible, conjugated polymer-fullerene-based bulk-heterojunction
solar cells: Basics, encapsulation, and integration. Journal of Materials Research, 2005.
20(12): p. 3224-3233.

9.

Jorgensen, M., K. Norrman, and F.C. Krebs, Stability/degradation of polymer solar cells.
Solar Energy Materials and Solar Cells, 2008. 92(7): p. 686-714.

10.

Chou, C.-T., et al., Transparent Conductive Gas-Permeation Barriers on Plastics by
Atomic Layer Deposition. Advanced Materials, 2013. 25(12): p. 1750-1754.

11.

Inagaki, N., S. Tasaka, and M. Makino, Plasma polymer deposition from mixture of
tetramethoxysilane and oxygen on PET films and their oxygen gas barrier properties.
Journal of Applied Polymer Science, 1997. 64(6): p. 1031-1039.

12.

Lape, N.K., E.E. Nuxoll, and E.L. Cussler, Polydisperse flakes in barrier films. Journal of
Membrane Science, 2004. 236(1): p. 29-37.

13.

DeRocher, J.P., et al., Barrier membranes with different sizes of aligned flakes. Journal of
Membrane Science, 2005. 254(1-2): p. 21-30.
89

14.

Nielsen, L.E., Models for the permeability of filled polymer systems. Journal of
Macromolecular Science-Chemistry, 1967. 1(5): p. 929-942.

15.

Gain, O., et al., Gas barrier properties of poly(ε-caprolactone)/clay nanocomposites:
Influence of the morphology and polymer/clay interactions. Journal of Polymer Science
Part B: Polymer Physics, 2005. 43(2): p. 205-214.

16.

Priolo, M.A., et al., Super Gas Barrier of Transparent Polymer-Clay Multilayer Ultrathin
Films. Nano Letters, 2010. 10(12): p. 4970-4974.

17.

Grunlan, J.C., et al., Effect of clay concentration on the oxygen permeability and optical
properties of a modified poly(vinyl alcohol). Journal of Applied Polymer Science, 2004.
93(3): p. 1102-1109.

18.

Alves, V.D., N. Costa, and I.M. Coelhoso, Barrier properties of biodegradable composite
films based on kappa-carrageenan/pectin blends and mica flakes. Carbohydrate
Polymers, 2010. 79(2): p. 269-276.

19.

Sanchez-Garcia, M.D., L. Hilliou, and J.M. Lagaron, Nanobiocomposites of
Carrageenan, Zein, and Mica of Interest in Food Packaging and Coating Applications.
Journal of Agricultural and Food Chemistry, 2010. 58(11): p. 6884-6894.

20.

Yang, Q., et al., Transparent cellulose films with high gas barrier properties fabricated
from aqueous alkali/urea solutions. Biomacromolecules, 2011. 12(7): p. 2766-2771.

21.

Aulin, C., M. Gallstedt, and T. Lindstrom, Oxygen and oil barrier properties of
microfibrillated cellulose films and coatings. Cellulose, 2010. 17(3): p. 559-574.

22.

Huang, H.-D., et al., High barrier graphene oxide nanosheet/poly(vinyl alcohol)
nanocomposite films. Journal of Membrane Science, 2012. 409-410(0): p. 156-163.

23.

Gokhale, A.A., J. Lu, and I. Lee, Immobilization of cellulase on magnetoresponsive
graphene nano-supports. Journal of Molecular Catalysis B: Enzymatic, 2013. 90(0): p.
76-86.

24.

Kim, H., Y. Miura, and C.W. Macosko, Graphene/Polyurethane Nanocomposites for
Improved Gas Barrier and Electrical Conductivity. Chemistry of Materials, 2010. 22(11):
p. 3441-3450.

25.
26.

Geim, A.K., Graphene: Status and Prospects. Science, 2009. 324(5934): p. 1530-1534.
Iler, R.K., Multilayers of Colloidal Particles. Journal of Colloid and Interface Science,
1966. 21(6): p. 569-&.

27.

Decher, G. and J.D. Hong, Buildup of Ultrathin Multilayer Films by a Self-Assembly
Process .1. Consecutive Adsorption of Anionic and Cationic Bipolar Amphiphiles on
90

Charged Surfaces. Makromolekulare Chemie-Macromolecular Symposia, 1991. 46: p.
321-327.
28.

Decher, G. and J.D. Hong, Buildup of Ultrathin Multilayer Films by a Self-Assembly
Process .2. Consecutive Adsorption of Anionic and Cationic Bipolar Amphiphiles and
Polyelectrolytes on Charged Surfaces. Berichte Der Bunsen-Gesellschaft-Physical
Chemistry Chemical Physics, 1991. 95(11): p. 1430-1434.

29.

Lee, I., Molecular Self-Assembly: Smart Design of Surface and Interface via Secondary
Molecular Interactions. Langmuir, 2013. 29(8): p. 2476-2489.

30.

Hendricks, T.R., et al., Effects of catalyst introduction methods using PAMAM
dendrimers on selective electroless nickel deposition on polyelectrolyte multilayers.
Langmuir, 2007. 23(13): p. 7404-7410.

31.

Yu, A.M., et al., Mesoporous silica particles as templates for preparing enzyme-loaded
biocompatible microcapsules. Advanced Materials, 2005. 17(14): p. 1737-+.

32.

Lvov, Y. and F. Caruso, Biocolloids with ordered urease multilayer shells as enzymatic
reactors. Analytical Chemistry, 2001. 73(17): p. 4212-4217.

33.

Shan, W.Q., et al., Polyelectrolyte multilayer films as backflushable nanofiltration
membranes with tunable hydrophilicity and surface charge. Journal of Membrane
Science, 2010. 349(1-2): p. 268-278.

34.

Dotzauer, D.M., et al., Catalytic membranes prepared using layer-by-layer adsorption of
polyelectrolyte/metal nanoparticle films in porous supports. Nano Letters, 2006. 6(10): p.
2268-2272.

35.

Hassler, B.L., et al., Renewable dehydrogenase-based interfaces for bioelectronic
applications. Langmuir, 2007. 23(13): p. 7127-7133.

36.

Devesh, S., et al., Neuropathy Target Esterase Biosensor in , 2010: INTECH: Intelligent
and Biosensors edited by Vernon S. Somerset. 386.

37.

Lu, J., et al., Simple fabrication of a highly sensitive glucose biosensor using enzymes
immobilized in exfoliated graphite nanoplatelets Nafion membrane. Chemistry of
Materials, 2007. 19(25): p. 6240-6246.

38.

Lu, J., et al., Nanometal-decorated exfoliated graphite nanoplatelet based glucose
biosensors with high sensitivity and fast response. Acs Nano, 2008. 2(9): p. 1825-1832.

39.

Sambanthamoorthy, K., et al., Identification of a Novel Benzimidazole That Inhibits
Bacterial Biofilm Formation in a Broad-Spectrum Manner. Antimicrobial Agents and
Chemotherapy, 2011. 55(9): p. 4369-4378.

91

40.

Mehrotra, S., et al., Time Controlled Protein Release from Layer by-Layer Assembled
Multilayer Functionalized Agarose Hydrogels (vol 20, pg 247, 2010). Advanced
Functional Materials, 2010. 20(6): p. 878-878.

41.

Mehrotra, S., I. Lee, and C. Chan, Multilayer mediated forward and patterned siRNA
transfection using linear-PEI at extended N/P ratios. Acta Biomaterialia, 2009. 5(5): p.
1474-1488.

42.

Mehrotra, S., et al., Time Controlled Release of Arabinofuranosylcytosine (Ara-C) from
Agarose Hydrogels using Layer-by-Layer Assembly: An In Vitro Study. Journal of
Biomaterials Science-Polymer Edition, 2012. 23(1-4): p. 439-463.

43.

Clark, S.L., M.F. Montague, and P.T. Hammond, Ionic effects of sodium chloride on the
templated deposition of polyelectrolytes using layer-by-layer ionic assembly.
Macromolecules, 1997. 30(23): p. 7237-7244.

44.

Yang, S.Y. and M.F. Rubner, Micropatterning of polymer thin films with pH-sensitive
and cross-linkable hydrogen-bonded polyelectrolyte multilayers. Journal of the American
Chemical Society, 2002. 124(10): p. 2100-2101.

45.

Hendricks, T.R., W. Wang, and I. Lee, Buckling in nanomechanical films. Soft Matter,
2010. 6(16): p. 3701-3706.

46.

Hendricks, T.R. and I. Lee, Wrinkle-free nanomechanical film: Control and prevention of
polymer film buckling. Nano Letters, 2007. 7(2): p. 372-379.

47.

Biswas, S., H. Fukushima, and L.T. Drzal, Mechanical and electrical property
enhancement in exfoliated graphene nanoplatelet/liquid crystalline polymer
nanocomposites. Composites Part a-Applied Science and Manufacturing, 2011. 42(4): p.
371-375.

48.

Fukushima, H., Graphite nanoreinforcements in polymer nanocomposites,Department of
Chemical Engineering and Materials Science, Michigan State University, East Lansing,
MI, USA, 2003.

49.

Wu, H. and L.T. Drzal, Graphene nanoplatelet paper as a light-weight composite with
excellent electrical and thermal conductivity and good gas barrier properties. Carbon,
2012. 50(3): p. 1135-1145.

50.

Jiang, X. and L.T. Drzal, Reduction in percolation threshold of injection molded highdensity polyethylene/exfoliated graphene nanoplatelets composites by solid state ball
milling and solid state shear pulverization. Journal of Applied Polymer Science, 2012.
124(1): p. 525-535.

92

51.

Hendricks, T.R., et al., Intact pattern transfer of conductive exfoliated graphite
nanoplatelet composite films to polyelectrolyte multilayer platforms. Advanced Materials,
2008. 20(10): p. 2008-+.

52.

