Synthesis Strategies for Highly Functional
Enzyme-based Conjugates
Gabriela C. Perhinschi
Problem Report submitted to the
Benjamin M. Statler College of Engineering & Mineral Resources
at West Virginia University
in partial fulfillment of the requirements for the degree of
Master of Science in
Mechanical Engineering
Cerasela Zoica Dinu, Ph.D., Chair
Edward Sabolsky, Ph.D.,
Nianqiang Wu, Ph.D.
Department of Mechanical and Aerospace Engineering
Morgantown, West Virginia
2016
Keywords: enzyme, activity, stability, functionality, nanosupport
ABSTRACT
“Greener-based” processes and products can reduce the environmental burden of
chemical synthesis and energy production. Such green-based technologies could be based on
enzymatic biocatalysis. Enzymes allow for reaction processes to operate under mild conditions
with greatly reduced waste generation while increasing yield in selected production scenarios.
However, to accomplish the implementation of such enzyme-based technologies, immobilization
and recovery of the working enzymes are necessary.
This Problem Report highlights the features of enzyme as biological catalysts and
provides examples of specific enzyme-based technologies used both for industrial and clinical
biocatalysis. Further, the report highlights the means for implementation of enzyme-based
immobilization techniques, as well as the advantages of such techniques in terms of enzymebased conjugates recovery, enzyme activity and stability, as well as product ease of separation.
Special consideration is given to carbon-based nanomaterials as supports for enzyme
immobilization because of their unique, advantageous properties such as high surface area,
exceptional conductivities and ease of functionalization. The last part of this report highlights my
own work on enzyme-based immobilization onto such materials, specifically discussing the
methods being applied, as well as advantages and applications of enzymes and enzymes
immobilization techniques on two types of such nanotubes respectively. Fundamental studies of
enzyme-based conjugates can lead the means for implementation of enzymatic catalysis for
designing efficient and effective processes and materials aimed at meeting the goals of enzymebased technologies for high product recovery, all while producing revenues from otherwise
inaccessible sources.
Table of Contents
Chapter 1 Introduction
1
Chapter 2 Enzymes for Industrial and Commercial Applications
3
2.1. Classification of Enzymes
3
2.2. Properties of Enzymes
5
2.3. Increasing Enzyme Activity and Stability
6
2.4. General Applications of Enzymes
7
Chapter 3 Enzyme Immobilization
10
3.1. Support Binding
11
3.1.1. Physical Adsorption
11
3.1.2. Covalent Bonding
12
3.1.3. Ionic Bonding
12
3.2. Entrapment
13
3.2.1. Occlusion within a Cross Linked Network
13
3.2.2. Microencapsulation
13
3.3. Cross-linking
13
Chapter 4 Carbon Nano-tubes for Enzyme Immobilization
15
4.1. CNT Classification
15
4.2. General Properties of CNT
15
4.3. CNT Production Methods
16
4.4. CNT Functionalization
17
4.5. Enzyme Immobilization on CNT
18
Chapter 5 Investigation of Effects of CNT Immobilization on Enzyme
Functionality
20
5.1. Material and Methods
20
5.1.1. Acid Oxidation of CNTs
20
5.1.2 Energy Dispersive X-ray Analysis (EDX) of CNTs
21
5.1.3. Scanning Electron Microscopy (SEM) of CNTs
21
5.1.4. Raman Spectroscopy of CNTs
21
iii
5.1.5. CNTs Solubility Measurement
21
5.1.6. Functionalization of CNT with Enzyme
22
5.1.7. CNTs Length and Morphology Measurement
23
5.1.8. Enzyme Loading
23
5.1.9. Enzyme Activity Assay
24
5.2. Results and Discussion
25
Chapter 6 Conclusions
35
References
36
Appendices
iv
List of Figures
Figure 1. Crystal structure of dithinonite reduced soybean ascorbate peroxidase mutant
W14A.
Figure 2. Chloroperoxidase f/propionate complex 1A8S.
Figure 3. (Reproduced after Campbell et al., Figure 1, ACS Appl. Mater. Interfaces
2014, 6, 5393−5403; [76]). Analysis of functional groups of the carbon-based materials. FTIR
and EDX spectra analysis of a) pristine and acid functionalized SWCNTs, b) pristine and acid
functionalized MWCNTs.
Figure 4. Reproduced after Dong et al., Applied Surface Science 264 (2013) 261–
268, [11]. (a) EDX elemental analysis of pristine SWCNTs; and b) MWCNTs. The inlets show
the changes in the contents of O and Fe with the time-controlled acid treatment.
Figure 5. Reproduced after Dong et al., Applied Surface Science 264 (2013) 261–
268, [11]. Scanning Electron Microscopy (SEM) images of a) pristine SWCNTs, b) pristine
MWCNTs, c) 6 h acids treated SWCNTs, and d) 6 h acids treated MWCNTs; the scale bar is 1
µm.
Figure 6. Atomic force microscopy (AFM) of pristine SWCNTs and acid treated for 6 h.
Figure 7. General mechanism of protein immobilization onto nanosupports. (a) Carbon
nanotube acid treatment. (b) Physical adsorption of enzymes onto nanosupports. (c) Covalent
binding of enzymes onto nanosupports.
Figure 8. a) Functionalized SWCNTs with SBP; b) Height profile of the enzymeSWCNT conjugates. The line was taken across the highlighted nanotube (in image a) and shows
a bead-like morphology of the immobilized enzymes.
v
Chapter 1
Introduction
Enzymes are organic chemical compounds (biological molecules) that act as catalysts for
an enormous number of biochemical reactions [1]. Briefly, an enzyme facilitates and accelerates
the transformation of an initial chemical compound or structure referred to as the substrate into a
final chemical compound referred to as the product without being modified itself in the process
or altering the overall reaction balance [2].
Critical processes in living cells and full organisms rely on activating enzymes to sustain
adequate rates of reaction to allow for the complexity of the biological system and its
functionality. As such, the individual chemical reactions influenced by the enzymes become
building blocks of highly complex processes or systems that allow for natural life thus making
the enzymes responsible for diverse biological functions from digestion, to antibody generation,
and from muscle contraction, to metabolic release of energy, bio-signal transfer, cell activity
regulation, and immune response, just to name a few [3].
Any particular enzyme can react with only one or a limited number of different substrates
to accelerate their conversion into products. The relationship between enzymes and substrates
has been often associated to either a “key-lock” or a “fit-model” interaction and it is being
studied and associated with enzyme-substrate model bindings. The two theories assume the
different means that the substrate plays in determining the final shape of the enzyme and
therefore the substrate binding affinity. The difference between the two theories is based on the
flexibility of the enzyme and whether the substrate distorts or not its chemical structure upon
binding. However, for both theories, the substrate needs to interact with the active site of the
enzyme through opposite charges to cause changes in the electron distribution of its chemical
bonds will ultimately lead to product formation. Subsequent release of the product from the
active site will allow for the active site regeneration and another enzymatic cycle to start.
The high specificity in substrate binding [4] distinguishes enzymes from inorganic
catalysts and makes them the preferable alternative for a wide range of industrial and commercial
applications since they always “stay on target”, they do not affect other substrates and do not
generate side-products or effects. Thus, enzymes are currently being used on a large scale in
processing food, leather, textile fibers, and paper pulp; in enhancing detergents and treating
1
waste; and for producing bio-fuel and oil-derived products [5]. In medicine, enzymes are being
used for the production and delivery of drugs, for diagnostic, and for treatment of enzyme-related
disorder and other diseases [6]. More recently, extensive research efforts have been invested to
expand the applicability of enzymes to areas such as biosensing [7], decontamination [8], or
tissue engineering [9].
However, in both the more established applications as well as in the newly investigated
ones, a major impediment is represented by the enzyme general lack of stability and high
sensitivity when exposed to changing environmental factors such as pH or temperature [10]. For
instance, once placed in a synthetic environment, the proper functionality of enzymes is only
maintained for a limited period of time with their catalytic characteristics degrading rapidly once
the narrow ranges of temperature and pH (normally associated and determined by the chemical
structure of the enzyme and its overall charge) are exceeded. Further, solubility in water and
other solvents may prevent recovery after use in industrial processes. To eliminate or mitigate
these problems, enzymes are immobilized on solid supports [11] either through weak physical or
chemical bonds, with such methods being expected to preserve enzyme’s catalytic properties
while enhancing its stability and robustness. Immobilization strategies thus opened a wide field
of study to establish the proper physical and chemical nature of the solid support, the potential
interactions between the support and the enzyme, and the potential effects on enzyme
functionality within the enzyme-support conjugate that will allow for preserved maximum
functionality of the enzyme.
In this brief research summary, a description of enzyme and their characteristics, as well
as a classification of the most widely used enzymes for industrial and commercial purposes are
provided. The general approaches for enzyme immobilization are also outlined. A special
consideration is being given to carbon nanotubes (CNT) and their usage as solid supports for
enzyme immobilization. Such strategies are outlined in the context of the work studied and
personally performed in the lab of Prof. Dinu (involved in my coordination as a graduate
student). The effects of different CNTs in conjunction with representative enzymes on conjugate
properties and functionality are presented followed by a summary of recommended future
strategies and approaches to improve and extend enzyme-based applicability.
2
Chapter 2
Enzymes for Industrial and Commercial Applications
2.1. Classification of Enzymes
The International Union of Biochemistry and Molecular Biology has formulated criteria
for the classification and the nomenclature of enzymes [12, 13]. Depending on the type of
reactions that they could catalyze, enzymes may belong to one of the six large classes known as
oxidoreductases, transferases, hydrolases, lyases, isomerases, and ligases. Briefly:
•
Oxidoreductases catalyze oxidation/reduction reactions in which hydrogen or oxygen
atoms or electrons are transferred among molecules (e.g. dehydrogenases, oxidases);
•
Transferases accelerate the transfer of functional groups (such as a methyl or a phosphate
group) among molecules (e.g. transaminase, kinases);
•
Hydrolases catalyze the hydrolysis or the reaction of substrate with water, resulting in
two products (e.g. estrases, lipases, peptidases);
•
Lyases accelerate the addition or removal of various groups by means other than
hydrolysis and oxidation (e.g. decarboxylases, aldolases);
•
Isomerases catalyze isomerization changes within a single molecule, which consists of an
intramolecular re-arrangement of the same atoms and groups (e.g. isomerase, fumarase,
mutase);
•
Ligases join two molecules with covalent bonds typically coupled with adenosinetriphosphate hydrolysis (e.g. glutamine synthetase, cobalt chelatase).
By international convention, an enzyme is labeled by a four-digit number following the
EC (Enzyme Commission) acronym, which specifies the class and sub-classes that the enzyme
belongs to. Two important enzymes will be referred to later in this study. The first one is
soybean peroxidase (SBP), with the systematic name phenolic donor: hydrogen-peroxide
oxidoreductase and the four-digit label EC 1.11.1.7 [14], presented in Figure 1 [15]. The second
is
chloroperoxidase
(CPO),
with
the
systematic
name
chloride:
hydrogen-peroxide
oxidoreductase and the four-digit label EC 1.11.1.10 [14] presented in Figure 2 [16].
3
Figure 1. Crystal structure of dithinonite reduced soybean ascorbate peroxidase mutant
W14A [15]. Alpha helices predominate in the enzyme structure. Structure reproduced from
Badyal, S.K., Metcalfe, C.L., Basran, J., Efimov, I., Moody, P.C.E., Raven, E.L. “Iron Oxidation
State Modulates Active Site Structure in a Heme Peroxidase”, Biochemistry, 47: 4403.
Figure 2. Chloroperoxidase f/propionate complex 1A8S [16]. Both alpha helixes and
beta sheets can be identified. Structure reproduced from Hofmann, B., Tolzer, S., Pelletier, I.,
Altenbuchner, J., van Pee, K.H., Hecht, H.J., “Structural investigation of the cofactor-free
chloroperoxidases”, J.Mol.Biol. 279: 889-900.
4
The two enzymes have been selected based on their known chemical composition and
properties as well as implementability in a variety of applications. Namely, soybean peroxidase
(SBP) is an anionic monomeric glycoprotein with a molecular weight of ~40 kDa that has high
thermostability and oxidation potential and was applied in diagnostics [17, 18] and waste-water
treatment [19, 20]. Complementary, chloroperoxidase (CPO) is a monomeric enzyme with a
molecular weight of ~42 kDa used for chiral organic synthesis [21] and decontamination [22],
just to name a few.
2.2. Properties of Enzymes
Enzymes are complex biological molecules, typically classified as proteins; they
accelerate high-energy reactions by replacing them with sequences of low energy reactions.
Enzyme building blocks consist of versatile amino acids joined together to form chainlike macro-molecular structure, i.e. the primary structure. When the primary structure organizes
in two dimensions, secondary (beta sheets, alpha helixes or coiled coils conformations) are being
formed. The complexity of these proteins and the structural arrangements of their amino acids
allows for understanding their catalytic properties. Specifically, structural geometry of these
molecules allows the exposure of specific sections of the molecular chain to contact with the
substrate. These sections form the so-called active sites to which the substrate may bind if
chemical compatibility exists. Therefore, for any enzyme, only a specific substrate (or a limited
number of closely similar substrates) will match and be targeted for a catalyzed chemical
reaction. The enzyme will be inert to any other substrates. There are several types of enzyme
specificity [23]:
•
Absolute substrate specificity - the enzyme will catalyze only one reaction affecting only
one substrate; it may also be stereochemical specific if it only reacts with specific
substrate isomers;
•
Dual specificity - the enzyme may act on one substrate to catalyze two different reactions
or act on two substrates for the same type of reaction;
5
•
Moderate or group specificity - the enzyme will act only on molecules that have specific
functional groups; it is specific not only to the type of bond but also to the molecular
structure;
•
Low or bond specificity - the enzyme will act on a particular type of chemical bond
regardless of the rest of the molecular structure.
Enzymes are not transformed at the end of the process and, they are not part of the
product. Typically they stay unaltered and can be active indefinitely in proper pH and
temperature conditions.
Combining their specificity and their biodegradable nature makes enzymes the most
environmentally friendly alternatives for industrial applications. However, their functionality
may degrade significantly, especially if temperature and pH exceed rather narrow optimal
ranges.