Lu, J., et al., Stable aqueous suspension and self-assembly of graphite nanoplatelets
coated with various polyelectrolytes. Journal of Nanomaterials, 2010. 2010: p. 2.

53.

Shiratori, S.S. and M.F. Rubner, pH-dependent thickness behavior of sequentially
adsorbed layers of weak polyelectrolytes. Macromolecules, 2000. 33(11): p. 4213-4219.

54.

Kim, H.M., J.K. Lee, and H.S. Lee, Transparent and high gas barrier films based on
poly(vinyl alcohol)/graphene oxide composites. Thin Solid Films, 2011. 519(22): p.
7766-7771.

55.

Yu, L., et al., A graphene oxide oxygen barrier film deposited via a self-assembly coating
method. Synthetic Metals, 2012. 162(7-8): p. 710-714.

56.

Nagel, C., et al., Free volume and transport properties in highly selective polymer
membranes. Macromolecules, 2002. 35(6): p. 2071-2077.

57.

Yang, Y.H., et al., Super Gas Barrier of All-Polymer Multilayer Thin Films.
Macromolecules, 2011. 44(6): p. 1450-1459.

58.

http://xgsciences.com/wp-content/uploads/2012/09/About-xGnP.pdf.

59.

Kim, S., I. Do, and L.T. Drzal, Multifunctional xGnP/LLDPE Nanocomposites Prepared
by Solution Compounding Using Various Screw Rotating Systems. Macromolecular
Materials and Engineering, 2009. 294(3): p. 196-205.

60.

Schneider, M., M. Brinkmann, and H. Mohwald, Adsorption of polyethylenimine on
graphite: An atomic force microscopy study. Macromolecules, 2003. 36(25): p. 95109518.

61.

Willner, I., et al., Photoinduced Electron-Transfer Processes Using Organized RedoxFunctionalized Bipyridinium Polyethylenimine Tio2 Colloids and Particulate Assemblies.
Journal of Physical Chemistry, 1993. 97(28): p. 7264-7271.

62.

Bequet, S., et al., New composite membrane for water softening. Desalination, 2000.
131(1-3): p. 299-305.

63.

Quinn, J.F., et al., Next generation, sequentially assembled ultrathin films: beyond
electrostatics. Chemical Society Reviews, 2007. 36(5): p. 707-718.

64.

Lavalle, P., et al., Comparison of the structure of polyelectrolyte multilayer films
exhibiting a linear and an exponential growth regime: An in situ atomic force microscopy
study. Macromolecules, 2002. 35(11): p. 4458-4465.
93

65.

DeLongchamp, D.M. and P.T. Hammond, Highly ion conductive poly(ethylene oxide)based solid polymer electrolytes from hydrogen bonding layer-by-layer assembly.
Langmuir, 2004. 20(13): p. 5403-5411.

66.

Picart, C., et al., Buildup mechanism for poly (L-lysine)/hyaluronic acid films onto a solid
surface. Langmuir, 2001. 17(23): p. 7414-7424.

67.

Laachachi, A., et al., Diffusion of Polyphosphates into (Poly(allylamine)montmorillonite) Multilayer Films: Flame Retardant-Intumescent Films with Improved
Oxygen Barrier. Langmuir, 2011. 27(22): p. 13879-13887.

68.

He, J.-A., et al., Oriented Bacteriorhodopsin/Polycation Multilayers by Electrostatic
Layer-by-Layer Assembly. Langmuir, 1998. 14(7): p. 1674-1679.

69.

Chatham, H., Oxygen diffusion barrier properties of transparent oxide coatings on
polymeric substrates. Surface and Coatings Technology, 1996. 78(1): p. 1-9.

70.

Zhang, Z.B., I.J. Britt, and M.A. Tung, Permeation of oxygen and water vapor through
EVOH films as influenced by relative humidity. Journal of Applied Polymer Science,
2001. 82(8): p. 1866-1872.

71.

Kalaitzidou, K., H. Fukushima, and L.T. Drzal, Multifunctional polypropylene
composites produced by incorporation of exfoliated graphite nanoplatelets. Carbon,
2007. 45(7): p. 1446-1452.

72.

Jiang, J., et al., Oxygen Barrier Coating Deposited by Novel Plasma-enhanced Chemical
Vapor Deposition. Journal of Applied Polymer Science, 2010. 115(5): p. 2767-2772.

73.

Fielding-Russell, G.S. and P.S. Pillai, A study of the crystallization of polyethylene
terephthalate using differential scanning calorimetry and X-ray techniques. Die
Makromolekulare Chemie, 1970. 135(1): p. 263-274.

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Chapter 5: Amperometric detection and quantification of nitrate ions using a
highly sensitive nanostructured PEDOT biosensor
(This chapter is submitted to a peer-reviewed journal and is currently undergoing revisions)

5.0 Abstract
In this work, we developed a novel one-step electropolymerized nitrate biosensor with detection
limit as low as 0.1 ppm. In our work, we exploit the unidirectional flow of electrical signal in 1D
conductive polyethylenedioxythiophene (PEDOT) nanowires to produce a fast acting, highly
sensitive biosensor. Nitrate reductase, an enzyme responsible for conversion of nitrate species to
nitrite is entrapped during the electropolymerization of PEDOT. Our work shows that the
polymerization conditions such as polymerization time and potential can play a major role in the
performance of the sensor. The sensor developed here has high resilience against interfering
species. Based on kinetic and operational parameters, we show that the nitrate biosensor
developed in this study shows impressive sensing and quantification capabilities as compared to
other electrochemical sensors.

5.1 Introduction
Human activities have contributed to as much as 30% increase in nitrogen and nitrogen based
compounds in the environment as compared to natural modes of fixation[1]. The increasing use
of fertilizers with nitrogen precursors and automobiles emitting nitrogen oxides have put
immense burden on the existing nitrogen cycle. The inability of natural sources to recycle the
excess nitrogen based materials makes nitrogen a major contributor to eutrophication. A
cascading set of events due to elevated nitrogen levels leading to ‘algal bloom’, formation of
‘dead zones’ because of depletion of dissolved oxygen and consequently loss of marine

95

biodiversity is well-documented[2, 3]. Various natural and biological processes are responsible
for transformation of nitrogen based compounds into nitrate. The presence of nitrate in water
supplies presents major health risks. To limit the consumption of nitrate, the U.S. Environmental
Protection Agency (EPA) and the World Health Organization (WHO) set the maximum limit of
nitrate in drinking water supplies as 10 ppm nitrate-nitrogen (equivalent to 45 ppm nitrate) and
11 ppm nitrate-nitrogen (same as 50 ppm nitrate), respectively[4]. Numerous studies have been
conducted in the past regarding determination of nitrate or nitrogen based compounds in water
supplies[5-7]. Using the data collected by the U.S. Geological Survey National Water-Quality
Assessment Program, Rupert[8] studied the general trend of nitrate concentration in groundwater supplies by evaluating data from a 24 well-network spread all over the US. Close to 30%
of the well network showed an increase in nitrate concentrations. Half of these wells recorded
nitrate concentration above the maximum limit prescribed by the regulatory agencies.
One of the most prominent ill effects of increased nitrate consumption in humans is the
low oxygen carrying capacity of the blood. The conversion of ingested nitrate to nitrite followed
by incorporation of nitrite with hemoglobin leads to the formation of a complex called
methemoglobin (MetHb)[9]. In infants there is greater incidence of nitrite because of the lower
gastric acidity[10]. Cellular anoxia is a likely consequence of lower oxygen transport throughout
the body. This disorder is commonly referred to as methemoglobinemia. Besides this, nitrate is
also responsible for the formation of several N-nitroso compounds (NOC) which in turn increase
the risk of cancer. Studies regarding the impact of nitrate uptake on occurrence of gastric and
abdominal cancers have been carried out[11-13]. Consumption of nitrate is also linked to growth
of cancerous lesions in the human thyroid gland[14]. Nitrate is also suspected to disrupt the

96

endocrine functioning of the thyroid gland by blocking the iodide symporter independently or in
conjugation with potent inhibitors such as perchlorate and thiocyanate[15].
Previous methods for detection and quantification of anions include ion chromatography
(IC), complexation electrospray mass spectrometry (cESI-MS) and Raman spectroscopy[16-18].
However, these methods require extensive instrumentation and are not particularly attractive for
instant field tests. Biosensors on the other hand have demonstrated a great potential for the
development of simple portable systems capable of detection and quantification even at minute
concentrations. Several examples of biosensors capable of accurate and sensitive analysis are
found in literature[19-21]. Detection of nitrate using an enzymatic biosensor has been
extensively studied in the past few decades. Nitrate reductase, an enzyme capable of facilitating
reduction of nitrate species has been used as the biorecognition element for the fabrication of
nitrate detecting biosensor. This enzyme also plays a significant role in the ‘denitrification cycle’
by participating in the nitrogen acquisition mechanism in higher plants[22]. One of the earliest
designs for a nitrate biosensor was proposed by Guilbault and co-workers[23]. Low sensitivity
and dependence on co-factors reduced the practical applicability of this device. Since then efforts
have been dedicated to improve the sensitivity, stability and ruggedness of the biosensor to make
it suitable for commercial application. Newer designs include use of metal nanoparticles[24],
colloidal clay[25], redox polymers[26, 27] and screen printing technology[28] to improve the
performance of the biosensors. The integration of these components helps to improve the
microenvironment in which the immobilized enzyme operates and maximizes the output
intensity facilitating easy detection of nitrate. However, unlike the biosensor described in this
study where we take the full advantage of the nanoscale dimensions by immobilizing the nitrate
reductase in upright PEDOT nanoarrays, most of the prior art relies on the incorporation of

97

nitrate reductase in thin films made from polymeric- nanomaterial matrix. Some of these films
may not always retain their nanoscale dimensions during the growth process. The enzyme is
likely to be buried deep inside the film blocking its accessibility to the substrate limiting the
performance of the sensor.
Conductive polymer nanowire/ nanotubes are attractive candidates to serve as supports
for enzyme immobilization. Nanostructures made from conductive polymers (CP) are known to
exhibit much higher conductivities than bulk materials. In some cases it can be as high as 15
orders of magnitude depending on the parameters such as dopant ions or monomer/dopant
ratio[29]. Besides the substrate-enzyme accessibility issues observed in thin films as pointed out
above, flat 2D films are also prone to lateral current shunting, reducing their signal intensities.
On the other hand, nanotubes/nanowires are the smallest dimensions that allow efficient
transport of electrons thus enabling detection of single molecules[29]. The use of the enzymepolymer nanowire matrix as shown in this work is therefore expected to show enhanced
electrical performance. This we believe has translated into superior LOD and instantaneous
response (2-5 seconds) making the sensor one of the frontrunners in the class of electrochemical
biosensors for detection of nitrate. Among the various techniques used to grow nanotubular/wire
structure, the templated route is a convenient approach to growth polymeric nanostructures of
well-defined morphology. Typically, template assisted polymerization enables the growth of
polymer nanostructures with enhanced molecular and supramolecular morphology[30]. Use of
hard templates such as track-etched polycarbonate (PC) and nanoporous anodic alumina
membrane is very common[31]. Templates allow highly oriented growth of polymers[32]. The
morphology of the polymer nanostructures is dictated by the shape and size of the pore in the
hard template and hence polymeric nanostructures of desired aspect ratio can be easily grown in

98

the pores. In our work, regulated growth of polymer/enzyme nanowire structures enabled precise
control over growth of nanostructures enabling improved operational stability and superior signal
to noise ratio. Using the standard CMOS processes and techniques such as dielectrophoretic
assembly, large arrays of nanowires made from conducting polymers can be easily aligned on
silicon chips. This readily available technology can be easily extended to our biosensor
application to produce commercial sensors in the near future. [33].
Compared to other heterocycles, PEDOT is a conducting polymer that retains its
conductivity and structural stability over a wide range of temperature[34]. The oxidation of
EDOT produces a transparent conductive polymer with high electrochemical stability and low
band-gap[35, 36]. EDOT also affords easy functionalization with redox active groups to improve
the overall solubility and processibility[37]. Because of the many advantages, PEDOT can serve
as an attractive candidate for applications such as optoelectronics[38, 39], electrochromics[40,
41], and supercapacitators[42, 43].
In this work, using a fast one-step electropolymerization process, PEDOT/enzyme
nanowires were grown in the pores of PC membrane. The insolubility of EDOT in aqueous
solution and high polymerization potential were overcome by adding sufficient quantity of
surfactant. This study shows that PEDOT serves an excellent matrix to entrap nitrate reductase.
PEDOT/enzyme biosensor was fabricated at specific process conditions such as applied potential
and polymerization times. By meticulously optimizing the process conditions we establish a
strong link between the performance of the biosensor and the morphology of the
PEDOT/enzyme nanowires.

As compared to other electrochemical nitrate reductase based

sensors in literature (comparison shown in subsequent section), the electropolymerized
nanobiosensor developed in this study shows several superior attributes: low limit of detection

99

(LOD) of about 0.108 ppm, fast fabrication (about 300 seconds), a very quick response time and
low background noise. The sensor developed in this study also showed a significant resilience
against interfering species. The kinetic parameters of nitrate reduction were estimated from the
initial rate data and compared with similar data obtained by replacing nitrate species with other
interfering substrates. Substrate binding affinity with nitrate was estimated to be several orders of
magnitude higher than other interfering ions such as perchlorate and carbonate.

5.2 Experimental
5.2.1 Materials
Sodium chloride (NaCl), sodium dodecyl sulfate (SDS), methyl viologen, potassium perchlorate,
potassium nitrate, sodium carbonate and ethylene dioxythiophene (EDOT) were purchased from
Sigma Aldrich. Conductive copper tape (adhesive on one side) was purchased from McmasterCarr, GA. Nucleopore track-etched polycarbonate (PC) membrane with pore size 200 nm was
purchased from Fisher Scientific. Nitrate reductase (Y-NaR) was purchased from Nitrate
elimination Co., MI.

5.2.2 Preparation of template assisted PEDOT/enzyme nanowire electrode
Templated growth of PEDOT nanostructures is described elsewhere[42, 44] [45] In our work, we
extend the templated growth of PEDOT nanostructures to coimmobilize nitrate reductase using a
one-step electropolymerization process. Based on our knowledge, this is the first attempt to grow
PEDOT/enzyme nanowire structures using single step process while still retaining the integrity
of the 1D nanowire structure. Unlike other biosensor fabrication techniques where a film of the
conducting polymer and enzyme is co-immobilized, in this work we retain the upright 1D

100

nanowire structure synthesized in the PC membrane to take full advantage of the electrical
conductivity of 1D nanowire structure as explained in the previous section.
The one step electropolymerization of PEDOT/nitrate reductase is summarized briefly as
given below. Commercially available 13 mm polycarbonate membrane was sputter-coated with a
15 nm thick gold layer using a Denton Vacuum sputter coater. A conductive copper tape was
attached to the other side of the PC membrane. This was covered with a circular insulating tape.
The sputter-coated side of the membrane was covered with a partial insulating disc such that the
effective electroactive area available for polymerization is about 0.2 cm2. The commercial nitrate
reductase is available in freeze-dried form. The nitrate reductase is added to the
electropolymerization bath consisting of 0.05 M EDOT, 0.07 M SDS and 0.1 M NaCl. About 1
ml of the electropolymerization bath solution is added to one vial of the freeze-dried enzyme
sample (protein concentration about 120-125 µg/ml ) and the mixture is allowed to equilibrate at
room temperature for at least 15 minutes.
Electrochemical polymerization of EDOT/Enzyme complex is carried out using a 3electrode system. The working electrode comprised on the Au- coated PC membrane as prepared
earlier. Saturated calomel electrode (SCE in saturated KCl) served as the reference electrode
whereas a thin platinum wire was used as a counter electrode. Electrochemical polymerization
was carried out at desired oxidation potential and time. The polymerization of EDOT/enzyme in
the pores of the PC membrane was confirmed by the formation of bluish-black composite on the
exposed portion of the membrane. To check the electroactivity of the working electrode, the
working electrode was immersed in 10 ml, 10 mM stirred phosphate buffer saline (PBS).1 mM
MV was added as a redox mediator to the PBS buffer. Prior to immersion of the electrode, the
buffer solution was deoxygenated by bubbling argon for 30 minutes.

101

5.2.3 Characterization of PEDOT/enzyme nanowires
The scanning electron microscopy (SEM) investigation for high-resolution imaging was carried
out using a JEOL 7500F with field emission. Transmission electron microscopy (TEM) studies
were carried out using a JEOL 100CX. After the electrochemical polymerization of
PEDOT/enzyme nanostructures, the PC membrane was dissolved using methylene chloride and
mild sonication. A few drops of the solution were put on a lacey carbon coated TEM grid and
dried overnight. The grids were then observed under SEM and TEM.

5.3 Results and Discussion
The growth of conductive polymers using a templated approach offers attractive opportunities to
fabricate electronic interfaces of desired aspect ratio. In this study, using a one-step
electropolymerization process we developed a highly sensitive PEDOT based nanostructured
biosensor with a LOD close to 108 ppb (0.108 ppm) nitrate. A schematic diagram depicting the
growth of PEDOT in a polycarbonate (PC) membrane is shown in Figure 5.1

102

Figure 5.1 Electrochemical polymerization of EDOT using a templated approach. (a) Formation
of polymer nanotubes (b) Formation of polymer nanowires

Along with the monomer, the electropolymerization bath consists of SDS (surfactant),
supporting electrolyte (NaCl) and nitrate reductase. Though two different types of structures
(hollow nanotubular (a) and filled nanowire (b) structure) are feasible depending on the process
conditions, in this paper we will be focusing on the growth of a filled nanowire structure. A
completely filled nanowire structure is a result of high concentration of monomer and relatively
103

longer polymerization time. High concentration of monomer ensures steady supply of EDOT to
the active sites in the pores of the PC membrane. A long polymerization time allows the polymer
to grow along the wall of the pore and then fill the pore cavity resulting in a well-formed
nanowire structure. Detailed discussion regarding the mechanism of growth of PEDOT
nanotubes vs. nanowire structure is available elsewhere[42, 44, 46].
Under oxidizing potential, most conducting polymers undergo p-doping followed by
entry of counter-ions from the supporting electrolyte solution to maintain electroneutrality[47].
In this case, the Cl- ions from the supporting electrolyte solution diffuse into the PEDOT
structure during the electropolymerization process. To overcome the issue of EDOT insolubility
in aqueous medium, sodium dodecyl sulfate (SDS) is added to the supporting electrolyte
solution. The anionic DS- ions of SDS bind with the EDOT+ radical to form a pseudo-complex
which is stable in aqueous medium[48]. The hydrophilic ends of SDS also wet the electrode
surface and allow the subsequent deposition of PEDOT. The addition of SDS also serves
another important purpose. It lowers the oxidation potential at which EDOT polymerizes [48,
49]. This is particularly important in biological systems which are known to deteriorate when
subjected to higher oxidation potential. Unlike electropolymerization of some conducting
polymers such as polyaniline[50] and polypyrrole[51] that need an acidic environment, the
electropolymerization of PEDOT aided by addition of SDS can be carried out in DI water
without pH adjustment. Acidic environment usually spurs the preferential complexation of H 3O+
ions which can reduce the nucleophilicity of water [52]. The use of surfactants for synthesis of
PEDOT can avoid this problem. The use of neutral aqueous medium can largely preserve the
biomolecule structure and function. Once synthesized, PEDOT largely retains its stability and

104

conductivity as opposed to polypyrrole which is likely to develop defects due to multiple , 
and ,-linkages [36].
The addition of nitrate reductase to the electropolymerization bath solution and the
subsequent one step electrochemical synthesis of polymer entraps the enzyme within the PEDOT
nanowires grown in the pores of the PC membrane. With the isoelectric point of nitrate reductase
close to 6.1, the enzyme attains a weak negative charge when added to the neutral aqueous
electrochemical bath solution. Weak electrostatic interactions are likely to exist between the
cationic EDOT+ radical and the enzyme. The hybrid electrochemical adsorptive process
involving electrophoretic migration of charged species located in the close vicinity of the
bioelectronic interface leads to the deposition of nitrate reductase in amounts exceeding that
obtained by regular adsorption [47]. Other advantages of one step electrochemical adsorptive
process include precise control over the growth of the PEDOT/enzyme nanostructure by
monitoring the charge build-up during the process, limited structural damage to the enzyme due
to lack of harsh chemical reactions and ability to electrogenerate the PEDOT/enzyme coating on
different types of electrodes with complex geometry [53].