2.3. Increasing Enzyme Activity and Stability
Enzyme applications are currently limited by enzyme’s lack of stability in a wide range
of pH and high temperatures. As a consequence, protein engineering is an active area of research
and involves attempts to create enzymes with novel properties or modify the existing enzyme to
increase their activity and stability as well as life time in a variety of synthetic environments
including changing pH, temperatures or solvents [11]. A strategy to improve the enzymatic
activity at a higher pH for instance consists of using a water-soluble atom transfer
polymerization reaction [24]. Using this strategy, the enzyme chymotrypsin has been modified
and the resulting conjugate proved active at a higher pH and temperature then the untreated or
pristine enzyme. Another much studied strategy to improve the activity and stability of enzymes
is by immobilization on different substrates. Out of all methods of immobilization investigated,
three methods proved to be more efficient and are more common: absorption, entrapment, and
covalent binding to a support. Research has demonstrated that by modification of biochemical
and physical properties of engineered or immobilized enzymes, problems related to stability,
activity, and selectivity can be solved [25].
6
2.4. General Applications of Enzymes
There are a variety of enzyme-based applications in both industrial and consumer
settings. Such applications do not only aim reducing the footprint of unstable or dangerous
byproducts but even further, intend reducing the costs associated with product specificity,
selectivity and yield. Such applications are being discussed next.
An example of an enzyme widely used in the pharmaceutical industry is tyrisinase.
Tyrisinase is a natural enzyme used in producing L-3, 4-dihydroxyphenylalanine or L-DOPA, a
chemical substance produced naturally by the human body. L-DOPA outside the human body
permits could be used for treating Parkinson disease. The tyrosinase also produces melanin, a
pigment with numerous applications not only in the pharmaceutical industry but also in cosmetic
industries, and food industries [26].
Lipase is known for its capability of removing fatty residues and cleaning of the clogged
drains that made the enzyme suitable to be added to detergents. The cleaning power of lipasebased detergents increases markedly however other enzymes, such as proteases, amylases,
cellulases and lipases are also currently being added to the detergents to improve their efficiency
[27]. Using the detergents with incorporated enzymes results into a more economical wash, at a
lower temperature and thus a shorter processing time.
Incorporating and using enzymes in the paper industry started back more effectively in
1986. Cellulase, ylanase, laccase and lipase are the most important enzymes used in the pulp and
paper processes. The key role and objectives of xylanases have been in the boosting of the
bleaching processes. Beside xylanases, accases have been primarily used in pulp production. The
fiber modification has been obtained by using cellulases and lipase [28].
The role of enzymes in the remediation of polluted environment became a new field of
research in the recent years. Specific enzymatic classes are studied for environmental
decontamination of different pollutants such as polyphenols, nitriles, polycyclic aromatic
hydrocarbons (PAHs), cyanides and heavy metals respectively. Such pollutants have been shown
to be harmful for human life as well as for environment. Therefore, the need to remove them in a
healthy and economical way has increased with the development of new industries producing
more and more of such residues. The use of enzymatic processes has been shown to be preferred
7
due to the fact that they can produce extensive transformations conducting to complete
conversion of pollutants into innocuous end products [29].
For instance, textile industry is a big producer of dyestuff as a residue of the chemical
processes. The persistent problem of contamination of the water resources is dangerous and has
increased as these industries developed and grew. The pollutants such as dyestuffs are very
resistant making their elimination from the wastewater difficult by classical methods (e.g.,
physical or chemical decontamination). The use of natural enzymes would be preferred because
they biodegrade and are easy to integrate in the decontamination platform. The class of enzymes
studied for use in bioremediation is the oxidoreductase, which exchanges transfer electrons with
the pollutant. Further, in the case of oxidoreductases at the end of the reaction the enzyme
regenerates itself and can be used again in a new cycle [29].
Gluconases are widely used due to their very high efficiency. Gluconases break down the
wheat and convert the carbohydrates into sugars that speed up the reaction of the beer
fermentation [30]. Amylases also used in beer brewing, splits starch into dextrins and sugars [31]
In dairy industry, enzymes such as protease and chymosin, which are coagulants, are
added to milk to hydrolyze the caseins and thus bring the milk into solid form [32]. Other
category of enzymes makes the products lactose free without compromising the taste [33].
Enzymes have also been used in the baking industry for decades. The yeast is responsible
for fermenting the sugars, transforming them in alcohol and carbon dioxide, resulting in the rise
of dough [34]. The main component of the wheat is the starch. Amylases degrade starch and
produce small dextrines for the yeast to act on [35].
Enzymes are applied in organic synthesis targeting several areas such as modifying the
reaction mechanism of the enzyme to catalyze new reactions, changing substrate specificity,
expanding substrate specificity, and improving substrate specificity. These modifications can be
obtained by redesigning the enzyme structure, or by mutagenesis methods followed by screening.
Both methods of enzyme engineering can be successful and are very useful for improving the
utility of enzymes for applied catalysis. [36].
Lastly, the applications of enzymes for the next generation of biosensors extended
enzyme incorporation in environmental testing, biowarfare agent detection and clinical testing.
Moreover, in the last few years, enzyme-based systems have been proposed for point-of-care
testing. Strategies for measuring enzyme activity by an essay have been studied. A paper-based
8
device has been for instance proposed and was shown to be capable of measuring the time for a
reference portion of the paper to change color to green relative to an assay region [37]. Such
applications are being driven by the need to minimize the costs of health care while reducing the
time for diagnosis.
9
Chapter 3
Enzyme Immobilization
Enzymes depend on strict conditions to function properly. Once placed in a synthetic
environment for industrial/commercial applications for instance, they could exhibit chemical
instability and high sensitivity to the encountered external factors. To be able to store and use
enzymes for long durations in an efficient and economical manner, they must be processed such
that these types of issues are minimized or completely solved.
Proposed strategies aim first to extract, isolate/purify the crude enzyme and subsequently
attach it to a solid insoluble support either through physical and/or chemical bonding; the
enzyme is said to be now immobilized [38-39]. The resulting enzyme-support conjugate is
expected to present increased stability, robustness, and resilience, while maintaining or
enhancing the catalytic and specificity properties of the enzyme alone. For commercial
applications, the immobilized enzymes present additional desirable characteristics, such as:
•
Easy removal and separation from product such that feedback inhibition is reduced and
re-use facilitated;
•
Easy packing for storage and use over long periods of time;
•
Increased thermal stability allowing higher operational temperature with additional
catalytic effect.
A large variety of support materials have been considered [38], including both organic and
inorganic materials. For instance:
1. Organic enzyme support materials
1.1. Natural polymers
1.1.1. Polysaccharides (e.g. cellulose, dextrans, agar, agarose, chitin, alginate)
1.1.2. Proteins (e.g. collagen, albumin)
1.1.3. Carbon
1.2. Synthetic polymers
1.2.1. Polystyrene
1.2.2. Other polymers (e.g. polyacrylates, polymethacrylates, polyacrylamides,
10
polyamides,vinyl, allyl-polymers)
while,
2. Inorganic enzyme support materials were comprised of:
2.1. Natural minerals (e.g. bentonite, silica, metal oxides)
2.2. Processed materials (e.g. nonporous glass, controlled pore glass, metals, controlled
poremetal oxides, ceramics)
The current main approaches for enzyme immobilization may be classified in three large
categories such as support binding, entrapment, and cross-linking [40]. The description of
various alternative methods, their advantages and disadvantages are briefly outlined next.
3.1. Support Binding
The support binding approach to enzyme immobilization aims to establish an intimate
connection between the enzyme and the support itself, which could be done either through
physical or chemical bonds. Depending on the nature of the desired bonds, methods have been
developed that rely on physical adsorption, covalent bonding, or ionic bonding respectively (just
to name the few most well studied and implemented).
3.1.1. Physical Adsorption
Physical absorption is based on weak bonds to be formed between the enzyme and the
surface of water-insoluble supports, such as those produced by van der Waals forces, hydrogen
bridges, electro-static forces, or hydrophobic interactions [41, 42]. The physical-based method is
known to produce reversible or weak types of binding. As such, the enzyme may be located on
the external or internal surface of the support, if porous materials are used for instance and could
be easily removed under harsher conditions such as sonication or dispersion in different
reactions. While external surface binding is easy to perform, it has the disadvantage of
potentially exposing the enzyme to microbial attacks or physical abrasion when synthetic
applications of enzymes are being consider/envisioned. The internal immobilization, on the other
11
hand, has to face the issue of pore diffusion and enzyme deformability at such interfaces, which
could result in enzyme loss of functionality and thus subsequently reduced product yield.
In general, physical adsorption, while not preventing enzyme alteration, is simple and
inexpensive, does not require complex and expensive reagents, possesses wide applicability, and
allows for high enzyme loading. However, practical methods may take long periods of time
(static process) and/or the application of mechanical agitation (dynamic process). Further,
desorption of the enzyme may occur under temperature and pH variations. In some situations, the
active sites of the enzyme may be blocked by the support that the enzyme is being immobilized
onto, thus significantly reducing the activity of the conjugate and further, affecting its future
implementation.
3.1.2. Covalent Bonding
Covalent bonding is the most frequently used method for enzyme immobilization [43].
Relying on strong covalent bonds between enzyme and the support being used, this method
produces irreversible binding with limited or inexistent enzyme leakage from the support. The
practical production of the conjugate itself is simple, may be performed under mild conditions,
and shows wide applicability. However, the method exposes the enzyme to possible chemical
modification and only low ratios of enzyme versus matrix may typically be achieved. Further,
one of the main disadvantages of such technique is that once the catalytic performance of the
enzyme decays, the matrix cannot be re-used since the bonds being formed cannot easily be
broken through physical means.
3.1.3. Ionic Bonding
Ionic bonding relies on compatible ion-exchanging capabilities of the enzyme and
support [44]. The method is typically simple and expected to produce reversible binding;
however, it is usually difficult to find adequate compatibility between enzyme and support such
that the enzyme remains strongly bound without possibly reducing its catalytic performance.
Incompatibility between a highly charged support and the substrates or products may also occur,
which can further significantly affect enzyme properties.
12
3.2. Entrapment
In entrapment [45], the enzyme molecules are not directly attached to the support surface
but rather trapped inside it. The support may typically consist of a polymeric network or support
microcapsules and allows the substrate and products to pass through, while retaining the enzyme.
3.2.1. Occlusion within a Cross Linked Network
Occlusion within a cross-linked network uses as a support matrix a gel [46] or a fiber [47]
entrapping. Entrapment may be performed by mixing the enzyme into a monomer solution,
followed by polymerization of that particular solution that could usually be initiated by a change
in temperature or by a chemical reaction. While the method ensures good preservation of
catalytic properties after immobilization, it is limited by the possible occurrence of enzyme
leakage and by the amount of mass that can be transferred throughout the network itself. This
may also prevent the substrate from penetrating deep inside the network thus limiting the amount
of the product being generated, i.e. the product yield.
3.2.2. Microencapsulation
Microencapsulation relies on the formation of small spherical capsules out of support
material in which the enzyme is included as a liquid or a suspension [48]. The polymeric
membrane of the capsules is selectively permeable allowing the transfer of substrate and product
while keeping the catalyst inside. With this approach, loss of enzyme reactivity is minimized.
However, the method is limited by the amount of mass that can be transferred through the
membranes of the microcapsules. This may also hinder the circulation of substrate and product
respectively.
3.3. Cross-linking
This method is based on the covalent bonding of enzyme molecules in the presence of
functional reagents [49, 50]. Very often, this process is achieved in the absence of any support.
13
Due to the nature of the covalent bonds, reduced enzyme desorption or leakage is recorded. The
operational stability is very high, even under demanding conditions. However, chemical
modifications of the enzyme surface, in particular the active sites, may occur and the diffusion
rates of both the substrate and the product may be negatively affected. The cross-linking agent
must be carefully selected such that the structural and functional properties of the enzyme during
the process of immobilization are preserved. This requirement typically leads to complex
reaction conditions and scenarios and the use of toxic reagents.
14
Chapter 4
Carbon Nanotubes for Enzyme Immobilization
Nanomaterials [51] have been investigated as support of choice for enzyme
immobilization primarily due to their promising capability to ensure high aspect ratios - relative
to their mass or volume - for contact area, enzyme mass adsorption, and effective enzyme
loading [52]. Among the various types of nanomaterials, such as nanoparticles, nanofibers, and
nanotubes, carbon nanotubes (CNT) have received substantial attention [53] because they are not
difficult or expensive to produce, they possess desirable mechanical, electrical, and thermal
properties which could subsequently lead to high biocompatibility with respect to wide classes of
enzymes [53]. CNTs are produced by various approaches out of graphitic sheets that are rolled
up into small cylinders with lengths in the range of micrometers and diameters in the range of
nanometers, with typical length-to-diameter ratio larger than 1 million.
4.1. CNT Classification
CNT are typically classified as multi-walled (MWCNT) and single-walled (SWCNT).
These graphite tubules have been discovered in the early 90’s [54, 55], and have ever since been
investigated as alternative support materials for enzyme immobilization. While both types share
the cylindrical shape, MWCNT contain at least two layers with outer diameters between 3 and
100 nm, while the SWCNT present one single layer with smaller diameters, usually between 1
and 2 nm. Because the carbon atoms form hexagonal rings on plane sheets, depending on the
direction about which the sheets are rolled up to form the cylinders, CNT can also be classified
as armchair, zigzag, or chiral CNT [56] which are known to determine their physical and
chemical functionality.
4.2. General Properties of the CNTs
The covalent bonds between carbon atoms and the catenation property are responsible for
the specific mechanical properties of the CNTs [57]. In particular, CNTs may reach densities of
around 1.3g/cm3, which represents 1/6 that of steel, while their stiffness in terms of Young’s
15
modulus may be 5 times higher [58]. The highest measured tensile strength at CNT breaking is
recorded at 50 times larger than that of steel [57]. The thermal conductivity of CNT may
typically be very high in the axial direction, but very low in the lateral direction.
Depending on their structure, CNTs can also exhibit a very wide range of electrical
properties. They can be excellent conductors, with an electrical conductivity that is hundreds of
times higher than that of copper, they can be insulators, or semiconductors [58]. It is the
direction of the graphene sheet rolling with respect to the hexagonal lattice and the diameter of
the tube that are responsible for impairing such electrical properties. Groups of armchair
SWCNT have been demonstrated to exhibit metallic/conductive electrical characteristics, while
groups of zigzag and chiral SWCNT have been demonstrated to exhibit semiconductor properties
[59]. These properties may be varied through variations of the structural and diameter
characteristics.
Generally, CNT present good chemical and environmental stability but at the same time
they are versatile enough to be biocompatible with wide classes of enzymes and form bonds to
other chemical compounds in synergistic compounds that could enhance their properties as well
as the one of the enzymes in the enzyme-CNT conjugate [60].
4.3. CNT Production Methods
A high-quality production process aims at producing CNT that are free from both
structural and chemical defects, especially the ones located along their tubular axis. While
numerous methodologies have been investigated and significant progress has been made, CNT
synthesis still faces the following major challenges [58]: 1) low-cost mass production; 2) strict
control and guarantees of final product consistent properties; 3) adequate location and orientation
of CNTs on substrate materials; 4) understand and control of the CNT growth mechanisms.
To produce CNTs, three main elements are necessary: a source of carbon, a catalyst, and
a sufficient amount of energy [61]. Typically, carbon atoms or groups of atoms are obtained
from a source through energy injection; then carbon particles are recombined in the presence of a
catalyst to generate the CNT of a specific structure and size. The most widely used methods for
CNT generation are: arc discharge, laser ablation, chemical vapor deposition, flame synthesis,
and ball milling. Specific details about these methods are included below:
16
The arc discharge method [62] relies on the vaporization of a carbon electrode and
carbon deposit on a second one due to a high temperature discharge between the two electrodes
in an enclosure filled with inert gas at low pressure or liquid nitrogen. This is the most common
and easiest way to produce CNTs; however, it requires subsequent purification.
The laser ablation method [63] uses a laser instead of an electrical arc to produce the
large temperatures necessary for inducing the carbon atoms re-organization into CNTs.
The chemical vapor deposition method [64] consists of the decomposition of volatile
carbon compounds in the presence of metallic catalysts at high temperature. Methane, carbon
monoxide, hydrocarbons, and alcohols have been the most frequently used sources of carbon. A
variety of approaches have been investigated to enhance the process through the use of plasma,
lasers, and combined action of catalysts and supports [58].
The flame synthesis method consists of inducing the formation of CNT in premixed
flames [65]. The approach appears to allow for the production of large quantities of CNT at low
costs.
The ball milling method [66] generates CNT through a combined mechanical and thermal
process from elemental graphite powders. Milling the powder is assumed to produce nanotube
nuclei that grow during the subsequent annealing process.
Many of these CNT generation methods require some type of product “purification”
which generally consists of separation of nanotubes from the added catalysts, process byproducts or contaminants. The largely used techniques used for CNT purification include
oxidation, acid treatment, annealing, sonication, and filtering.
4.4. CNT Functionalization
As previously mentioned, the diverse properties of the CNTs have made them attractive
for a variety of applications. However, their versatility and implementation in such applications
requires application-dependent specificity, in other words, the functionality of the CNT must be
tuned, enhanced, or extended to support a particular requirement. Examples of techniques to be
applied include modifications of CNT solubility, proper dispersion, unbundling, selective
chemical reactivity, enhancement of mechanical and electrical properties, or biocompatibility
with specific enzymes.
17
CNT functionalization methods [67] are classified into chemical (or covalent) and
physical (or non-covalent) methods. They may affect specific areas of the nanotubes such as the
interior or exterior of the walls or the open ends. Covalent functionalization forms covalent
bonds between the carbon atoms of the CNT and the functional entities involved in the process.
They are likely to result in a functionalized CNT whose physical properties may be significantly
different than the original or pristine tubes. In some instances, this may produce undesirable
effects and thus a non-covalent method may be preferable in order to maintain their structure of
unaffected or unchanged.
Due to the high potential for applications in the bio-medical field, bio-functionalization
of CNTs is currently investigated quite extensively [66]. DNA, proteins, polypeptides,
carbohydrates, and others have been used to prepare biocompatible CNT. The functionalization
of CNTs, when used in CNT-enzyme conjugates may still affect the catalytic properties of the
enzyme [68, 69].
4.5. Enzyme Immobilization onto CNT
Once attached to a solid support, enzymes were shown to typically exhibit higher
physical stability and reusability, however changes in their catalytic properties usually gated
towards reduced functionality have also been observed. Generally, the ideal support for enzyme
immobilization should have the capability to immobilize large amounts of enzyme while
ensuring their optimal functionality. The support should also be chemically stable (i.e., not
degrade or deteriorate) and should not negatively affect the catalytic properties of the enzyme
(i.e., not interact with the enzyme to change or modify its chemical structure and thus its
functionality). Lastly, the support should be inert to microbial contamination such that the
immobilized enzyme remains unaltered and active for longer periods of time [70].
CNTs can fit such support-profile-based requirements through their large contact surfaceto-volume ratio and physico-chemical properties as highlighted in the previous sections.
However, CNT characteristics and their influences on the enzyme functionality and stability are
still to be investigated. Different methods using CNT as support for enzyme immobilization have
been tested. For instance, successful reports of enzyme immobilization on CNTs include direct
18
physical absorption [71], absorption on CNTs functionalized with polymers [72], direct covalent
binding [73], covalent binding with linking molecules [74], and entrapment [75].
19
Chapter 5
Enzyme Functionality if a Function of the Enzyme-Nanotube Interface
Note: All the results and discussion presented next are part of the research I was involved
with in Dr. Dinu’s lab at West Virginia University. These results have been published (please see
references [11, 76], with the two articules being included in the Appendix). I was also included
as an author on these two articles for my direct contribution.
This part of the lab work presented next aimed at understanding the effects of the CNT
onto enzyme characteristics (such as activity and functionality); such effects were investigated
upon enzyme immobilization onto both single wall and multi wall carbon nanotubes (SW and
MWCNTs respectively). Concrete examples of the immobilization techniques being used are
provided. Aspects related to the conditions that need to be chosen in order to preserve enzyme
functionality at the CNT interface are also being discussed, especially in the context of two
different model enzyme and two different types of supports, i.e., enzyme and supports with
different physical and chemical characteristics.
5.1. Material and Methods
5.1.1. Acid Oxidation of CNTs
Commercial SWCNTs (85% purity, Unidym Inc.) and MWCNTs (95% purity, Nanolab
Inc.,) were incubated in a concentrated sulfuric (96.4%, Fisher, USA) and nitric acid (69.5%,
Fisher, USA) mixture in a ratio of 3:1 (V/V). The CNTs/acids mixture was subsequently
sonicated in an ice bath (Branson 2510, Fisher, USA) for 3 or 6 h, at a constant temperature of
23°C. When the required time elapsed, CNTs/acids mixture was diluted with deionized (di)
water and filtered through a GTTP 0.2 µm polycarbonate filter membrane (Fisher, USA). Several
cycles of resuspension in di water were employed to remove acidic residues or catalysts. The
CNTs were isolated on the filter, subsequently dried in a vacuum desiccator and stored at room
temperature for further use.
20
5.1.2 Energy Dispersive X-ray Analysis (EDX) of CNTs
Energy dispersive X-ray analysis (EDX) was used for quantitative elemental analysis of
pristine and acid oxidized CNTs. For such analysis, samples (1 mg/ml in di water) were
deposited on silica wafers and dried under vacuum. The experiments were performed on a
Hitachi S-4700 Field Emission Scanning Electron Microscope (USA) with a S-4700 detector
combining secondary (SE) and backscattered (BSE) electron detection (all in a single unit),
operating at 20 kV. Results are presented as a percent of elements relative to the most dominant
element.
5.1.3. Scanning Electron Microscopy (SEM) of CNTs
Samples (1 mg/ml in di water of both pristine and acid treated CNTs) were dried on silica
wafers under vacuum and imaged using a Hitachi S-4700 Field Emission Scanning Electron
Microscope (USA) with a field emission gun at 10 kV.
5.1.4. Raman Spectroscopy of CNTs
Raman spectroscopy was performed on a Renishaw InVia Raman Spectrometer, CL532100, 100 mW, USA and allowed determination of the chemical structure and any modifications
resulted from the acids oxidation of both pristine and acids treated CNTs. Briefly, CNTs were
deposited on glass slides (Fisher, USA) were excited through a 20 X microscope objective using
an Argon ion (Ar+) laser beam with a spot size of < 0.01 mm2 operating at 514.5 nm. Detailed
scans were taken in the 100 to 3200 cm-1 range; low laser energy (i.e., < 0.5 mV) and exposure
time of 10 sec were used to prevent unexpected heating effects.
5.1.5. CNTs Solubility Measurement
The solubility of CNTs (pristine and acids oxidized CNTs) was evaluated in both di water
(pH 6.25) and Phosphate Saline Buffer (PBS, pH 7, 100 mM ionic strength). Briefly, CNTs were
diluted in the solvent of interest to yield to a 3 mg/ml solution. The suspension was then
21
centrifuged at 3000 rpm for 5 min; subsequently, part of the supernatant (0.8 mL) was removed
and filtered through a 0.2 µm GTTP filter membrane.
The filter membrane was then dried under vacuum and the amount of CNTs was
weighted. The solubility of the CNTs was calculated based on the volume used for suspension
and the initial starting amount.
5.1.6. Functionalization of CNT with Enzyme
•
Enzyme Immobilization by Physical Adsorption
One mg/mL enzyme, soybean peroxidase (SBP, Bioresearch, USA) or 0.5 mg/mL
chloroperoxidase (CPO, Bioresearch, USA) solution was prepared in phosphate-buffered saline
(PBS, 100mM, pH7, Sigma Aldrich) for SBP and citric acid buffer (CAB 50mM, pH 4.8, Sigma
Aldrich) for CPO.
For physical binding 2 mg of 3 or 6 h acid treated SW or MWCNTs were first dispersed in 2
mL of the enzyme solution prepared as previously described. The mixture was incubated with
shaking at 200 rpm for 2 h at room temperature. The immobilized enzyme was recovered by
filtration on the GTTP 0.2 µm polycarbonate filter membrane. The supernatant was isolated;
subsequently, the conjugates on the filter were washed at least six times (1 mL for each wash) to
remove loosely bound enzyme. The supernatant and the first two washes were kept and used to
determine the concentration of the enzyme. The supernatant and the first two washes were kept
and used to determine the concentration of the enzyme being washed out.
•
Enzyme Immobilization by Covalent Bonding
SBP or CPO were covalently attached to 3 or 6 h acid treated SW or MWCNTs using 1ethyl-3-[3-dimethylaminopropyl] carbodiimide hydrochloride (EDC; Acros Organics, USA) and
N-hydroxysuccinimide (NHS, Pierce, USA) [32]. Briefly, 2 mg CNTs (SW and MWCNTs) were
dispersed in 160 mM EDC and 80 mM NHS (total volume of 2 mL in MES (2-(N-morpholino)
ethane sulfonic acid sodium salt, 50 mM, pH 4.7, Sigma, USA) for 15 min at room temperature
with shaking at 200 rpm. The activated SWCNTs and MWCNTs were next filtered through the
22
0.2 µm GTTP filter membrane, washed thoroughly with the appropriate buffer to remove any
ester residues, immediately dispersed in 2 mL of 1 mg/mL SBP solution in PBS (100 mM, pH
7.0) or 0.5mg/mL CPO in CAB and incubated for 3 h at room temperature with shaking at 200
rpm. The resulting SBP/CPO-SW or MWCNT conjugates were filtered and washed extensively
with the correspondent buffer at least 6 times to remove any unbound enzyme. The supernatants
and two washes were collected to quantify enzyme loading.
5.1.7. CNTs Length and Morphology Assessment
An atomic force microscope (AFM, Asylum Research, USA) was used to evaluate the
length of pristine and acids treated CNTs. For this, the Si tip (Asylum Research, 50-90 KHz
AC240TS, USA) helped perform tapping mode in air. CNTs samples (i.e., pristine, 3 or 6 h acids
treated SW and MWCNTs) were dispersed in di water (to yield solutions of 0.1 mg/ml
concentration), deposited on mica surfaces (9.5 mm diameter, 0.15- 0.21 mm thickness, Electron
Microscopy Sciences, USA) and allowed to dry overnight under vacuum. Scan images of 10, 5
or 1 (µm x µm) areas were acquired. For each sample, at least 30 individual CNTs were counted
and measured to obtain average length distribution.
Sample morphology upon enzyme immobilization was also investigated; briefly, 0.1
mg/ml concentration of solutions containing enzyme-CNTs conjugates (both SW and MWCNTs)
have been deposited on the mica surfaces and allowed to dry overnight under vacuum. Scan
images of 10, 5 or 1 (µm x µm) areas were acquired to evaluate enzyme binding to the CNTs.
Height distribution measurements have also been collected to demonstrate enzyme attachment
onto the nanosupport being studied.
5.1.8. Enzyme Loading
The amount of SBP/CPO attached to SWCNTs and MWCNTs (i.e., SBP or CPO loading)
was determined using standard BCA assay kit (Pierce, USA) and subtracting the amount of
enzyme washed out in the supernatant and the first two washes from the initial amount of
enzymes added to the CNTs, during immobilization process. Briefly, the working reagent (1000
μL) was prepared by mixing 50 parts of reagent A with 1 part of reagent B (the reagents are
23
provided with the kit). The mixture of reagents A and B was further added to 50 μL solutions of
enzymes containing samples (i.e. the samples isolated in the form of the supernatant and washes
respectively). The resulting solutions were incubated at 37oC for 30 min. Absorbance at 562 nm
was determined on a spectrophotometer (Fisher, USA). Control calibration curves were prepared
by serial dilutions of the enzyme (free in solution) into the working reagent.