5.3.1 Anodic polymerization of EDOT
Oxidation onset potential (Eox

onset)

of EDOT depends on the components of the

electropolymerization bath solution more particularly the concentration of the surfactant used for
polymerization[52, 54]. The Eox

onset

was determined from linear sweep voltametry curves as

shown in Figure 5.2. The magnitude of current density is plotted as a function of potential. At
lower potential, EDOT polymerization is mostly subdued. However at a potential of 0.82V, the
current starts rising and a slight bulge-shaped profile can been seen. With further increase in the
applied potential, the current starts increasing rapidly as more and more EDOT starts getting
105

polymerized. The LSV profile for EDOT polymerized in presence of 0.07 M SDS and 0.1 M
NaCl is indicated by curve a. A qualitative confirmation of the participation of nitrate reductase
in the EDOT polymerization process can be obtained from curve b. Curve b shows the LSV
profile when nitrate reductase was added to the electropolymerization bath solution. The addition
of enzyme delays the onset polymerization of EDOT. E ox onset for this curve is close to 0.90-0.92
V. Also the magnitude of the current density of the EDOT polymerization in presence of nitrate
reductase is much lower. This could be because the EDOT + cations can form weak electrostatic
complexes with negatively charged nitrate reductase. The polymerization of EDOT +/nitrate
reductase (in presence of DS - ions) can differ from traditional EDOT +/DS- complexes and the
same needs to be probed in detail using more quantitative tools.

Figure 5.2 Linear sweep voltammetry (LSV) profile of EDOT polymerization (a) in absence of
enzyme (b) in presence of enzyme.

5.3.2 Chronoamperometric growth of PEDOT/enzyme nanowires
Chronoamperometry is a potentiostatic technique that has been frequently used for oxidative
growth of conducting polymers[55, 56]. Unlike cyclic voltammetry, chronoamperometry allows
the polymerization to occur at user defined fixed potential thus preventing the system from
106

reaching an overoxidation state. The one step electropolymerization of EDOT/nitrate reductase
was carried out at three different potentials namely 0.85 V, 1.1 V and 1.4 V respectively. The
electropolymerization time was fixed at 300 seconds in each case. For comparison, the electrodes
were subjected to EDOT polymerization in absence of nitrate reductase using the same
conditions as before. Six systems were subjected to polymerization and the variation of current
density w.r.t time is shown in Figure 5.3. All the six cases show a common trend.

Figure 5.3 Chronoamperometric plot of current density as a function of electropolymerization
time. Inclusion of enzyme reduces the creation of active sites for polymerization evidenced by
lower current density during the initial phase of polymerization.
The chronoamperometric graphs can be divided into three separate regions. The first region lasts
for a few seconds but shows a sharp increase in the current. In this region, active sites within the
pores of the membrane (along the walls and on the gold electrode) where the EDOT monomer is
likely to polymerize are created. Zeng and co-workers[57] found similar behavior when choline
oxidase was added to the electropolymerization bath solution consisting of EDOT monomer.
Region II shows a gentle decrease in the current as the system tries to stabilize. It is reported that
it is in this region that the diffusion and deposition of monomer units occurs [57]. In the process,
107

the active sites within the pores of the membrane are used up. Region III shows the current
reaching a steady plateau value indicating no further creation of active sites. Linear growth of the
polymer/enzyme nanowire is expected to occur to occur in this region. The addition of enzyme
to the electropolymerization bath solution is expected to slow down the polymerization process.
As seen earlier, the slight increase in the E ox onset potential confirms this. Addition of enzyme
reduces the number of active sites available for polymerization. Hence the three cases that
include incorporation of the enzyme in the PEDOT matrix show lower current in region I.

5.3.3 Morphology of PEDOT /enzyme nanostructures
In the previous section, we observed from the chronoamperometric plots that application of
varied potentials can lead to differential creation of active sites. Initially at low potentials, the
degree of polymerization is very low. However as the potential crosses the E ox onset , the degree of
polymerization increases and this is represented by increase in current density. Since 0.85 V is
less than Eox onset (for enzyme incorporated system), the driving force for creation of active sites in
the pores of the membrane is subdued. The morphology of PEDOT/enzyme nanowires formed at
this potential is therefore not very distinct. At 0.85 V, the nanowires are short, clustered with loss
in form especially at the ends. This is evident from the SEM and TEM images of
PEDOT/enzyme nanowires synthesized at 0.85 V shown in Figures 5.4 (a) and (b) respectively.
However, as the potential is raised beyond the E ox onset, the nanowires appear more distinct and
their morphology at the ends is well-formed. The SEM and TEM images shown in Figure 5.4 (c)
and (d) confirm this. As the potential is raised further to 1.4 mV, more and more polymer gets
deposited in the pores of the membrane. As seen from Figure 5.4 (e) and (f) , irregularly shaped
nanowires are commonly produced at this potential. At higher potential, overoxidation of
PEDOT becomes the dominating factor. In case of most polythiophenes, increasing oxidation
108

potential results in ‘overoxidation’ that results in rapid degradation and irreversible loss in
polymer conductivity[58]. Du and Wang[59] have established that effects of over-oxidation
cannot be neglected when the electropolymerization potential exceeds 1.1 V. Another possible
impediment in carrying out EDOT polymerization at higher potential is the possible breakdown
of water. The decomposition of water at 1.23 V allows only a limited potential window for the
polymerization of EDOT [60, 61].

109

Figure 5.4 Microscopy images of PEDOT nanowires at different oxidation potential. SEM
images of nanowires polymerized at (a) 0.85 V (c) 1.1 V (e) 1.4 V. Corresponding TEM images
(b) 0.85 V (d) 1.1 V (f) 1.4 V.
110

Digital images of the polymer suspension in dichloromethane following the dissolution for PC
membrane (for preparation of SEM/TEM samples) shown in Figure 5.5 give us a qualitative idea
of the degree of polymerization at different potentials. PEDOT has a characteristic bluish-black
color. Upon dissolution of the PC membrane, the polymer/enzyme nanowires get dispersed in the
organic solvent. At 0.85 V, the extent of polymerization is low, so very little polymer is formed.
Hence the solution is mostly clear. The formation of the polymer at higher potential namely 1.1
V and 1.4 V is confirmed by the characteristic bluish-black color of PEDOT as seen from Figure
5.5. Upon visual inspection, the bluish-black color of the PEDOT solution obtained at 1.4 V
appears to be more intense than the one at 1.1 V.

Figure 5.5 Digital images of PEDOT in dichloromethane solution. EDOT polymerized at 0.85 V
yields clear solution indicative of low degree of polymerization. At 1.1 and 1.4 V, bluish-black
PEDOT is formed.
5.3.4 Amperometric detection of nitrate ions
The electropolymerized biosensor developed in the previous section was subjected to addition of
potassium nitrate. With each successive addition of potassium nitrate, the cumulative nitrate
concentration was evaluated. As mentioned before, nitrate reductase is an enzyme that operates
111

under anaerobic conditions. Therefore prior to the test, the PBS buffer was thoroughly purged
with argon gas for about 30 minutes. Besides argon purging, other methods include using oxygen
scavengers such as sodium sulfite [62]. However, we avoided using external scavengers since
their effect on the electropolymerized PEDOT sensor is relatively unknown. However in future,
use of oxygen scavengers prior to the amperometric test seems to be a viable option for carrying
out instant field tests. 1mM MV was added to the PBS buffer prior to argon purging. Methyl
viologen is a well-known redox mediator that can aid shuttling of electrons between the
bioelectronics interface and the active site of the enzyme. The addition of MV proves to be
useful particularly in case of nitrate reductase based biosensors because the active site catalyzing
the reduction of nitrate to nitrite is buried deep within the enzyme[22, 27]. The presence of MV
in the buffer solution helps in efficient transmission of electrical signal enhancing the chances of
achieving a quick response.
Biosensors fabricated at three different potentials namely 0.85 V, 1.1 V and 1.4 V were
subjected to amperometric test. The sensor polymerized at 0.85 V comprises of polymer/enzyme
nanowires with morphological defects (especially at the ends). The polymer/emzymes nanowires
are also short and mostly buried within the pores of the membrane. The nitrate ions suffer from
accessibility issues since they need to diffuse through the pores of the membrane and reach the
polymer/enzyme nanowires. Hence the biosensor developed at this potential fails to give a
measurable response. However, detection of nitrate ions is still possible. This is seen from Figure
5.6. Addition of nitrate leads to spike in the current. However, the poorly formed nanostructures
are unable to sustain the increase in the current. Within a few seconds the current starts tailing
off. The inability of the biosensor to achieve steady state current after successive addition of
nitrate ions makes it difficult to quantify the analyte content in the solution.