5.1.9. Enzyme Activity Assay
Enzyme retained specific activity was determined using colorimetric reactions monitored
on a UV-Vis spectrophotometer (Thermo Scientific EVO300). The specific activity was
calculated by comparing the activity of immobilized enzyme to the activity of free enzyme in
solution when enzymes where used at the same amount. Specifically, the activity of SBP was
determined by monitoring the oxidation of (2,2’-Azinobis[3-ethylbenzothiazoline-6-sulfonic
acid]) (ABTS, Sigma Aldrich) by SBP in the presence of H2O2 (Sigma Aldrich) at 412 nm.
Briefly, 20 µL of the SBP solution to be tested (free or immobilized) was added to 650 µL of
0.25 mg/mL ABTS and mixed in a plastic cuvette. Next, 20 µL of 6.5 mM H2O2 was added to
initiate the reaction and the cuvette was immediately placed in the spectrophotometer; the rate of
absorbance change was monitored for 2 min. The initial reaction rate was calculated from the
time-course slope and reported in µM µg-1 s-1.
The activity of CPO was determined by monitoring the conversion of 2-chloro-5,5dimethyl-1,3-cyclohexanedione (monochlorodimedon, Alfa Aesar) to dichlorodimedon by CPO
in the presence of Cl- and H2O2 at 278 nm. Briefly, 500 µL of CAB, 440 µL of 227.27 mM NaCl
(ACROS), 20 µL of 5 mM monochlorodimedon, and 20 µL of the CPO sample to be tested were
first mixed in a quartz cuvette. Then, 20 µL of 50 mM H2O2 was added to initiate the reaction
and the cuvette was immediately placed in the spectrophotometer and rate of absorbance change
monitored for 2 min. The initial reaction rate was calculated from the time-course slope and
reported in M µg-1 s-1.
The activity of the immobilized enzyme is reported as specific activity relative to free
enzyme activity. The activity of the free enzyme was determined using an equivalent amount of
free enzyme (based on loading data) and the protocol provided above.
24
5.2. Results and Discussion
Using liquid phase oxidation in strong sulfuric and nitric acids mixture, both SWCNTs
and MWCNTs functionalization is reported. Sonication at room temperature is known to attack
the graphene sheets on the C-C bands, introducing defects and oxidizing the CNT at their defect
sites thus resulting in shorter nanotubes and O-related functional groups used subsequently to
immobilize two model enzymes, namely soybean peroxidase (SBP) and chloroperoxidase (CPO).
Alan S. Campbell, Chenbo Dong, Fanke Meng, Jeremy Hardinger, Gabriela Perhinschi,
Nianqiang (Nick) Wu, and Cerasela Zoica Dinu investigated the effects of acids treatment onto
the structure and morphology of the CNTs. They confirmed the presence of the O-related
functionalities, as well as the removal of catalysts upon the employment of such acid treatment.
The results have been reported using well-implemented techniques namely using scanning
electron microscopy (SEM), atomic force microscopy (AFM), energy dispersive X-ray analysis
(EDX) and Fourier transform infrared spectroscopy (FTIR).
Selected results as reported by Campbell et al., are shown in Figure 3 [76]. Specifically,
FTIR confirmed the insertion of O-related functionalities onto both SW and MWCNTs (Figure
4a, and b respectively). The observed 3450 cm-1 peak was assigned to the hydroxyl moiety while
the ~2900 cm-1 peak was assigned to the stretching modes of the C-H groups. The 1750 cm-1
band corresponded to the C=O bond in the carbonyl and carboxylic moiety while the bands at
1550~1660 cm-1 were associated with the carbon-carbon bonds. Lastly the bands in the range of
1300-950 cm-1 were characteristic to the carbon-oxygen bond formation, thus further confirming
the presence of large amounts of hydrated surface oxides and O-related functionalization upon
the investigated acid-based treatment functionalization.
Further, a selected example of the chemical composition of pristine and acid treated
CNTs as investigated using EDX is shown in Figure 4 (reproduced after Dong et al., Applied
Surface Science 264 (2013) 261– 268, [11]). Results are plotted of X-ray counts vs. energy (in
KeV). The results revealed a high contents of carbon (C), oxygen (O) and iron (Fe) as metal
catalysts in both SW and MWCNTs. The presence of other elements such as Al, Si, Cl, S was
also detected, however such elements appeared in much lower amounts. Analysis showed that
the Fe content was generaly higher in the SWCNT sample when compared to the MWCNT one,
25
this being due to the fact that the purity of pristine (as purchased) SWCNTs was 85% compared
with 95% of the MWCNTs (both values were provided by the manufacturer).
Figure 3. (Reproduced after Campbell et al., Figure 1, ACS Appl. Mater. Interfaces
2014, 6, 5393−5403; [76]). Analysis of functional groups of the carbon-based materials. FTIR
and EDX spectra analysis of a) pristine and acid functionalized SWCNTs, b) pristine and acid
functionalized MWCNTs.
26
a)
b)
Figure 4. Reproduced after Dong et al., Applied Surface Science 264 (2013) 261– 268, [11].
(a) EDX elemental analysis of pristine SWCNTs; and b) MWCNTs. The inlets show the changes
in the contents of O and Fe with the time-controlled acid treatment.
As it can be seen in the inlets in Figure 4 reproduced after after Dong et al., Applied
Surface Science 264 (2013) 261– 268, [11], the Fe content decreased with the acid treatment
time for both SW and MWCNT samples, thus demonstrating the efficient removal of the metal
catalyst, and further confirming free O-related functional groups formation. Analysis also
27
showed that the removal of Fe was higher for the SWCNT sample relative to the MWCNT one,
presumably due to its lower purity (as stipulated in the Materials and Methods section and based
on the manufacturer characteristics provided to our lab). The O content increased with the acid
treatment time presumably due to more O-related groups being formed at the defect sites after an
extended acid incubation period when compared with a shorter incubation time. The decrease of
all other elements was due to impurities being removed by acid treatment and the shortening of
the nanosupports that is known to be leading to the formation of amorphous carbon.
Figure 5. Reproduced after Dong et al., Applied Surface Science 264 (2013) 261– 268, [11].
SEM images of a) pristine SWCNTs, b) pristine MWCNTs, c) 6 h acids treated SWCNTs, d) 6 h
acids treated MWCNTs; the scale bar is 1 µm.
The typical morphologies of SWCNTs and MWCNTs respectively were investigated by
SEM and are shown above. Analysis reprinted after Dong et al., Applied Surface Science 264
28
(2013) 261– 268, [11] did not show any significant morphological changes between the acid
treated samples when compared to their pristine counterparts (Figure 5).
AFM, in tapping mode allowed morphology and length analysis of both pristine and acid
treated CNT samples. The study of Campbell et al. (Campbell et al., Figure 1, ACS Appl. Mater.
Interfaces 2014, 6, 5393−5403; [76]) showed that acid treatment reduced SWCNTs length from
760 ± 276 nm to 516 ± 277 nm, while the length of MWCNTs was reduced from 6,049± 2,954
nm to 452 ± 213 nm, both after 6 h of nanotubes incubation in the nitric and sulfuric acids
mixture. A representative AFM scan of such sample is shown in Figure 6 (please note that these
are analysis performed in parallel to the Campbell et al., Figure 1, ACS Appl. Mater. Interfaces
2014, 6, 5393−5403; [76] and not previously published).
Figure 6. AFM pristine SWCNTs acid treated for 6 h.
Having established that the acid functionalization influences both the chemical and
physical properties of the CNTs, we proceeded to assess whether such functionalization will also
29
influence CNTs biocompatibility. For this purpose, the SBP and CPO were immobilized on SW
and MWCNTs respectively, either through physical or covalent binding (Figure 7).
Defect
Acid functionalization
O-related groups
(a)
SW or
Enzyme
(b
Physical
Adsorption
EDC/NHS
Activation
Enzyme-CNT conjugate
Ester Functionalized Groups
(c)
Covalent
Binding
Enzyme-CNT conjugate
Figure 7. General mechanism of protein immobilization onto nanosupports of carbon nanotubes.
(a) Carbon nanotube (either SW or MWCNT) under acid treatment. (b) Physical adsorption of
enzymes onto nanosupports. (c) Covalent binding of enzymes onto nanosupports using EDC and
NHS chemistry as highlighted in the materials and methods.
The two model enzymes are heme enzymes and were selected based on their different
chemical caracteristics as well as their extended applications in synthetic environment.
Specifically, SBP is an anionic monomeric glycoprotein (pI 3.9) with a molecular weight of ~40
kDa known for its unusual thermostability and a high oxidation potential, while CPO is a
monomeric enzyme with a molecular weight ~42 kDa known for its catalitic contribution in
halogenation reaction, except fluorination. Specific applications of these enzymes were also
listed earlier in this report.
Morphology analysis of the enzyme-CNTs conjugates were also performed using AFM,
with representative images being shown in Figure 8.
30
a)
b)
Figure 8. a) Functionalized SWCNTs with SBP; b) Height profile of the enzyme-SWCNT
conjugates. The line was taken across the highlighted nanotube in image and shows a bead-like
morphology in which the beads were immobilized enzymes.
31
The amount and activity of SBP or CPO immobilized through either physical or covalent
binding onto the nanosupport were quantified using standard spectroscopic assays (see
description in the Materials and Methods section). Both the loading and activity are reported in
terms of mean ± standard deviation and the data is averaged over at least 5 samples in order to
ensure relevant statistics (Table 1).
The specific retained activity of the immobilized SBP varied greatly with the nanosupport
being tested. SBP showed the highest activity when immobilized onto MWCNTs using covalent
binding (about 28 % retained specific activity relative to free enzyme activity). The highest
activity for the physically bound SBP was also observed for the enzyme immobilized onto
MWCNTs (about 25 % of the free enzyme); however, the same method of immobilization
allowed retention of only about 15 % specific activity onto SWCNTs.
Table 1: SBP Loading and Activity Data. Reproduced after Campbell et al., ACS Appl. Mater.
Interfaces 2014, 6, 5393−5403; [76].
Nanosupport
(Immobilization method)
SWCNTs
(Physical)
SWCNTs
(Covalent)
MWCNTs
(Physical)
MWCNTs
(Covalent)
Loading
(mg enzyme /
mg nanosupport)
Specific
Retained
Activity (%)
0.19 ± 0.03
14.81 ± 6.77
0.08 ± 0.02
4.38 ± 1.49
0.15 ± 0.05
25.28 ± 4.04
0.24 ± 0.10
28.01 ± 5.01
Campbell et al. complementary studies confirmed that MWCNTs nanosupports provided
the optimum nanointerfaces to preserve the selected enzymes catalytic behavior and activities.
Specifically, CPO bound to MWCNTs expressed retained specific activities of up to 29%, and
49% for physical adsorption, and covalent binding respectively (Table 2). These activities were
32
approximately 27% and 46% higher for each respective immobilization method when compared
to the activity of the same enzyme immobilized onto SWCNTs.
Table 2: CPO Loading and Activity Data. Reproduced after Campbell et al., ACS Appl. Mater.
Interfaces 2014, 6, 5393−5403; [76].
Nanosupport
Loading
(Immobilization
(mg enzyme /
method)
mg nanosupport)
SWCNTs
(Physical)
SWCNTs
(Covalent)
MWCNTs
(Physical)
MWCNTs
(Covalent)
Specific Retained
Activity (%)
0.09 ± 0.02
1.49 ± 0.16
0.06 ± 0.01
2.06 ± 0.35
0.06 ± 0.01
28.81 ± 9.78
0.08 ± 0.02
48.84 ± 11.56
Based on the statistical analyses provided herein, covalent binding onto MWCNT
nanosupports have benefited CPO more than it benefited SBP, when considering both the
enzyme loading as well as its activity. This was presumably due to the higher ability of CPO to
bind away from its active site and thus lead to less conformational changes of this site even upon
enzyme binding to the nanotube [77-78]. Further, even though both enzymes have been
considered as models based on their known primary and secondary structures and further, even
though both enzymes have been treated similarly in terms of lab conditions being used for the
experiments related above, the individual net positive charge of CPO is known to be higher than
that of the SBP which will presumably lead to its different ability to bind to the MWCNT
structure. Further, at the working pH considered in this study i.e., CAB (pH 4.8) for CPO (pI 4.0)
when compared with PBS (pH 7) for SBP (pI 3.9), the SBP is being even more negatively
charged.
Complementary, previous analyses have showed that enzymes levels of activity also
33
depend on the radius of curvature that the enzyme is being immobilized onto [76]. In particular,
Campbell et al. proved that a higher radius of curvature of the nanosuports resulted onto a higher
center to center distance between two adjacent immobilized enzymes which was shown to be
beneficial for both enzyme loading and activity. Contrary to that, increased protein−protein
interactions caused by a less curved surface could result in a more dramatic activity loss over
time and in a harsh environment as authors have demonstrated when they considered graphene as
an example of a totally flat surface.
Lastly, it is known that the electrical behavior of SWCNTs as a result of their structure as
one graphene sheet is either semiconductor or metal. In the same time, the MWCNTs having a
multi graphene sheet structure, behave as metals if at least one of the sheets is a metal [79]. Due
to their difference in charge, the CPO is more compatible with the MWCNTs that exhibited a
stronger metallic nature thus limiting deformation of its more rigid active site. The SBP, which
have a higher negative charge will be more compatible with the SWCNTs however it will also
lead to a larger deformation of its more flexible binding site.
34
Chapter 6
Conclusions
Enzyme applications are currently limited by enzyme’s lack of stability in a wide range
of pH and high temperatures, therefore intense research has been conducted to overcome such
impediments. One area of research is in finding the optimum enzyme immobilization strategy to
ensure enzyme functionality for extended time and in harsh environments. As such, studies on
immobilization of enzymes on different supports have demonstrated that the support has to
exhibit specific properties, such as physical resistance to compression, hydrophilicity, inertness
toward enzymes, ease of functionalization, biocompatibility, resistance to microbial attack, and
availability at low cost. Further, the enzyme and the support must be compatible such that the
enzyme-support conjugate itself exhibits the desirable properties as listed above.