112

Figure 5.6. Amperometric profile of nitrate biosensor fabricated at polymerization potential of
0.85 V. Detection of nitrate species is possible but quantification information is hard to establish.
Similarly, the sensor fabricated using a polymerization potential of 1.4 V suffers from
quantification issues. (See Figure 5.7)

Figure 5.7 Amperometric profile of nitrate biosensor fabricated at polymerization potential of 1.4
V. Detection of nitrate species is possible but no quantification information is hard to establish.

113

However, unlike the poor/incomplete oxidation of EDOT as observed at 0.85 V, the
polymerization at 1.4 V suffers from overoxidation. This means that though the polymer/enzyme
nanowires are formed in the pores of the membrane, a loss of conductivity due to overoxidation
can result in poor signal transmission. Also higher potential can also result in some nanowires
outgrowing the pores. The outgrown nanowires can combine and form polymer/enzyme thin film
on the surface of the membrane. This can damage the advantages afforded by 1D nanowire
structure. We found higher incidence of nanowires growing up to 9 microns in length at this
polymerization condition. (See Figure 5.8). Such long nanowires were not observed routinely
observed at other conditions.

(a)
Figure 5.8. TEM micrographs of PEDOT nanowires fabricated at polymerization potential of 1.4
V. (a) length of nanowire close to 6.5 microns (b) length of nanowire close to 9 microns. Growth
is irregular.
114

Figure 5.8 (cont’d)

(b)

Another configuration of PEDOT/enzyme biosensor was tested by polymerization at 1.1
V. The selection of 1.1 V was made keeping in mind that the E ox onset for the enzyme incorporated
system is close to 0.9-0.92 V. Fabrication of the biosensor at 1.1 V should eliminate
morphological defects resulting from incomplete polymerization and/or overoxidation. Also 1.1
V is also lower than the theoretical potential at which water undergoes decomposition.

The

biosensor showed improved performance when fabricated at this condition. The step-wise
increase in current (see Figure 5.9a) occurs as a result of successive addition of KNO 3 solution.
This facilitates easy quantification of the added analyte. Since 1.1 V also lies in the small
potential window where the oxidation of EDOT yields a highly conductive polymer, we
observed a quick response of the biosensor to the addition of nitrate solution. The sensor records
115

a rise in current with 2-5 seconds on addition of the analyte solution. The sensor also enjoys very
low detection limit (LOD: 108 ppb). As seen from Figure 5.9b, the sensor shows a linear
response to nitrate addition in the range 200 ppb-1100 ppb.

(a)
Figure 5.9 (a) Amperometric profile of nitrate biosensor fabricated at 1.1 V, 300 sec. in presence
of nitrate species. Inset image shows step-wise increase in current after addition of 200 ppb
nitrate after every 50 seconds (b) Linear range of operation of the sensor.

116

Figure 5.9 (cont’d)

(b)
In order to understand the effect of the polymerization time on the performance of the
biosensor, we fabricated a sensor using a polymerization potential of 0.95 V (just enough to
ensure optimum polymerization of EDOT) and polymerization times of 300 seconds and 600
seconds respectively. At 300 sec, just like before detection of nitrate species was observed but
quantification trend was hard to establish. (See Figure 5.10).

117

Figure 5.10. Amperometric profile of nitrate biosensor fabricated at polymerization potential of
0.95 V (300 sec.). The nitrate species are detected but quantitative trend difficult to establish.
But on increasing the polymerization time to 600 sec, a step-wise increase in current with
fast response was observed (See Figure 5.11a). The biosensor fabricated at this condition shows
a LOD of around 363 ppb. The linear range is from 550 ppb-1600 ppb (Refer Figure 5.11b).
Increasing the polymerization time to 600 seconds allows higher degree of EDOT
polymerization. Also longer time allows more enzyme to get entrapped within the PEDOT
matrix thus improving the response of the sensor. However SNR for this sensor is low. Thus by
increasing the polymerization time, a quantifiable set of data can be obtained. However, high
levels of noise in the system can impede the practical applicability of the sensor fabricated at this
condition.

118

(a)
Figure 5.11 (a) Amperometric profile of nitrate biosensor fabricated at 0.95 V, 600 sec. in
presence of nitrate species. Inset image shows step-wise increase in current after addition of 200
ppb nitrate after every 50 seconds (b) Linear range of operation of the sensor.

119

Figure 5.11 (cont’d)

(b)
5.3.5 Effect of other reducible substrates on the nitrate reductase biosensor
Presence of other reducible substrates is likely to affect the performance of the nitrate reductase
biosensor. Like all other analytical methods, the nitrate reductase based biosensors are
susceptible to interferences due to substrates that share a similar structural affinity. The
specificity of nitrate reductase enzyme plays an important role in minimizing the effects of these
interferences. The thermodynamics of perchlorate reduction closely matches that of nitrate
reduction and this is evident by comparing the Gibb’s free energy of both systems (ΔG 0= -801
kJ/mol acetate for perchlorate reduction as compared to ΔG 0=-792 kJ/mol acetate for nitrate
reduction) [63]. This makes it a potent interfering ion. Carbonate species (obtained by replacing
N in nitrate with C) normally associated with hard water is also a known interfering material
largely due to the structural similarity [64].

The kinetic parameter kcat (enzyme turnover

number) commonly used in determination of substrate specificity cannot be accurately estimated

120

due to the lack of information regarding the extent of enzyme loading during the polymerization
process. This is one drawback associated with using one-step enzyme/polymer coimmobilization route and has been duly noted in literature [53]. However other kinetic
parameters relevant to enzyme kinetics can be obtained by from the Lineweaver Burk plot.
In this work, we carried out studies to ascertain the affinity of the enzymatic biosensor
towards the substrates containing nitrate, perchlorate and carbonate species. The biosensor was
fabricated at 1.1 V and effect of the above substrates was estimated by adding potassium nitrate,
potassium perchlorate and sodium carbonate separately. Refer Figures 5.12 (a) and 5.13 (a) for
the amperometric profile obtained after adding perchlorate and carbonate substrates.

(a)
Figure 5.12. (a) Amperometric profile of nitrate biosensor fabricated at 1.1 V, 300 sec. in
presence of perchlorate species. Inset image shows step-wise increase in current after addition of
2.5 ppm perchlorate after every 50 seconds (b) Linear range of operation of the sensor.

121

Figure 5.12 (cont’d)

(b)

(a)
Figure 5.13. (a) Amperometric profile of nitrate biosensor fabricated at 1.1 V, 300 sec. in
presence of carbonate species. Inset image shows step-wise increase in current after addition of
25 ppm nitrate after every 50 seconds (b) Linear range of operation of the sensor.
122

Figure 5.13 (cont’d)

(b)
Adding lower concentrations of perchlorate and carbonate did not yield a quantifiable
signal. The sensor shows a linear response in the range from 2.5 ppm- 21.5 ppm for perchlorate
(Figure 5.12b) and 9.8 ppm-21.5 ppm for carbonate ions (Figure 5.13b). The biosensor has high
sensitivity towards detecting and quantifying nitrate substrates at low concentrations. However,
the same electrode requires a much higher concentration of perchlorate and carbonate species to
show a quantifiable response. The estimation of the kinetic parameters enables us to make a
comparative study of the performance of the biosensor when subjected to different substrates.
Figure 5.14 shows the Lineweaver Burk plot for different configurations. The slope of the
equation is given by Km/Vmax. The y-intercept denotes 1/Vmax. While Vmax denotes the saturation
current, Km denotes the binding affinity shared by the enzyme with the substrate under
consideration. Therefore measurement of Km can provide us with a partial confirmation of the
specificity of biosensor towards the substrate under consideration.

123

(a)

(b)
Figure 5.14 Lineweaver-Burk plots for nitrate biosensor (a) Sensor fabricated at 1.1 V, 300 sec.
in presence of nitrate species. (b) Sensor fabricated at 0.95 V, 600 sec. in presence of nitrate
species. (c) Sensor fabricated at 1.1 V, 300 sec. in presence of perchlorate species. (d) Sensor
fabricated 1.1V, 300 sec. in presence of carbonate species.

124

Figure 5.14 (cont’d)

(c)

(d)

125

As seen from Table 1, the Km value for the biosensor in response to addition of nitrate
ions is close to 62.8 µM (sensor fabricated at 1.1V) and around 174.7µM (sensor fabricated at
0.95 V). The value reaches 214.3 µM for perchlorate and 38.2 mM for carbonate species. Lower
value of Km for KNO3 indicates higher binding affinity towards nitrate ions. The immobilized
enzyme requires higher perchlorate and carbonate concentration to catalyze the reduction
process. Based on Km values, the nitrate reductase biosensor (at 1.1V) developed in this work has
over three folds affinity for nitrate as compared to perchlorate substrates. The affinity for
carbonate species is almost 600 times lower than nitrate ions;
Table 5.1. Estimation of kinetic parameter based on Lineweaver-Burk approximation
Sensor fabrication
conditions
0.95 V, 600 sec
1.1 V, 300 sec
1.1 V, 300 sec
1.1 V, 300 sec

Substrate

Km

Vmax( µA/cm2)

nitrate
nitrate
perchlorate
Carbonate

174.71 µM
62.74 µM
214 µM
38.2 mM

11.136
29.15
22.52
156.25

5.3.6 Operational stability of the biosensor
The operational stability of the biosensor can be determined by successive addition of substrate
and monitoring the rise in current (See Inset of figures 6a, 7a, 8a and 9a). In each case equal
concentration of substrate was added and the response of the biosensor was measured by
estimating the rise in current for each addition. Operational parameters such as signal to noise
ratio (SNR) and limit of detection (LOD) were calculated. The SNR was estimated by taking a
ratio of mean signal to standard deviation. LOD can be estimated using co-relations described
elsewhere[65]. The comparison of these operational parameters for biosensors fabricated at
different process conditions in presence of desired substrates is presented in Table 2. Similarly
some recent examples of highly sensitive nitrate reductase based biosensors that have low LOD