The investigation presented in this report defines a methodology that can be extended in
order to identify the best parameters and ideal conditions for synthetic applications of
biocatalyst-based conjugates. The investigation uses CNTs and proved that they are compatible
supports for immobilization of model enzymes thus opening the door for their wide
implementation in a variety of applications. From these fundamental studies, means of
implementation of enzymatic catalysis through CNTs immobilization could be derived and seen
as the next steps in designing efficient and effective processes and materials aimed at meeting
the goals of enzyme-based technologies as highlighted in the first chapters of this problem
report. In particular, to ensure high enzyme activity and functionality at nanotube interfaces, one
needs to control both the nanotube physico-chemical properties as well as consider the structure
and individual charge of the enzymatic system to be immobilized. Only manipulation of both
interfaces will lead to optimum catalytic efficiencies and further implementation of such
enzyme-based conjugates into both consumer and industrial applications. Such enzymatic
biocatalysis-based systems for instance, will not only allow for high product recovery since the
enzyme is not mixed with the product but rather immobilized, but further, they could have the
capability to be utilized to reduce the release of environmentally harmful molecules, while
producing revenues from otherwise inaccessible sources.
35
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43
Appendices
44
Research Article
www.acsami.org
Enzyme Catalytic Efficiency: A Function of Bio−Nano Interface
Reactions
Alan S. Campbell,† Chenbo Dong,† Fanke Meng,‡ Jeremy Hardinger,† Gabriela Perhinschi,†
Nianqiang Wu,‡ and Cerasela Zoica Dinu*,†
†
Department of Chemical Engineering and ‡Department of Mechanical and Aerospace Engineering, West Virginia University,
Morgantown, West Virginia 26506, United States
S Supporting Information
*
ABSTRACT: Biocatalyst immobilization onto carbon-based
nanosupports has been implemented in a variety of applications
ranging from biosensing to biotransformation and from decontamination to energy storage. However, retaining enzyme
functionality at carbon-based nanosupports was challenged by
the non-specific attachment of the enzyme as well as by the
enzyme−enzyme interactions at this interface shown to lead to loss
of enzyme activity. Herein, we present a systematic study of the
interplay reactions that take place upon immobilization of three
pure enzymes namely soybean peroxidase, chloroperoxidase, and
glucose oxidase at carbon-based nanosupport interfaces. The
immobilization conditions involved both single and multipoint
single-type enzyme attachment onto single and multi-walled
carbon nanotubes and graphene oxide nanomaterials with properties determined by Fourier transform infrared spectroscopy
(FTIR), energy dispersive X-ray analysis (EDX), scanning electron microscopy (SEM), and atomic force microscopy (AFM).
Our analysis showed that the different surface properties of the enzymes as determined by their molecular mapping and size work
synergistically with the carbon-based nanosupports physico-chemical properties (i.e., surface chemistry, charge and aspect ratios)
to influence enzyme catalytic behavior and activity at nanointerfaces. Knowledge gained from these studies can be used to
optimize enzyme−nanosupport symbiotic reactions to provide robust enzyme-based systems with optimum functionality to be
used for fermentation, biosensors, or biofuel applications.
KEYWORDS: enzyme immobilization, bio−nano interface, symbiotic behavior, catalytic tuning
■
INTRODUCTION
Enzymes are a naturally occurring class of proteins that possess
unique properties including high catalytic activity, selectivity, and
specificity. Enzymes are environmentally friendly and produce
fewer harsh byproducts than their chemical counterparts.1,2
Because of such properties, enzymes are now key players in
various industrial processes from waste treatment3,4 to food
processing5 and from biodiesel production6 to the petroleum
refining industry.7 More recently, enzyme-based conjugates
obtained by immobilization of enzymes onto nanoscale solid
supports have shown applicability in biosensing,8,9 drug
delivery,10 and decontamination.11,12 In particular, Besteman et
al. reported on the use of single-walled carbon nanotubes as
supports for immobilization of glucose oxidase for biosensing
applications,13 Luckarift et al. examined the use of biomimetic
silica supports for butyrylcholinesteras immobilization for flow
through reactors,14 and Fernandez-Lafuente et al. showed that
coupling immobilization and site-directed mutagenesis can
improve biocatalyst or biosensor performance,15 while Dinu et
al. reported on immobilization of enzyme perhydrolase S54 V
onto carbon nanotubes to be used for generating decontamination platforms.11
© 2014 American Chemical Society
In a favorable nanoenvironment, enzyme immobilization was
shown to lead to increased enzyme stability and improved
specificity1,2,16 and allowed for prolonged enzyme functionality
through chemical (e.g., cross-linking)17 and physical treatment
(e.g., pH enhancement or lyophilization).18 For instance,
immobilization onto carbon-based nanosupports was shown to
increase enzyme turnover and to allow for prolonged enzymebased conjugate usage.11,17,19 Nanosupport immobilization
studies have also proved that, while the high aspect ratio of the
nanosupports allows enzyme-based conjugate retention in
solution, multiple usages and ease of conjugate recovery via
filtration, the nonspecific binding of the enzyme at the
nanointerface can result in enzyme active site deformation (i.e.,
change in the active site conformation)19 and thus increased
enzyme−nanosupport interactions with subsequent reduced
enzyme activity.16,19−22 Future developments in enzyme-based
applications of enzyme-based-nano conjugates need to account
for increased enzyme functionality, high operational stability,
Received: June 28, 2013
Accepted: March 25, 2014
Published: March 25, 2014
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subsequently rediluted in 710 mL of DI water, preheated to 35 °C, and
incubated with 30 mL of 30% hydrogen peroxide (H2O2, Sigma
Aldrich). Finally, the solution was filtered and washed using DI water at
35 °C until the effluent was clear and the pH was kept constant at 6. The
resulting product was dried in a vacuum oven; the obtained brown
powder was stored at room temperature for future use.
Carbon-Based Material Acids Treatment. Functionalized
carbon-based materials (CMATs; SWCNTs, 85% purity, Unidym
Inc.; MWCNTs, 95% purity, NanoLab Inc.; or GON) were prepared via
acids treatment as previously described.50 Briefly, 100 mg of pristine
CMATs were added to a 60 mL mixture of 3:1 (V:V) H2SO4 and nitric
acid (HNO3, Fisher Scientific, 69.6%). The mixture was ultrasonicated
for 6 h (Branson 2510, Fisher Scientific) at a constant temperature of
approximately 23 °C. Next, the solution was diluted in DI water and
filtered through a GTTP 0.2 μm polycarbonate membrane (Fisher
Scientific). Several cycles of redispersion and filtration in DI water were
performed to remove acidic residues or catalysts and impurities. The
CMATs isolated on the filter membrane were dried in a vacuum
desiccator and stored at room temperature until use.
CMATs Characterization. Chemical structure, morphology, and
elemental composition of CMATs were investigated using Fourier
transform infrared spectroscopy (FTIR), scanning electron microscopy
(SEM), and energy-dispersive x-ray spectroscopy (EDX), respectively.
For FTIR, 2 mg pellets of samples were collected and analyzed under the
transmission mode by using KBr pellet on a Thermo Nicolet
Instrument. For SEM and EDX characterizations, samples (1 mg/mL
in DI water) were deposited on silica wafers and dried under vacuum.
Experiments were performed on a Hitachi S-4700 field emission
scanning electron microscope with a S-4700 detector combining
secondary (SE) and backscattered (BSE) electron detection (in a single
unit).
The length of pristine and acids treated SWCNTs and MWCNTs
were quantified using atomic force microscopy (AFM) and a silicon tip
(Asylum Research, 50-90 kHz AC240TS) operating in air tapping mode.
Briefly, nanotube samples in DI water (0.1 mg/mL) were deposited
onto mica surfaces (9.5 mm diameter, 0.15−0.21 mm thickness,
Electron Microscopy Sciences) and dried overnight under vacuum.
Scans of 10 μm × 10 μm and 1 μm × 1 μm areas were acquired.
To evaluate the CMATs’ degree of hydrophilicity/hydrophobicity,
dispersity tests were performed in DI water (pH 6.25), phosphate
buffered saline (PBS, 100 mM, pH 7, Sigma Aldrich), and citric acid
buffer (CAB 50 mM, pH 4.8, Sigma Aldrich). Briefly, CMATs were first
dispersed in each of the different solvents at a concentration of 3 mg/
mL. The suspension was subsequently centrifuged at 3000 rpm for 5 min
and 0.8 mL of the generated supernatant was removed and filtered
through the GTTP 0.2 μm polycarbonate filter membrane. The filter
was subsequently dried under vacuum, and the amount of CMATs on
the filter was weighed. Dispersity was calculated based on the volume
suspended, the initial amount used in the dispersion test, and the final
amount isolated on the filter paper.
Enzyme Immobilization. Soybean peroxidase (pure SBP, Bioresearch, Rz = 2), glucose oxidase (pure GOX, Type VII, Sigma, Rz = 1.3),
and chloroperoxidase (pure CPO, Bioresearch, Rz = 1.3) were
immobilized onto CMATs using either physical or covalent binding.
For physical binding 2 mg of CMATs were first dispersed in 2 mL of
enzyme solution (1 mg/mL in PBS for SBP, 0.5 mg/mL in PBS for GOX,
or 0.5 mg/mL in CAB for CPO) via brief sonication. The solution was
then incubated at room temperature for 2 h with shaking at 200 rpm.
Next, the enzyme−CMAT conjugates were recovered by filtration
through the GTTP 0.2 μm polycarbonate filter membrane. The
supernatant was isolated and its volume recorded. The conjugates
isolated on the filter were washed at least 6 times using the
corresponding buffer (2 mL for each wash) to remove loosely bound
enzyme, with the first two washes being isolated and their volumes
recorded. Finally, the conjugates were redispersed in 2 mL of their
corresponding buffer and stored at 4 °C.
For covalent binding, 2 mg of CMATs were first activated using 1ethyl-3-[3-dimethylaminopropyl] carbodiimide (EDC, Acros Organics)
and N-hydroxysuccinimide (NHS, Pierce) chemistry. Specifically,
CMATs were dispersed via sonication in 160 mM EDC and 80 mM
efficiency, yield of recovery and conversion, and reduced enzyme
inhibition. However, while previous examples show that a
significant amount of research has been directed towards
understanding the interactions of known enzymes with nanosupports, the molecular mechanisms and synergistic reactions
that take place at the nanosupport interface upon enzyme
immobilization have yet to be fully understood.
We hypothesized that fine control of the enzyme−nanosupport interface through the control of the enzyme immobilization process as well as nanosupport characteristics can lead to
enhanced enzyme catalytic efficiency. To test our hypothesis, we
used pure glycosylated enzymes with different properties (e.g.,
surface chemistry, molecular weight, isoelectric point, etc.),
nanosupports with different characteristics (both physical and
chemical), and different immobilization techniques (i.e., physical
or chemical). Specifically, soybean peroxidase (SBP), an anionic
monomeric glycoprotein (pI 3.9)23 with a molecular weight of
∼40 kDa24 known for its unusual thermostability and a high
oxidation potential, chloroperoxidase (CPO), a monomeric
enzyme with a molecular weight of ∼42 kDa,25 and glucose
oxidase (GOx), a homodimer flavoenzyme oxidoreductase with a
molecular weight of ∼180 kDa,26 were used as models. The
choice in enzymes was based on their extended applications with
SBP being used for diagnostics27−29 and waste-water treatment
industrial implementation;30−32 CPO for chiral organic synthesis,33−35 decontamination,36 and the petroleum industry,37,38
and GOX for biosensing,39,40 biofuel cell formation,41 and food
processing applications.42 The selected carbon-based nanosupports encompassed single-walled carbon nanotubes
(SWCNTs), multi-walled carbon nanotubes (MWCNTs), and
graphene sheets (GON) with different physical and chemical
properties as demonstrated by Fourier transform infrared
spectroscopy (FTIR), energy dispersive X-ray analysis (EDX),
scanning electron microscopy (SEM), and atomic force
microscopy (AFM). The choice of the nanosupports was based
on their extended implementation in a wide variety of
applications from biosensing40,43 to large-scale industrial
processing and waste remediation.44−46 Lastly, the chosen
immobilization techniques were aimed to offer different enzyme
attachment mechanisms at nanointerfaces, namely single or
multipoint attachment.47−49
Our systematic studies on the underlying mechanisms that
control enzyme activity and catalytic behavior at nanointerfaces
seek to reveal whether there is an optimum support to be used for
a specific enzyme immobilization in order to lead to maximum
catalytic efficiency of that enzyme. Discovering an optimum
strategy that could be used in the future when the formation of
bio−nano conjugate systems with increased enzyme functionality is considered can fill the gap in developing robust enzymebased systems with applications in fermentation, biosensoring, or
biofuel production.
■
MATERIALS AND METHODS
Graphene Oxide Nanosheet Synthesis. Graphene oxide nanosheets (GON) were produced from graphite powder (Alfa Aesar, 99.8%
purity). First, 10 g of the graphite powder and 5 g of sodium nitrate
(NaNO3, Sigma Aldrich, 99.0%) were added to 230 mL of concentrated
sulfuric acid (H2SO4, Fisher Scientific, 96.4%) in a 2000 mL flask; the
flask was subsequently placed in an ice bath and the mixture was stirred
slowly. A 30 mg portion of potassium permanganate (KMnO4, Sigma
Aldrich, 99.0%) was added slowly to the flask to ensure that the
temperature of the mixture remained below 20 °C. Next, the solution
was heated to 35 °C for 30 min, diluted in 460 mL of deionized (DI)
water, and again quickly heated to 98 °C for 15 min. The mixture was
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Figure 1. Characterization of carbon-based materials (CMATs). FTIR and EDX spectra analysis of (a) pristine and acid-functionalized SWCNTs, (b)
pristine and acid-functionalized MWCNTs, and (c) pristine and acid-functionalized graphene oxide nanosheets (GON). FTIR and EDX spectra
confirmed the presence of carboxyl (COOH) functionalizations upon acid mixture incubation of CMATs.
NHS in 2-(N-morpholino)ethanesulfonic acid sodium salt buffer (MES,
50 mM, pH 4.7) with a final volume of 2 mL and incubated at room
temperature for 15 min with shaking at 200 rpm. Subsequently, the
mixture was filtered through a GTTP 0.2 μm polycarbonate filter
membrane and washed thoroughly with MES buffer to remove any ester
residues. Next, the activated CMATs were immediately dispersed in 2
mL of the selected enzyme solution (consistent with physical binding)
and incubated at room temperature for 3 h with shaking at 200 rpm.
Enzyme−CMAT conjugates were then recovered and washed, with the
supernatant, and the two washes were recovered (consistent with
physical binding). Finally, the conjugates were redispersed in 2 mL of
the corresponding buffer and stored at 4 °C.
For covalent binding through a spacer, 2 mg of the selected CMATs
were first activated using EDC/NHS chemistry as previously described
(see covalent binding), subsequently dispersed in 5 mL of 1 mg/mL
Amino-dPEG8-COOH (PEG, 32.2 Å, Quanta Biodesign) in the
designated buffer, and incubated at room temperature for 3 h with
shaking at 200 rpm. The resulting conjugates were then filtered and
washed with their corresponding buffer. Finally, the selected enzyme
was attached to the PEG linker as previously described. After the time
elapsed, enzyme−PEG−CMAT conjugates were recovered and washed,
and the supernatant and the two washes were recovered to quantify the
enzyme loading (consistent with physical and covalent binding).
Conjugates were redispersed in 2 mL of their corresponding buffer and
stored at 4 °C.
Covalent binding was confirmed by incubating the enzyme-carbonbased conjugates in 1 M NaCl solution for 10 min at 200 rpm; upon
incubation, the conjugates were filtered using the GTTP 0.2 μm
polycarbonate filter membrane and washing thoroughly with their
corresponding buffers. The resulting supernatant and washes were
recovered to evaluate any enzyme removal.
Enzyme Loading onto CMATs. The amount of the immobilized
enzyme relative to the amount of CMATs being used (i.e., the enzyme
loading) was estimated using standard BCA Assay (Pierce) and
subtracting the amount of enzyme washed out in the supernatant and
the first two washes (see above) from the initial amount of enzyme
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added during the immobilization process. Briefly, 1 mL of working
reagent containing 50 parts reagent A with 1 part reagent B (reagents
were provided stock with the BCA Assay kit) was mixed with 50 μL of
enzyme solution (either from the supernatant or the washes) and
incubated at 37 °C for 30 min. Absorbance at 562 nm was recorded for
each sample using a UV−vis spectrophotometer (Thermo Scientific
EVO300) and compared to a calibration curve of known concentrations
of the respective enzyme (free in solution) in the working reagent.
Loadings were estimated as the difference between the amount of
enzyme washed out from the initial amount of enzyme added during the
incubation relative to the amount of CMATs being used.
Determine the Specific Retained Activity of the Enzyme
Immobilized Onto CMATs. Immobilized enzyme retained specific
activity was determined using colorimetric reactions monitored on a
UV−vis spectrophotometer (Thermo Scientific EVO300). The specific
activity was calculated by comparing the activity of immobilized enzyme
to the activity of free enzyme in solution at the same amount.
Specifically, the specific activity of SBP was determined by monitoring
the oxidation of 2,2′-azinobis[3-ethylbenzothiazoline-6-sulfonic acid]
(ABTS, Sigma Aldrich) by SBP in the presence of H2O2 (Sigma Aldrich)
at 412 nm. Briefly, 20 μL of the SBP solution to be tested (free or
immobilized) was added to 650 μL of 0.25 mg/mL ABTS and mixed in a
plastic cuvette. Next, 20 μL of 6.5 mM H2O2 was added to the mixture to
initiate the reaction, and the cuvette was immediately placed on the
spectrophotometer; the rate of absorbance change was monitored for 2
min. The initial reaction rate was calculated from the time-course slope
and reported in micromolar per microgram second.
For the specific activity of GOX, 400 μL of PBS, 250 μL of 0.25 mM
glucose (Across), 250 μL of 0.25 mg/mL ABTS, and 50 μL of 0.5 mg/
mL SBP were first mixed in a plastic cuvette. Then, 50 μL of the GOX
solution to be tested was added to initiate the reaction and the cuvette
was immediately placed in the spectrophotometer; the rate of
absorbance change was monitored for 2 min. The initial reaction rate
was calculated from the time-course slope and reported in micromolar
per microgram second.
The specific activity of CPO was determined by monitoring the
conversion of 2-chloro-5,5-dimethyl-1,3-cyclohexanedione (monochlorodimedon, Alfa Aesar) to dichlorodimedon by CPO in the
presence of Cl− and H2O2 at 278 nm. Briefly, 500 μL of CAB, 440 μL of
227.27 mM NaCl (ACROS), 20 μL of 5 mM monochlorodimedon, and
20 μL of the CPO sample to be tested were first mixed in a quartz
cuvette. Then, 20 μL of 50 mM H2O2 was added to initiate the reaction
and the cuvette was immediately placed in the spectrophotometer; the
rate of absorbance change was monitored for 2 min. The initial reaction
rate was calculated from the time-course slope and reported in
micromolar per microgram second.
Enzyme Kinetic Parameters Determination. The kinetic
parameter, Km (where Km is the Michaelis−Menten constant in
micromolar), Vmax (where Vmax represents the maximum rate of reaction
in micromolar per microgram second), and kcat (enzyme turnover, 1/s),
values of the free and immobilized enzyme were determined by
measuring the initial rates of reaction in the respective activity assays (as
described above), with varying substrate concentrations and using
nonlinear regression. Specifically, for SBP the concentration of H2O2
was varied from 0 to 0.04 mM, for GOX the concentration of glucose was
varied from 0 to 100 mM, and for CPO the concentration of H2O2 was
varied from 0 to 4 mM.
Statistical Analysis. All results are presented as mean ± standard
deviation with at least six trials for each conjugate.
properties of the pristine carbon-based materials (CMATs), we
used Fourier transform infrared spectroscopy (FTIR), energy
dispersive X-ray analysis (EDX), scanning electron microscopy
(SEM), and atomic force microscopy (AFM).
Our FTIR analysis showed that acids treatment led to grafting
of carboxyl (COOH) functionalities onto all the CMATs being
tested. Specifically, the analysis of the chemical structure of both
SWCNTs and MWCNTs (Figure 1a and b, respectively) showed
a peak at 3450 cm−1 corresponding to the hydroxyl moiety and a
∼2900 cm−1 peak corresponding to the stretching mode of C
H groups. The 1750 cm−1 band corresponded to the CO bond
in the carbonyl and carboxylic moiety while the bands at 1550−
1660 cm−1 were associated with the CC bonds formation.51
The bands in the 1300−950 cm−1 range were characteristic of
CO bond formation and, thus, confirmed the presence of large
amounts of hydrated surface oxides and CMATs-COOH
functionalization. The FTIR spectrum of the GON is shown in
Figure 1c. The large peak in the 3400−3200 cm−1 range is
indicative of formation of hydroxyl groups at the surface of the
GON.52 The peak at ∼1740 cm−1 is a result of the CO bonds
in the COOH groups as well as in carbonyl moieties, while the
∼1620 cm−1 peak confirmed the presence of CC bonds
resulted from unoxidized regions of the graphene. Finally, the
large band at 1400−1060 cm−1 confirmed the presence of
COOH groups in epoxy or alkoxy groups formed at the surface of
the CMATs.52
EDX analysis (Figure 1, table format) further confirmed
COOH functionalization of CMATs; specifically, the O content
increased in the acids-treated CMATs while the C and other
elements content decreased.53 The decrease in other elements
was a result of metal catalyst residues and other impurities being
removed upon acid treatment as well as nanosupports being
shortened thus leading to the formation of amorphous carbon.50
To confirm the shortening of the nanotubes we compared
COOH-functionalized SWCNTs and MWCNTs with their
pristine counterparts using tapping mode AFM. Our analysis
showed that acid treatment reduced the length of SWCNTs from
760 ± 276 to 516 ± 277 nm and the length of MWCNTs from
6049 ± 2954 to 452 ± 213 nm. The diameters of the nanotubes
were however unaffected by the treatment; similarly, the
dimensions of the GON were maintained constant. Further,
SEM showed no significant morphological changes for the acidtreated samples when compared to their pristine counterparts
(Supporting Information Figure S1). Our results are in
agreement with previous studies, which showed that liquid
phase oxidation with a strong acid mixture introduces structural
changes and adds free COOH groups to nanomaterials.50,51,53
Carboxyl functionalization upon acids treatment improved
CMAT dispersity in several solvents (Supporting Information
Table S1). Specifically, in DI water (pH 6.25), the dispersity of
SWCNTs, MWCNTs, and GON improved by 9.3-, 6.8-, and 6.5fold, respectively. Similarly, in PBS (pH 7) the dispersity of
SWCNTs, MWCNTs, and GON improved by 4.8-, 3.8-, and 1.4fold. Further, in CAB (pH 4.8) the dispersity of SWCNTs,
MWCNTs, and GON improved by 8.3-, 9.3-, and 13.5-fold
relative to their pristine counterparts. The increase in dispersity
upon acids treatment is attributed to the increase in the number
of COOH groups and thus increased carboxylate anion
formation through deprotonation of these groups in waterbased environments.54 The poor dispersion observed at lower
pH values (i.e., in CAB) can be attributed to the aggregation of
CMATs through H bonding in these conditions.55 The acidstreated MWCNTs and GON had the lowest dispersity in PBS
■
RESULTS AND DISCUSSION
Morphology and Structure Characterization of Carbon-Based Materials (CMATs). Pristine SWCNTS (diameter
= 0.8−1.2 nm, length = 760 ± 276 nm), pristine MWCNTs
(diameter = 10−20 nm, length = 6049 ± 2954 nm), and pristine
GON (sheets of 500−5000 nm) were acids treated by incubation
in a nitric and sulfuric acids mixture for 6 h50 to generate
nanosupports with different characteristics. To investigate
whether acids treatment changed the physical and chemical
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Figure 2. Concept schematic of soybean peroxidase (SPB) immobilization onto CMATs. (a) SBP catalyzes the oxidation of ABTS to ABTS+. (b) SPB
immobilization onto CMATs with different surface curvatures and aspect ratios led to enzyme-based conjugates.
enzyme loading is given in Figure 3a. Our data showed that the
specific retained activity of the immobilized SBP varied
significantly with the nanosupport being tested. In particular,
SBP retained the highest specific activity when immobilized onto
MWCNTs using covalent binding (about 28% retained specific
activity relative to the free enzyme) while the lowest specific
activity was displayed by the enzyme immobilized onto GON
using covalent binding with PEG linker (about 1.6 % of the
specific activity of the free enzyme). The highest specific activity
for the physically bound SBP was observed for the enzyme
immobilized onto MWCNTs (about 25 % of the specific activity
of the free enzyme); however, the same immobilization method
allowed retention of only about 15% and 1.7% specific activity
onto SWCNTs and GON, respectively. Covalent immobilization
also yielded to only about 4% of the specific activity of the free
enzyme activity both onto SWCNTs and GON nanosupports,
while covalent binding through the PEG linker led to the highest
specific retained activity for the enzyme immobilized onto
MWCNTs (about 20% of the specific activity of the free enzyme)
and only about 8% and 1.6% onto SWCNTs and GON,
respectively. Control experiments have been also performed to
validate the feasibility of the covalent binding. Specifically, the
enzyme-carbon-based nanosupports have been incubated in high
salt concentrations known to remove the enzymes bound
through nonspecific electrostatic interactions; subsequent
evaluation of the enzyme loading showed that such high salt
incubation removed <3% of the immobilized enzyme. However,
upon such incubation, the remaining immobilized enzyme lost
about 70% of its initial activity possibly due to the accelerated
covalent multipoint attachment.58
Enzyme catalytic behavior at the different CMAT nanointerfaces was assessed under varying concentrations of hydrogen peroxide (Figure 3b−d); the kinetic parameters Vmax (i.e., the
presumably due to the higher ionic strength of this buffer that
could have induced aggregation of their carboxylated anions.55
This effect was not observed for SWCNTs since these nanotubes
have a reduced number of defects and thus a lower rate of COOH
functionalization relative to both MWCNTs and GON.
Influence of the Bio−Nano Interface on Enzyme
Catalytic Behavior. Previously characterized CMATs were
used as nanosupports for model enzyme soybean peroxidase
(SBP) immobilization (SBP, Figure 2a); the high dispersity of
the CMATs was required to ensure uniform loading of the
nanosupports. The different radii of curvature of the CMATs
were required to determine the geometrical congruence and thus
the degree of enzyme−nanosupport interactions.11,19,48
Three independent immobilization techniques, i.e., physical
adsorption, covalent binding, and covalent binding through a
PEG linker, were used. The different immobilization techniques
aimed to provide a variety of enzyme−nanosupport interactions.
In particular, physical binding provides a multipoint attachment;48,56 however, such a process was previously shown to lead
to deformation of the enzyme (e.g., change in active’s site
conformation or change in the enzyme’s footprint) at the
nanointerface.11 Covalent binding might reduce such deformation while theoretically serving as a zero-length single point
attachment technique.19,47,49,57 Lastly, covalent binding through
an arm spacer could serve as a single-point immobilization
method that brings the enzyme away from the nanosupport while
increasing its substrate binding capability.11,12 Figure 2b shows
the concept of SBP immobilization onto the COOH-functionalized CMATs.
The amounts of SBP attached to the different CMATs relative
to the amount of CMATs being used (i.e., the enzyme loadings)
are shown in Supporting Information Table S2, while the specific
retained activities of the enzyme-based conjugates relative to the
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maximum rate of reaction), Km (i.e., Michaelis−Menten
constant), and kcat (i.e., enzyme turnover) were calculated
using nonlinear regression36,59 and compared with the kinetic
parameters of the free enzyme in solution (Table 1). The Vmax of
SBP physically immobilized onto MWCNTs decreased by 87%,
while the Vmax of the SBP immobilized onto GON decreased by
about 98% relative to Vmax of free enzyme in solution. Km values
of the immobilized SBP also showed an overall decrease however
within the same order of magnitude with the Km values of the free
enzyme. The catalytic efficiency (kcat/Km) of the immobilized
enzyme (generally used as a comparator for the rate at which the
immobilized enzyme catalytically transforms its substrate)36,60
was much lower than that of the free SBP and varied both with
the nanosupport and immobilization technique being used. For
example, the catalytic efficiencies of SBP covalently bound onto
SWCNTs, MWCNTs and GON were about 10%, 11%, and 3%
of that of the free enzyme in solution. Further, the lowest activity
of immobilized SBP was obtained at the flat surface of GON.