126

values are included in Table 2. Table 2 shows that the sensor developed in this study (process
conditions: 1.1 V, 300 seconds) has lowest LOD among the tabulated values.
Table 5.2. Operational parameters for different types of biosensors.
Sensor fabrication
conditions
PEDOT/nitrate
reductase nanowire 0.95
V, 600 sec*
PEDOT/nitrate
reductase nanowire 1.1
V, 300 sec*
PEDOT/nitrate
reductase nanowire 1.1
V, 300 sec*
PEDOT/nitrate
reductase nanowire 1.1
V, 300 sec*
CNT/Polypyrrole nitrate
reductase[66]
MV/Nafion nitrate
reductase[67]
Ultrathin film composite
membrane(UTFCM)[68]
MV/Nafion nitrate
reductase glassy
carbon[64]
Polypyrrole-Nitrate
reductase-Azure A[26]
Polypyrrole-Ag/nitrate
reductase[24]
Nitrate reductase/Glassy
carbon[62]
Screen printed nitrate
biosensor[62]

Substrate
Potassium nitrate

Limit of Detection
(LOD) ppb
363.2

Signal to Noise
Ratio(SNR)
1.43

Potassium nitrate

108

3.75

Potassium
perchlorate

1113

3.75

Sodium Carbonate

888

4.75

Sodium nitrate

14450

3

Sodium nitrate

425

3

Potassium nitrate

525

-

Sodium nitrate

<255

-

Potassium nitrate

1010

-

Sodium nitrate

425

-

350

3

468

3

Sodium nitrate

* Reported in this study
The nitrate reductase based biosensor was tested for long term stability. Sensor stored at 4 0C and
tested after 36 hours revealed little or negligible activity towards nitrate species. One possible
explanation for this poor performance could be as a result of deterioration of PEDOT at low
127

temperature.

During the electrochemical polymerization, we observed that the EDOT

microemulsion becomes unstable at low temperatures.

When refrigerated at 4 0C, the

microemulsion breaks down with the EDOT /surfactant solution showing high levels of turbidity.
We also tested the performance of the sensor stored at room temperature. After 36 hours, the
biosensor showed about 58% reduction in current intensity when subjected to the same
concentration of nitrate species. This indicates that the enzyme and/or the PEDOT conducting
nanowires need additional stabilization. Our future studies are focused in this direction.

5.4 Conclusion
In summary, we showed the development of a highly sensitive PEDOT biosensor capable of
detecting and quantifying minute concentrations of nitrate ions. A surfactant assisted templated
approach was used to grow hybrid PEDOT/nitrate reductase nanowires. As compared to other
nitrate reductase based biosensors, the use of PEDOT/nitrate reductase nanowire structure in this
work enabled improved sensitivity, low limit of detection and instantaneous response. The
performance of the PEDOT/nitrate reductase biosensor is a function of the fabrication
conditions. Parameters like oxidation potential and time influence the degree of EDOT
polymerization and may govern the extent of enzyme entrapment within the polymer matrix.
Depending on the process conditions, nanowires of different morphologies can be grown. An
optimized biosensor with LOD close to 0.108 ppm (108 ppb) and a sensitivity of 2.6 µA/µM cm2
was obtained by fixing the oxidation potential to 1.1 V and polymerization time to 300 seconds.
The sensor shows a linear behavior in the range 200 ppb-1100 ppb. The binding efficiency of
nitrate reductase incorporated in the PEDOT biosensor is higher than interfering ions such as
perchlorate (3.5 times higher) and carbonate (600 times higher) indicating potentially high
substrate specificity. Future work will focus on improving the long term stability of the sensor by
128

investigating effect of temperature on the PEDOT sensor and preventing enzyme leaching using
chemical cross-linking agents. Methods such as irreversible entrapment of enzyme obtained by
adsorbing a mixture of enzyme and monomer on the electrode surface followed by
electropolymerization can provide quantifiable information regarding the degree of enzyme
immobilization and help develop robust comparative standards for kinetic characterization and
optimization[53, 69].

129

REFERENCES

130

REFERENCES

1.

Ward, M.H., et al., Workgroup report: Drinking-water nitrate and health-recent findings
and research needs. Environmental Health Perspectives, 2005. 113(11): p. 1607-1614.

2.

Conley, D.J., et al., ECOLOGY Controlling Eutrophication: Nitrogen and Phosphorus.
Science, 2009. 323(5917): p. 1014-1015.

3.

Anderson, D., P. Glibert, and J. Burkholder, Harmful algal blooms and eutrophication:
Nutrient sources, composition, and consequences. Estuaries, 2002. 25(4): p. 704-726.

4.

World Health, O., Guidelines for Drinking-water Quality: First Addendum to Third
Edition, Volume 1, Recommendations2006: World Health Organization.

5.

Binghui, Z., Z. Zhixiong, and Y. Jing, Ion chromatographic determination of trace
iodate, chlorite, chlorate, bromide, bromate and nitrite in drinking water using
suppressed conductivity detection and visible detection. Journal of Chromatography A,
2006. 1118(1): p. 106-110.

6.

Daniel, W.L., et al., Colorimetric Nitrite and Nitrate Detection with Gold Nanoparticle
Probes and Kinetic End Points. Journal of the American Chemical Society, 2009.
131(18): p. 6362-6363.

7.

Dasary, S.S.R., et al., Gold Nanoparticle Based Label-Free SERS Probe for
Ultrasensitive and Selective Detection of Trinitrotoluene. Journal of the American
Chemical Society, 2009. 131(38): p. 13806-13812.

8.

Rupert, M.G., Decadal-scale changes of nitrate in ground water of the United States,
1988-2004. Journal of Environmental Quality, 2008. 37(5): p. S240-S248.

9.

Avery, A.A., Infantile methemoglobinemia: reexamining the role of drinking water
nitrates. Environmental Health Perspectives, 1999. 107(7): p. 583.

10.

Fraser, P. and C. Chilvers, Health-Aspects of Nitrate in Drinking-Water. Science of the
Total Environment, 1981. 18(Apr): p. 103-116.

11.

Gulis, G., M. Czompolyova, and J.R. Cerhan, An ecologic study of nitrate in municipal
drinking water and cancer incidence in Trnava District, Slovakia. Environmental
Research, 2002. 88(3): p. 182-187.

12.

Moralessuarezvarela, M.M., A. Llopisgonzalez, and M.L. Tejerizoperez, Impact of
Nitrates in Drinking-Water on Cancer Mortality in Valencia, Spain. European Journal of
Epidemiology, 1995. 11(1): p. 15-21.

131

13.

Sandor, J., et al., Association between gastric cancer mortality and nitrate content of
drinking water: Ecological study on small area inequalities. European Journal of
Epidemiology, 2001. 17(5): p. 443-447.

14.

Ward, M.H., et al., Nitrate Intake and the Risk of Thyroid Cancer and Thyroid Disease.
Epidemiology, 2010. 21(3): p. 389-395.

15.

Braverman, L.E., et al., The effect of perchlorate, thiocyanate, and nitrate on thyroid
function in workers exposed to perchlorate long-term. Journal of Clinical Endocrinology
& Metabolism, 2005. 90(2): p. 700-706.

16.

Blount, B.C. and L. Valentin-Blasini, Analysis of perchlorate, thiocyanate, nitrate and
iodide in human amniotic fluid using ion chromatography and electrospray tandem mass
spectrometry. Analytica chimica acta, 2006. 567(1): p. 87-93.

17.

Niedzielski, P., I. Kurzyca, and J. Siepak, A new tool for inorganic nitrogen speciation
study: Simultaneous determination of ammonium ion, nitrite and nitrate by ion
chromatography with post-column ammonium derivatization by Nessler reagent and
diode-array detection in rain water samples. Analytica chimica acta, 2006. 577(2): p.
220-224.

18.

Senapati, D., et al., A Label-Free Gold-Nanoparticle-Based SERS Assay for Direct
Cyanide Detection at the Parts-per-Trillion Level. Chemistry-A European Journal, 2011.
17(30): p. 8445-8451.

19.

Lu, J., et al., Nanometal-decorated exfoliated graphite nanoplatelet based glucose
biosensors with high sensitivity and fast response. ACS nano, 2008. 2(9): p. 1825-1832.

20.

Lu, J., et al., Simple fabrication of a highly sensitive glucose biosensor using enzymes
immobilized in exfoliated graphite nanoplatelets nafion membrane. Chemistry of
Materials, 2007. 19(25): p. 6240-6246.

21.

Kohli, N., et al., Nanostructured biosensor for measuring neuropathy target esterase
activity. Analytical Chemistry, 2007. 79(14): p. 5196-5203.

22.

Campbell, W.H., Nitrate reductase structure, function and regulation: bridging the gap
between biochemistry and physiology. Annual review of plant biology, 1999. 50(1): p.
277-303.

23.

Kiang, C.-H., S.S. Kuan, and G.G. Guilbault, Enzymic determination of nitrate:
electrochemical detection after reduction with nitrate reductase and nitrite reductase.
Analytical Chemistry, 1978. 50(9): p. 1319-1322.

24.

Atmeh, M. and B.E. Alcock-Earley, A conducting polymer/Ag nanoparticle composite as
a nitrate sensor. Journal of Applied Electrochemistry, 2011. 41(11): p. 1341-1347.

132

25.

Cosnier, S., et al., Electrochemical nitrate biosensor based on poly (pyrrole–viologen)
film–nitrate reductase–clay composite. Bioelectrochemistry, 2008. 74(1): p. 47-51.

26.

Sohail, M. and S.B. Adeloju, Fabrication of Redox-Mediator Supported Potentiometric
Nitrate Biosensor with Nitrate Reductase. Electroanalysis, 2009. 21(12): p. 1411-1418.

27.