The observed changes in the kinetic parameters indicate that
the different characteristics of the nanosupports influenced
directly the catalytic behavior of the immobilized enzyme. Even
though no significant changes in the enzyme active site
conformation occurred (i.e., the Km of the immobilized enzymes
were in the same order of magnitude with the ones of the
corresponding free enzymes), the multipoint attachment
resulted from the enzyme physical binding could explain both
the decrease in the rate of reaction and the reduced catalytic
efficiency. In particular, the multipoint attachment led to
decreased substrate-binding ability for the immobilized enzyme
relative to free enzyme in solution.11,47−49,61
These results are in agreement with previous reports that
showed that enzymes immobilized onto nanosupports with
smaller diameters and thus higher radii of curvature (i.e.
SWCNTs (0.8−1.2 nm) or MWCNTs (10−20 nm) relative to
GON (500−5000 nm)) tend to retain higher levels of
activity.11,62,63 Higher radius of curvature of the nanosupports
ensures an increased center-to-center distance between two
adjacent immobilized enzymes (Figure 2b), which could
potentially reduce the unwanted interactions between neighboring proteins and also reduce their multi-attachment points to the
nanosupports. Contrary to that, increased protein−protein
interactions caused by a less curved surface could result in a
more dramatic activity loss over time and in a harsh
environment.11,12,36,63
The nanosupport’s curvature trend was not confirmed for the
enzyme immobilized onto SWCNTs relative to the enzyme
immobilized onto MWCNTs; in particular, SBP showed the
highest enzyme activity at the MWCNTs interface which has a
larger radius of curvature than that of the SWCNTs. The
apparent discrepancy in the reported results is due to the bio−
nano interface being also influenced by the enzyme structure and
its surface energy.24 Specifically, at the working pH (PBS pH 7),
SBP carries a negative charge (pI 3.9).64 The presence of a larger
density of COOH groups onto the MWCNTs surface effectively
lowers their pI more so than that of the SWCNTs.65 Thus, the
SWCNTs will carry a weaker negative charge compared to that of
the MWCNTs leading to less repulsion of the enzyme at their
nanointerface. This effect coupled with the relatively large
dimensions of the SBP (6.1 nm × 3.5 nm × 4.0 nm)66 when
compared to the diameter of the SWCNTs (0.8−1.2 nm) could
also lead to an increase in protein−protein interactions and thus
account for the lower activity and reduced catalytic efficiency as
observed at this nanointerface.
Figure 3. Catalytic behavior of model enzyme SBP immobilized onto
different CMATs. (a) Comparison of the specific retained activity of
SBP upon immobilization onto SWCNTs, MWCNTs, and GON.
Physical adsorption, covalent binding, and covalent binding with a PEG
linker were used. The nanosupport diameter increases from left to right.
Michaelis−Menten kinetics data of SBP immobilized using physical
adsorption (filled square), covalent binding (filled circle), and covalent
binding via PEG linker (filled triangle) onto (b) SWCNTs, (c)
MWCNTs, and (d) GON. Enzyme retained specific activity and kinetics
depend on the nanosupport characteristics, both physical and chemical.
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Table 1. Soybean Peroxidase (SBP) Michaelis−Menten Kinetics
nanosupport and immobilization method
Vmax (μM/μg s)
Km (μM)
kcat (1/s)
kcat/Km
SWCNTs (physical)
SWCNTs (covalent)
SWCNTs (covalent with PEG)
MWCNTs (physical)
MWCNTs (covalent)
MWCNTs (covalent with PEG)
GON (physical)
GON (covalent)
GON (covalent with PEG)
free SBP
0.005 ± 0.001
0.012 ± 0.003
0.022 ± 0.011
0.017 ± 0.007
0.011 ± 0.004
0.008 ± 0.003
0.003 ± 0.001
0.005 ± 0.001
0.002 ± 0.001
0.128 ± 0.042
3.7 ± 1.0
1.9 ±0.7
1.9 ± 0.8
7.2 ± 2.3
1.6 ± 0.4
2.9 ± 0.4
1.4 ± 1.0
2.6 ± 0.3
2.8 ± 2.7
1.9 ± 0.8
0.14 ± 0.04
0.33 ± 0.12
0.61 ± 0.43
0.47 ± 0.27
0.30 ± 0.16
0.22 ± 0.12
0.08 ± 0.04
0.14 ± 0.04
0.03 ± 0.04
3.53 ± 1.64
0.04 ± 0.02
0.20 ± 0.07
0.33 ± 0.09
0.07 ± 0.02
0.22 ± 0.13
0.08 ± 0.03
0.12 ± 0.13
0.06 ± 0.01
0.02 ± 0.02
2.01 ± 0.63
Specificity of the Bio−Nano-Interface Reaction. To
assess whether there is a symbiotic relationship between the
immobilized enzyme and nanomaterial characteristics that
influence such catalytic behavior at nanointerfaces and whether
there is an optimal nanosupport that can be used when aiming to
preserve enzyme catalytic behavior, we extended our initial study
of the SBP to two additional biocatalysts, namely chloroperoxidase (CPO) and glucose oxidase (GOX). The additional studies
however excluded GON as a nanosupport because of the low
activity and increased protein−protein interactions observed
when SBP was used as an example.
Our complementary studies confirmed that MWCNTs
nanosupports provided once again the optimum nanointerfaces
to preserve the additionally selected two-enzyme catalytic
behavior and activities. In particular, CPO bound onto
MWCNTs retained about 29%, 49%, and 30% specific activities
after physical adsorption, covalent binding, and covalent binding
through the PEG linker, respectively, when compared to free
enzyme in solution (Supporting Information Table S3). These
specific activities were ∼27%, 46%, and 27% higher for each
respective immobilization method when compared to the
specific activity of the enzyme immobilized onto SWCNTs.
Further, covalent binding onto MWCNTs seemed to have
benefited CPO more than it benefited SBP. This was presumably
due to the higher ability of CPO to bind away from its active site
when compared to SBP. Specifically, even though both enzymes
have similar sizes and molecular weights,24,25,67 the different
mapping of their amino acid sequences as well as their different
number of lysine groups (five lysine for CPO and only three for
SBP) influenced their different binding ability at nanointerfaces.
Furthermore, at the working pH values, i.e., CAB (pH 4.8) for
CPO (pI 4.0) and PBS (pH 7) for SBP (pH 3.9), each enzyme is
negatively charged, but SBP is more so.64,68 Lastly, the decrease
in activity observed upon utilization of the PEG linker was
attributed to the nonspecific interactions of the PEG linker with
the active site of both CPO and SBP containing histidine groups.
Specifically, studies have shown that histidine group interactions
with PEG could potentially lead to substrate inhibition and
decreased enzyme activity.69,70 The greater impact seen for CPO
is presumably due to the inherent rigidity of its active site when
compared to the more flexible one of SBP.69,71
The kinetic behavior of the immobilized CPO (Supporting
Information Table S4) was also assessed using varying
concentrations of hydrogen peroxide (Figure 4a−c). The Vmax
of CPO covalently immobilized onto MWCNTs was larger than
that of the enzyme immobilized through both physical and
covalent through the PEG linker techniques. Km values for the
immobilized CPO were on the same order of magnitude as for
the free enzyme, indicating that no significant enzyme active site
conformational change occurred upon immobilization. Further,
kcat/Km catalytic efficiency of the CPO physically immobilized
onto SWCNTs and MWCNTs decreased to about 99% and 78%
relative to the catalytic efficiency of the free enzyme.
The more complex and larger GOX (6.0 nm × 5.2 nm × 7.7
nm)72 showed however an increase in the retained specific
activity upon immobilization using covalent binding, with a
further increase upon the utilization of the PEG linker
(Supporting Information Table S5). Namely, GOX bound to
MWCNTs physically, covalently, and covalently with the PEG
linker resulted in retained specific activities of around 20%, 44%,
and 63%, respectively, relative to the activity of the free enzyme
in solution. The active site inhibition was less likely to occur in
the GOX trials due to its extended numbers of lysine residues (i.e.,
60 lysine groups present on the enzyme structure compared to
only five or three for CPO and SBP, respectively)73 that would
thus offer multiple binding sites for the specific covalent
immobilization. Further, the benefit of the PEG linker was
obvious for this large enzyme, presumably due to the reduced
interactions of the GOX or reduced enzyme−enzyme interactions at the nanosupports.11,74,75
The catalytic behavior of the immobilized GOX (Supporting
Information Table S6) was also evaluated using varying
concentrations of glucose (Figure 4d−f). Specifically, GOX
bound to MWCNTs physically, covalently, and covalently
through a PEG linker yielded Vmax values of around 0.098,
0.218, and 0.234, respectively. These trends correspond to those
resulting from specific retained activity determination. Km values
for GOX were on the same order of magnitude as for the free
enzyme, confirming that there was no significant enzyme active
site conformational change upon immobilization.
Optimum Nanosupport for Optimum Catalytic Behavior. Our studies showed that the impact on enzyme binding at
nanosupport interfaces is a function of both the enzyme and the
nanosupport characteristics. Thus, in order to ensure maximum
catalytic efficiency of bio−nano conjugates for selected
applications, there is an optimum nanosupport and an optimum
immobilization method to be used. For instance, our results have
shown that nanosupports of MWCNTs 10−20 nm in diameter
functionalized with COOH groups are the most suitable for
being used for immobilization of enzymes with a footprint of half
of this diameter or as large as the nanosupport itself (Table 2).
Further, our results have shown that the catalytic behavior of the
enzymes upon immobilization is a function of the overall enzyme
isoelectric properties and changes in the surrounding environment. While our studies have used three selected enzymes and
three selected nanosupports, they can further be extended to
identify the best parameters and thus conditions to be considered
for synthetic applications of such biocatalyst-based conjugates.
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Figure 4. Catalytic behavior of chloroperoxidase (CPO) and glucose oxidase (GOX) immobilized onto different nanosupports. (a) Specific retained
activity comparison of the CPO immobilized onto SWCNTs and MWCNTs via physical adsorption, covalent binding, and covalent binding via PEG
linker. Michaelis−Menten kinetics of CPO immobilized using physical adsorption (filled square), covalent binding (filled circle), and covalent binding
via PEG linker (filled triangle) onto (b) SWCNTs and (c) MWCNTs. (d) Specific retained activity comparison of GOX immobilized onto SWCNTs
and MWCNTs via physical adsorption, covalent binding, and covalent binding via PEG linker. Michaelis−Menten kinetics data of GOX immobilized
using physical adsorption (filled square), covalent binding (filled circle), and covalent binding via PEG linker (filled triangle) onto (e) SWCNTs and (f)
MWCNTs.
Table 2. MWCNT-Based Conjugates as Optimum Nanosupports to Provide High Catalytic Behavior
enzyme (immobilization method)
Vmax (μM/μg s)
Km (μM)
kcat (1/s)
kcat/Km
SBP (covalent)
CPO (covalent)
GOX (covalent with PEG)
0.011 ± 0.004
12.42 ± 2.43
0.234 ± 0.032
1.6 ± 0.4
120 ± 8
2600 ± 700
0.30 ± 0.16
521.77 ± 102.14
42.12 ± 8.15
0.22 ± 0.13
4.48 ± 0.86
0.018 ± 0.008
well as the symbiotic reactions that take place at this interface,
can be tailored to lead to maximum retained enzyme activity
while augmenting recovery of active enzyme−nanomaterial
conjugates. Providing user-directed feedback for individual
For instance, one can envision comparing even lower surface
curvatures (i.e., spheres or gold nanorods) in order to understand
how nanomaterial characteristics and physico-chemical properties and the interplay at the enzyme−nanosupport interface, as
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application, and accounting for biochemical data relaying on the
characteristics of both the nanosupport and the biocatalyst being
tested, is empirically necessary for enzymes immobilized onto
carbon-based nanosupports to reach their full operational
potential.
■
CONCLUSIONS
We showed that controlling the interplay as well as the symbiotic
reactions that take place at the enzyme−carbon-based nanointerfaces lead to enzyme-based conjugates with higher catalytic
behavior. In particular, we showed that activity of the enzyme−
carbon-based conjugates can be tuned by the user by controlling
the immobilization conditions, the local curvature of the
nanosupport, and its physico-chemical properties. Further, our
studies showed that user manipulation of the immobilization
conditions as well as careful nanosupport and enzyme selection
are required for the optimum catalytic efficiency of these
conjugates. The detailed characterization and optimization of the
enzyme−nanointerface reactions will potentially result in
improved interfacial interactions, stable catalytic behaviors, and
thus a greater understanding of the molecular requirements and
symbiotic reactions at such interfaces for integrated technological applications of bio−nano conjugates in pharmacological
industry, biosensors, biofuel cells and bioactive coatings
formation.
■
ASSOCIATED CONTENT
S Supporting Information
*
In-depth characterization of the carbon-based nanomaterials
used in this study. Briefly, Figure S1 contains the SEM images of
these carbon-based nanomaterials, while Table S1 contains the
nanomaterial dispersity analysis. Tables S2−S6 contain
information on the loading, activity data, and Michaelis−Menten
kinetics for each individual enzyme−nanosupport configuration.
This material is available free of charge via the Internet at http://
pubs.acs.org.
■
AUTHOR INFORMATION
Corresponding Author
*Mailing address: Department of Chemical Engineering, West
Virginia University, Benjamin M. Statler College of Engineering
and Mineral Resources, P.O. Box 6102, Morgantown, WV
26506, United States. E-mail: [email protected].
Tel.: 1 304 293 9338. Fax: 1 304 293 4139.
Notes
The authors declare no competing financial interest.
■
ACKNOWLEDGMENTS
This work was supported by the National Science Foundation
(NSF-CBET: 1033266). The authors acknowledge NanoSAFE
and WVU Chemical Engineering for the shared facilities. The
authors thank undergraduate Andrew Maloney for his
involvement in the initial stage of the project.