Willner, I., A. Riklin, and N. Lapidot, Electron-transfer communication between a redox
polymer matrix and an immobilized enzyme: activity of nitrate reductase in a viologenacrylamide copolymer. Journal of the American Chemical Society, 1990. 112(17): p.
6438-6439.

28.

Albanese, D., M. Di Matteo, and C. Alessio, Screen printed biosensors for detection of
nitrates in drinking water. Computer Aided Chemical Engineering, 2010. 28: p. 283-288.

29.

Wanekaya, A.K., et al., Nanowire-based electrochemical biosensors. Electroanalysis,
2006. 18(6): p. 533-550.

30.

Duvail, J.L., et al., Tuning electrical properties of conjugated polymer nanowires with the
diameter. Applied Physics Letters, 2007. 90(10).

31.

Long, Y.Z., et al., Recent advances in synthesis, physical properties and applications of
conducting polymer nanotubes and nanofibers. Progress in Polymer Science, 2011.
36(10): p. 1415-1442.

32.

Huang, J.Y., K. Wang, and Z.X. Wei, Conducting polymer nanowire arrays with
enhanced electrochemical performance. Journal of Materials Chemistry, 2010. 20(6): p.
1117-1121.

33.

Dan, Y.P., et al., Dielectrophoretically assembled polymer nanowires for gas sensing.
Sensors and Actuators B-Chemical, 2007. 125(1): p. 55-59.

34.

Corradi, R. and S.P. Armes, Chemical synthesis of poly(3,4-ethylenedioxythiophene).
Synthetic Metals, 1997. 84(1-3): p. 453-454.

35.

Jonas, F. and L. Schrader, Conductive Modifications of Polymers with Polypyrroles and
Polythiophenes. Synthetic Metals, 1991. 41(3): p. 831-836.

36.

Kros, A., N.A.J.M. Sommerdijk, and R.J.M. Nolte, Poly(pyrrole) versus poly (3,4ethylenedioxythiophene): implications for biosensor applications. Sensors and Actuators
B-Chemical, 2005. 106(1): p. 289-295.

37.

Kros, A., R.J.M. Nolte, and N.A.J.M. Sommerdijk, Poly (3,4-ethylenedioxythiophene)based copolymers for biosensor applications. Journal of Polymer Science Part a-Polymer
Chemistry, 2002. 40(6): p. 738-747.

133

38.

Dobbelin, M., et al., Innovative materials and applications based on poly(3,4ethylenedioxythiophene) and ionic liquids. Journal of Materials Chemistry, 2010. 20(36):
p. 7613-7622.

39.

Perepichka, I.F. and D.F. Perepichka, Handbook of thiophene-based materials:
applications in organic electronics and photonics2009: Wiley Online Library.

40.

Il Cho, S. and S.B. Lee, Fast electrochemistry of conductive polymer nanotubes:
Synthesis, mechanism, and application. Accounts of Chemical Research, 2008. 41(6): p.
699-707.

41.

Poverenov, E., et al., Major Effect of Electropolymerization Solvent on Morphology and
Electrochromic Properties of PEDOT Films. Chemistry of Materials, 2010. 22(13): p.
4019-4025.

42.

Liu, R., S. Il Cho, and S.B. Lee, Poly( 3,4-ethylenedioxythiophene) nanotubes as
electrode materials for a high-powered supercapacitor. Nanotechnology, 2008. 19(21).

43.

Ryu, K.S., et al., Poly(ethylenedioxythiophene) (PEDOT) as polymer electrode in redox
supercapacitor. Electrochimica Acta, 2004. 50(2-3): p. 843-847.

44.

Xiao, R., et al., Controlled electrochemical synthesis of conductive polymer nanotube
structures. Journal of the American Chemical Society, 2007. 129(14): p. 4483-4489.

45.

Cho, S.I., et al., Electrochemical synthesis and fast electrochromics of poly(3,4ethylenedioxythiophene) nanotubes in flexible substrate. Chemistry of Materials, 2005.
17(18): p. 4564-4566.

46.

Cho, S.I., et al., Nanotube‐Based Ultrafast Electrochromic Display. Advanced materials,
2005. 17(2): p. 171-175.

47.

Vidal, J.C., E. Garcia-Ruiz, and J.R. Castillo, Recent advances in electropolymerized
conducting polymers in amperometric biosensors. Microchimica Acta, 2003. 143(2-3): p.
93-111.

48.

Sakmeche, N., et al., Improvement of the electrosynthesis and physicochemical properties
of poly(3,4-ethylenedioxythiophene) using a sodium dodecyl sulfate micellar aqueous
medium. Langmuir, 1999. 15(7): p. 2566-2574.

49.

Sakmeche, N., et al., Application of sodium dodecylsulfate (SDS) micellar solution as an
organized medium for electropolymerization of thiophene derivatives in water. Synthetic
Metals, 1997. 84(1-3): p. 191-192.

50.

Bhadra, S., et al., Progress in preparation, processing and applications of polyaniline.
Progress in Polymer Science, 2009. 34(8): p. 783-810.

134

51.

Zhou, M. and J. Heinze, Electropolymerization of pyrrole and electrochemical study of
polypyrrole. 2. Influence of acidity on the formation of polypyrrole and the multipathway
mechanism. Journal of Physical Chemistry B, 1999. 103(40): p. 8443-8450.

52.

Zhang, S.S., et al., Electrochemical polymerization of 3,4-ethylenedioxythiophene in
aqueous solution containing N-dodecyl-beta-D-maltoside. European Polymer Journal,
2006. 42(1): p. 149-160.

53.

Cosnier, S., Biomolecule immobilization on electrode surfaces by entrapment or
attachment to electrochemically polymerized films. A review. Biosensors &
Bioelectronics, 1999. 14(5): p. 443-456.

54.

Wen, Y.P., et al., Electrochemical polymerization of 3,4-ethylenedioxythiophene in
aqueous micellar solution containing biocompatible amino acid-based surfactant.
Journal of Electroanalytical Chemistry, 2009. 634(1): p. 49-58.

55.

Torres-Gomez, G. and P. Gomez-Romero, Conducting organic polymers with
electroactive dopants. Synthesis and electrochemical properties of hexacyanoferratedoped polypyrrole. Synthetic Metals, 1998. 98(2): p. 95-102.

56.

Kim, Y., et al., Photoelectrochemical oxidative polymerization of aniline and its
application to transparent TiO2 solar cells. Journal of Photochemistry and Photobiology
a-Chemistry, 2009. 204(2-3): p. 110-114.

57.

Zeng, H.J., et al., Choline oxidase immobilized into conductive poly(3,4ethylenedioxythiophene) film for choline detection. Applied Surface Science, 2008.
254(20): p. 6337-6340.

58.

Lapkowski, M. and A. Pron, Electrochemical oxidation of poly(3,4ethylenedioxythiophene) - "in situ" conductivity and spectroscopic investigations.
Synthetic Metals, 2000. 110(1): p. 79-83.

59.

Du, X. and Z. Wang, Effects of polymerization potential on the properties of
electrosynthesized PEDOT films. Electrochimica Acta, 2003. 48(12): p. 1713-1717.

60.

Fic, K., E. Frackowiak, and F. Beguin, Unusual energy enhancement in carbon-based
electrochemical capacitors. Journal of Materials Chemistry, 2012. 22(46): p. 2421324223.

61.

Wen, Y.P., et al., Poly(3,4-ethylenedioxythiophene methanol)/ascorbate oxidase/nafionsingle-walled carbon nanotubes biosensor for voltammetric detection of Vitamin C.
Chinese Journal of Polymer Science, 2012. 30(3): p. 460-469.

62.

Quan, D., et al., Electrochemical determination of nitrate with nitrate reductaseimmobilized electrodes under ambient air. Analytical Chemistry, 2005. 77(14): p. 44674473.
135

63.

Bardiya, N. and J.H. Bae, Dissimilatory perchlorate reduction: A review. Microbiological
Research, 2011. 166(4): p. 237-254.

64.

Glazier, S.A., E.R. Campbell, and W.H. Campbell, Construction and characterization of
nitrate reductase-based amperometric electrode and nitrate assay of fertilizers and
drinking water. Analytical Chemistry, 1998. 70(8): p. 1511-1515.

65.

Borgmann, S., et al., Amperometric Biosensors, in Advances in Electrochemical Science
and Engineering, D.M.K. R. C. Alkire, J. Lipkowski, Editor 2011, Wiley-VCH Verlag
GmbH & Co. KGaA,Weinheim, Germany. p. 1-83.

66.

Can, F., et al., Amperometric nitrate biosensor based on Carbon
nanotube/Polypyrrole/Nitrate reductase biofilm electrode. Materials Science and
Engineering: C, 2012. 32(1): p. 18-23.

67.

Wang, X., et al., Development of a conductometric nitrate biosensor based on Methyl
viologen/Nafion< sup>®</sup> composite film. Electrochemistry communications,
2006. 8(2): p. 201-205.

68.

Moretto, L.M., et al., Nitrate biosensor based on the ultrathin-film composite membrane
concept. Analytical Chemistry, 1998. 70(10): p. 2163-2166.

69.

Cosnier, S. and C. Innocent, A novel biosensor elaboration by electropolymerization of
an adsorbed amphiphilic pyrrole-tyrosinase enzyme layer. Journal of Electroanalytical
Chemistry, 1992. 328(1–2): p. 361-366.

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Chapter 6: Conclusions and future work
6.1 Conclusions
The supramolecular assembly of different components opens up exciting opportunities for the
development of novel materials. In the preceding chapters, detailed descriptions regarding design
of an improved biocatalyst (for cellulosic ethanol production) and development of
nanostructured synergistic interfaces (conductive gas barrier films and bioelectronic enzymatic
interface for nitration detection and quantification) using supramolecular techniques have been
presented. The versatility of the applications presented here is an indication in itself of the huge
potential this field can hold for future research. The subtle manipulations brought about by
supramolecular interactions are reminiscent of the pathways evolved by nature over years of
perfection. Hence most of these forces operate at ambient process conditions (mild temperature,
pressure, pH) inflicting minimum stress on the environment and the same is observed in the
development of all the three applications discussed here. However, supramolecular forces are
also prone to drastic changes in the microenvironment when subjected to small process changes.
From the preceding discussion, this fact is attested by variation in colloidal stability of the
biocatalyst as a result of the swelling action of the spacer polyelectrolyte (Chapter 3), change in
the film morphology and hence permeation and conductance due to change in deposition pH
(Chapter 4) and loss of accurate nitrate quantification capability due to changes in the
electropolymerization of polymer/enzyme nanowires (Chapter 5). In view of the several facets
observed during the design and development of the novel materials discussed here, a
recapitulation of some important findings for each application is in order. The same are
presented in this final chapter.

137

The first project involved development of a novel nanobiocatalyst that is capable of overcoming
the geometric limitation (due to 2D geometry) and steric hindrance as a result of enzyme
immobilization to deliver improved catalytic activity. The biocatalyst uses a supramolecular
bottom-up approach that incorporates a plethora of charged components such as polyelectrolytes,
nanoparticles and cellulase enzymes. The incorporation of each component imparts additional
functionality to the biocatalyst architecture and the same has been discussed in chapter 3. Salient
features of these nanobiocatalyst are improved dispersion of high surface area graphene platelets
stabilized by supramolecular assembly of strong polylelectrolytes, inclusion of iron-oxide
nanoparticles to facilitate recycle and recuse of the immobilized enzyme and pH tunable swelling
of polyacrylic acid brushes to manipulate colloidal stability and hence enzymatic activity. The
immobilized enzymes developed in this study show an activity as high as 80% compared to free
enzymes with retention in specific enzyme activity up to 55% after four cycles of reuse.
In Chapter 4, the supramolecular assembly of cationic polyelectrolyte functionalized graphene
platelets along with anionic polyelectrolyte was discussed and the material properties of these
films wrt oxygen barrier and conductance were reported. Three separate types of assemblies
were identified by making subtle changes in the deposition conditions to produce predominantly
(a) hydrogen bonded films (b) electrostatic films (c) composite films. In all the three cases, the
same cationic and anionic polyelectrolytes were used. However, a variation in the pH condition
during the film deposition process resulted in change in the charge density of these
polyelectrolytes. As a result, films of different types of interactions were fabricated. The
composite films developed in this work have oxygen barrier properties comparable to
commercial grade silicon oxide coatings and resistance that decreased by high orders of
magnitude. With more modifications, a further decrease in electrical sheet resistance is possible

138

which would improve the chances of these films to be used in the semiconductor/electronics
industry.
In Chapter 5, an electropolymerized PEDOT/nitrate reductase nanobiosensor was developed
from a surfactant stabilized supramolecular matrix. The sensor enjoys low LOD (close to 0.1
ppm nitrate) and shares a high binding affinity towards nitrate species as compared to other
interfering anions. Manipulating the electropolymerization time and potential causes a change in
the morphology of the PEDOT/nitrate reductase nanowires. It is expected that the level of
enzyme entrapment with the polymer matrix can be tweaked by proper optimization of these two
parameters. Another important feature of this sensor is the retention of 1D nanowire dimension.
Because of the templated approach used to grow the nanowire, the PEDOT/enzyme nanowires
return their upright nanowire dimensions contributing to increased signal efficiency and hence
better detection and quantification.

6.2 Future outlook
The design of the supramolecular nanostructures discussed in the preceding chapters holds
immense promise to expand the existing work to several new applications. For example, in
Chapter 3, iron-oxide doped graphene platelets were used as nanocarriers for immobilization of
cellulase in the saccharification reaction. However, because of the sheer robustness and
simplicity of assembly, it is possible to substitute cellulase with other enzymes. This means that
the same graphene based support can be used in a number of biocatalytic reactions. Besides this,
a very novel extension of using this nanostructured support can be in the field of targeted drug
delivery applications. Several variations of magnetic carbonaceous composites for biomedical
applications such as cancer treatment and magnetic resonance imaging (MRI) are already
available in literature [1-4]. While the ferrous component of the composite helps in guiding or
139

tracking the flow of the drug/tracer in the body, the carbonaceous component provides an
excellent contrast for imaging[5]. Other than the biobased applications, a potential extension of
graphene-iron oxide carriers (without the enzyme component) for selective oxidation of
secondary alcohols is possible since carbon -iron oxide carriers have been shown to be active in
catalytic oxidation reactions with performance comparable to noble-metals[6]. Likewise, the
layer-by-layer assembly of polyelectrolyte functionalized graphene platelets also holds immense
promise for incorporation of additional components to suit the requirements. Chapter 4
specifically dealt with fabrication of oxygen barrier films. However based on recent research by
making minor modifications in these films robust corrosion resistant films can be easily built[7,
8]. Polyelectrolytes are effective in covering several types of substrates shielding the existing
defects in their microstructure from direct attack of air and water[8-10]. This property has been
shown to provide a comfortable degree of protection to the substrate in presence of a corrosive
environment. The LbL assembly also allows for the incorporatation of several non-homogenous
components such as cells, nanoparticles charged species within the multilayer structure. This can
be exploited to ensure value addition to the supramolecular films. The inclusion of corrosion
inhibitors[8] and hydrophobic silica nanoparticles[11, 12] are some excellent options to limit the
effect of corrosion and exposure to water respectively. Likewise, the third application discussed
in this dissertation namely development of nanostructured biosensor for detection of contaminant
ions can be made more user-friendly by use of oxygen scavengers. The nitrate quantification
studies shown here used deoxygenated test solution obtained after nitrogen/argon purging for
estimation. Further improvements such as addition of oxygen scavenging agents like sodium
sulfite or cysteine can prevent the necessity of purging argon/nitrogen through the solution.
Similarly, enzymes such as oxyrase or catalase consume oxygen[13]. It has been shown that

140

supramolecular biocompatible polyelectrolyte films can be used for the controlled release of
enzymes, drugs and nanoparticles entrapped within the multilayers by adjusting process
condition such as pH and temperature[14-17]. Supramolecular assembly of oxygen scavengers
post electropolymerization followed by release in the test solution is another interesting option
that can be explored in the future. This should help in the elimination of the inert gas purging
step and improve the portability of the sensor.

141

REFERENCES

142

REFERENCES

1.

Ma, Y., et al., Plasma synthesis of carbon magnetic nanoparticles and immobilization of
doxorubicin for targeted drug delivery. Journal of Biomaterials Science, Polymer Edition,
2004. 15(8): p. 1033-1049.

2.

Bystrzejewski, M., A. Huczko, and H. Lange, Arc plasma route to carbon-encapsulated
magnetic nanoparticles for biomedical applications. Sensors and Actuators B: Chemical,
2005. 109(1): p. 81-85.

3.

Rudge, S., et al., Preparation, characterization, and performance of magnetic iron–
carbon composite microparticles for chemotherapy. Biomaterials, 2000. 21(14): p. 14111420.

4.

Leconte, Y., et al., Continuous production of water dispersible carbon–iron
nanocomposites by laser pyrolysis: Application as MRI contrasts. Journal of colloid and
interface science, 2007. 313(2): p. 511-518.

5.

Harrison, B.S. and A. Atala, Carbon nanotube applications for tissue engineering.
Biomaterials, 2007. 28(2): p. 344-353.

6.

Gao, Y., et al., Layered‐Carbon‐Stabilized Iron Oxide Nanostructures as Oxidation
Catalysts. Angewandte Chemie International Edition, 2011. 50(43): p. 10236-10240.

7.

Shchukin, D.G., et al., Layer‐by‐Layer Assembled Nanocontainers for Self‐Healing
Corrosion Protection. Advanced materials, 2006. 18(13): p. 1672-1678.

8.

Andreeva, D.V., E.V. Skorb, and D.G. Shchukin, Layer-by-layer polyelectrolyte/inhibitor
nanostructures for metal corrosion protection. Acs Applied Materials & Interfaces, 2010.
2(7): p. 1954-1962.

9.

Farhat, T.R. and J.B. Schlenoff, Corrosion control using polyelectrolyte multilayers.
Electrochemical and solid-state letters, 2002. 5(4): p. B13-B15.

10.

Andreeva, D.V., et al., Buffering polyelectrolyte multilayers for active corrosion
protection. Journal of Materials Chemistry, 2008. 18(15): p. 1738-1740.

11.

Zhai, L., et al., Stable superhydrophobic coatings from polyelectrolyte multilayers. Nano
Letters, 2004. 4(7): p. 1349-1353.

12.

Bravo, J., et al., Transparent superhydrophobic films based on silica nanoparticles.
Langmuir, 2007. 23(13): p. 7293-7298.
143

13.

Patel, J., et al., Comparison of oxygen scavengers for their ability to enhance
resuscitation of heat-injured Listeria monocytogenes. Journal of Food Protection®, 1995.
58(3): p. 244-250.

14.

Chung, A. and M. Rubner, Methods of loading and releasing low molecular weight
cationic molecules in weak polyelectrolyte multilayer films. Langmuir, 2002. 18(4): p.
1176-1183.

15.

Zhu, Y., et al., Stimuli‐Responsive Controlled Drug Release from a Hollow Mesoporous
Silica Sphere/Polyelectrolyte Multilayer Core–Shell Structure. Angewandte Chemie, 2005.
117(32): p. 5213-5217.

16.

Qiu, X., et al., Studies on the drug release properties of polysaccharide multilayers
encapsulated ibuprofen microparticles. Langmuir, 2001. 17(17): p. 5375-5380.

17.

Shi, X. and F. Caruso, Release behavior of thin-walled microcapsules composed of
polyelectrolyte multilayers. Langmuir, 2001. 17(6): p. 2036-2042.

144