■
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Applied Surface Science 264 (2013) 261–268
Contents lists available at SciVerse ScienceDirect
Applied Surface Science
journal homepage: www.elsevier.com/locate/apsusc
Effects of acid treatment on structure, properties and biocompatibility of carbon
nanotubes
Chenbo Dong a , Alan S. Campell a , Reem Eldawud a , Gabriela Perhinschi a ,
Yon Rojanasakul b , Cerasela Zoica Dinu a,∗
a
b
Department of Chemical Engineering, West Virginia University, Morgantown, WV 26506, USA
Department of Basic Pharmaceutical Sciences, West Virginia University, Morgantown, WV 26506, USA
a r t i c l e
i n f o
Article history:
Received 28 July 2012
Received in revised form
25 September 2012
Accepted 28 September 2012
Available online 23 October 2012
Keywords:
Nanotubes
Acid treatment
Structure modification
Cytotoxicity
Biocompatibility
a b s t r a c t
Carbon nanotubes (CNTs) are promising to be the next generation of viable tools for bioapplications.
Further advances in such bioapplications may depend on improved understanding of CNTs physical and
chemical properties as well as control over their biocompatibility. Herein we performed a systematic
study to show how acid oxidation treatment changes CNTs physical and chemical properties and leads
to improved CNTs biocompatibility. Specifically, by incubating CNTs in a strong acid mixture we created
a user-defined library of CNTs samples with different characteristics as recorded using Raman energy
dispersive X-ray spectroscopy, atomic force microscopy, or solubility tests. Systematically characterized
CNTs were subsequently tested for their biocompatibility in relation to human epithelial cells or enzymes.
Such selected examples are building pertinent relationships between CNTs biocompatibility and their
intrinsic properties by showing that acid oxidation treatment lowers CNTs toxicity providing feasible
platforms to be used for biomedical applications or the next generation of biosensors.
© 2012 Elsevier B.V. All rights reserved.
1. Introduction
Carbon nanotubes (CNTs) are nanoscale diameter materials
of tubular shape and micrometer length with many interesting
properties that make them viable candidates for a wide range of
applications including electrical circuits [1], hydrogen storage [2],
fiber optics [3], and conductive plastics [4]. In recent years, CNTs
functionalization with biomolecules such as proteins [5], enzymes
[6,7] or nucleic acids [8] opened up exciting bioapplications in
biolabeling [9], biosensing [10], drug delivery [11], bioseparation
[12] and tissue engineering [13]. However, further development
of such bioapplications is hindered by: (1) CNT’s limited available surface area for biomolecule functionalization [14], (2) lack
of understanding of CNTs growth mechanisms in uncontaminated
forms [15], (3) CNTs structural instability since larger nanotubes
are prone to kinking and collapsing [16,17], and (4) CNTs cytotoxicity and associated health risks posed during their manufacturing
and processing [18]. These challenges are mainly associated with
the fact that as-produced CNTs form large aggregates in liquid
∗ Corresponding author at: Department of Chemical Engineering, West Virginia
University, Benjamin M. Statler College of Engineering and Mineral Resources, PO
Box 6102, Morgantown, WV 26506, USA. Tel.: +1 304 293 9338;
fax: +1 304 293 4139.
E-mail address: [email protected] (C.Z. Dinu).
0169-4332/$ – see front matter © 2012 Elsevier B.V. All rights reserved.
http://dx.doi.org/10.1016/j.apsusc.2012.09.180
enviroments since their hydrophobic walls are prone to van der
Waals interactions [19]. Thus, in order to increase CNTs bioapplications [20] and reduce their aggregation [21] and cytotoxicity [22], it
is critical to overcome their intrinsic hydrophobicity and tendency
to form conglomerates in solution.
Numerous attempts have been made to overcome CNTs
hydrophobicity and increase their hydrophilicity; these include
gas- [23] and liquid-phase activation [24], and oxidation with
strong oxidants including hydrogen peroxide [25], potassium permanganate [26], potassium hydroxide [27], and nitric and/or
sulfuric acid [6,7,28]. Among these attempts, nitric and sulfuric
acid oxidation is regarded as the most prevalent treatment since
it is easy to implement in both laboratory and industrial settings [20]. When CNTs are oxidized with such aggressive acids,
their hydrophilicity is increased by the introduction of oxygencontaining functional groups, i.e., carboxyl [29], carbonyl [26,29],
and phenol groups [30]. Moreover, during such oxidation treatments amorphous carbon [31] and residual metal catalyst particles
are removed, possibly resulting in reduced intrinsic toxicity of CNTs
[22]. Despite the fact that wide evaluations of the effects of acid oxidation on CNTs have been carried out, systematic investigations of
changes in physical and chemical properties and how such changes
can be further employed for increasing CNTs biocompatibility and
thus bioapplications are still lacking.
Herein we performed a systematic study of the changes in physical and chemical properties of pristine CNTs upon user-controlled
262
C. Dong et al. / Applied Surface Science 264 (2013) 261–268
treatment with nitric and sulfuric acids. Further, we assessed how
these changes affect CNTs biocompatibility in relation to cellular
and enzymatic systems [6,7,10]. Our hypothesis was that selected
biological examples will help build pertinent relationships between
CNTs biocompatibility and their intrinsic properties and demonstrate how interface reactions between a biological molecule and
the nanomaterial can be further used to provide systems with lower
toxicity to be used for selected bioapplications as well as feasible
platforms for the next generation of biosensors.
2.5. CNTs solubility measurement
2. Materials and methods
The solubility of CNTs (pristine and acids oxidized) was evaluated in di water (pH 6.25) and Phosphate Saline Buffer (PBS, pH 7,
100 mM ionic strength). Briefly, CNTs were diluted in the solvent
of interest to yield to a 3 mg/ml solution. The suspension was then
centrifuged at 3000 rpm for 5 min; subsequently, part of the supernatant (0.8 ml) was removed and filtered through a 0.2 ␮m GTTP
filter membrane. The filter membrane was then dried under vacuum and the amount of CNTs was weighted. The solubility of the
CNTs was calculated based on the volume used for suspension and
the initial starting amount.
2.1. Acid oxidation of CNTs
2.6. CNTs length measurement
Acid oxidation treatment of single- and multi-walled carbon
nanotubes (SW- and MWCNTs, respectively) was employed to generate a library of samples with different physical and chemical
properties. Specifically, commercial SWCNTs (85% purity, Unidym
Inc.) and MWCNTs (95% purity, Nanolab Inc. (PD15L5-20)) were
incubated in a concentrated sulfuric (96.4%, Fisher, USA) and nitric
acid (69.5%, Fisher, USA) mixture in a ratio of 3:1 (V/V). The
CNTs/acids mixture (where CNTs can refer to either SW- or MWCNTs) was subsequently sonicated in an ice bath (Branson 2510,
Fisher, USA) for 1, 3, or 6 h, at a constant temperature of 23 ◦ C.
When the required time elapsed, CNTs/acids mixture was diluted
with deionized (di) water and filtered through a GTTP 0.2 ␮m
polycarbonate filter membrane (Fisher, USA). Several cycles of
resuspension in di water were employed to remove acidic residues
or catalysts. The CNTs were isolated on the filter, subsequently dried
in a vacuum desiccator and stored at room temperature for further
use.
An atomic force microscope (AFM, Asylum Research, USA) was
used to evaluate the length of pristine and acids treated CNTs. A
Si tip (Asylum Research, 50–90 kHz AC240TS, USA) helped perform
tapping mode in air. CNTs samples (i.e., pristine, 1, 3 or 6 h acids
oxidized SW and MWCNTs) were dispersed in di water (to yield
solutions of 0.1 mg/ml concentration), deposited on mica surfaces
(9.5 mm diameter, 0.15–0.21 mm thickness, Electron Microscopy
Sciences, USA) and allowed to dry over night under vacuum. Scan
images of 10, 5 or 1 (␮m × ␮m) areas were acquired. For each sample, at least 30 individual CNTs were counted and measured to
obtain average length distribution.
2.2. Energy dispersive X-ray analysis (EDX) of CNTs
Energy dispersive X-ray analysis (EDX) was used for quantitative elemental analysis of pristine and acid oxidized CNTs. Samples
(1 mg/ml in di water) were deposited on silica wafers and dried
under vacuum. The experiments were performed on a Hitachi S4700 Field Emission Scanning Electron Microscope (USA) with a
S-4700 detector combining secondary (SE) and backscattered (BSE)
electron detection (all in a single unit), operating at 20 kV. Results
are presented as a percent of elements relative to the most dominant element.
2.3. Scanning Electron Microscopy (SEM) of CNTs
Samples (1 mg/ml in di water of both pristine and acid treated
CNTs) were dried on silica wafers under vacuum and imaged using a
Hitachi S-4700 Field Emission Scanning Electron Microscope (USA)
with a field emission at 10 kV.
2.7. Cell culture and treatment with CNTs
Non-tumorigenic human bronchial epithelial cells (BEAS-2B)
were purchased from American Type Culture Collection (ATCC,
USA). The cells were cultured in DMEM medium supplemented
with 5% fetal bovine serum (FBS), 2 mM l-glutamine and 100units/ml penicillin/streptomycin (all reagents were purchased
from Invitrogen, USA). Cells were passaged weekly using 0.05%
trypsin (Invitrogen, USA) and kept in 5% CO2 at 37 ◦ C.
Pristine and acids oxidized SWCNTs were dispersed in di water
by sonication, filtered through the 0.2 ␮m GTTP filter membrane,
resuspended in cellular media and sonicated at room temperature to form stable dispersions. For treatment, BEAS-2B cells were
seeded overnight in a 12 well plates (Fisher, USA) at a density of
3.5E5 cells/well, and allowed to reach confluence. Subsequently,
the cells were exposed to 100 ␮g/ml SWCNTs; 24 h post exposure,
the cells were incubated with 6.5 ␮g/ml Hoechst 33342 dye (Molecular Probes, USA) for 30 min at 37 ◦ C and analyzed for apoptosis
by scoring the percentage of cells with intensely condensed chromatin and/or fragmented nuclei using fluorescence microscopy
(Leica Microsystems, USA). Approximately 1000 cell nuclei from ten
random fields were analyzed for each sample. The apoptotic index
was calculated as the percentage of cells with apoptotic nuclei relative to the total number of cells. At least 3 independent trials were
performed for each sample.
2.8. Functionalization of CNTs with enzyme
2.4. Raman spectroscopy of CNTs
Raman spectroscopy (performed on a Renishaw InVia Raman
Spectrometer, CL532-100, 100 mW, USA) allowed determination
of the chemical structure and any modifications resulted from the
acids oxidation of both pristine and acids treated CNTs. Briefly, CNTs
deposited on glass slides (Fisher, USA) were excited through a 20×
microscope objective using an Argon ion (Ar+ ) laser beam with a
spot size of <0.01 mm2 operating at 514.5 nm. Detailed scans were
taken in the 100–3200 cm−1 range; low laser energy (i.e., <0.5 mV)
and exposure time of 10 s were used to prevent unexpected heating
effects.
Soybean peroxidase (SBP, Bioresearch, USA) was covalently
attached to 1, 3 or 6 h acid treated MWCNTs using 1-ethyl-3[3-dimethylaminopropyl] carbodiimide hydrochloride (EDC; Acros
Organics, USA) and N-hydroxysuccinimide (NHS, Pierce, USA)
[32]. Briefly, 2 mg CNTs (MWCNTs) were dispersed in 160 mM
EDC and 80 mM NHS (total volume of 2 ml in MES (2-(Nmorpholino)ethanesulfonic acid sodium salt, 50 mM, pH 4.7, Sigma,
USA) for 15 min at room temperature with shaking at 200 rpm. The
activated MWCNTs were next filtered through the 0.2 ␮m GTTP
filter membrane, washed thoroughly with MES buffer to remove
any ester residues, immediately dispersed in 2 ml of 1 mg/ml SBP
C. Dong et al. / Applied Surface Science 264 (2013) 261–268
solution in PBS (100 mM, pH 7.0) and incubated for 3 h at room temperature at room temperature with shaking at 200 rpm. The resulting SBP–MWCNT conjugates were filtered and washed extensively
with PBS to remove any unbound enzyme [32]. The supernatants
and washes were collected to quantify enzyme loading.
2.9. Enzyme loading
The amount of SBP attached to MWCNTs (i.e., SBP loading) was
determined using standard BCA assay kit (Pierce, USA) and subtracting the amount of enzyme washed out in the supernatant and
washes from the amount of SBP initially added to the MWCNTs.
Briefly, the working reagent (1000 ␮l) was prepared by mixing 50
parts of reagent A with 1 part of reagent B (the reagents are provided
with the kit). The mixture of reagents A and B was further added
to 50 ␮l solutions of SBP-containing samples (i.e., the samples isolated in the form of the supernatant and washes). The resulting
solutions were incubated at 37 ◦ C for 30 min. Absorbance at 562 nm
was determined on a spectrophotometer (Fisher, USA). Control calibration curves were prepared by serial dilutions of SBP (free in
solution) into the working reagent.
2.10. Enzyme activity assay
The activity of SBP was measured by monitoring the oxidation reaction of (2,2 -Azinobis [3-ethylbenzothiazoline-6-sulfonic
acid]) (ABTS, Sigma, USA) in the presence of hydrogen peroxide
(H2 O2 , Sigma, USA). 20 ␮l of the SBP–MWCNTs conjugates were
added to 0.65 ml ABTS solution (0.5 mM final concentration, Pierce,
USA) and mixed; subsequently, 20 ␮l H2 O2 solution (0.2 mM final
concentration) was added to the sample in order to initiate the
reaction. The change in absorbance was monitored spectrophotometrically at 412 nm immediately upon addition of H2 O2 . The
initial reaction rate was calculated from the slope of the linear timecourse. The extinction coefficient of the oxidized ABTS product
is 32,400 M−1 cm−1 at 412 nm [33]. The activity of the immobilized enzyme is reported as specific activity relative to free enzyme
activity. The activity of the free enzyme was determined using an
equivalent amount of free enzyme (based on loading data) and the
protocol provided above.
2.11. Statistical analysis
All results are presented as mean ± standard deviation.
3. Results and discussion
We prepared a library of single- and multi-walled carbon nanotubes (SW- and MWCNTs) using liquid phase oxidation with a
strong nitric and sulfuric acids mixture [6,7]. The approach is shown
in Scheme 1; sonication in the acids mixture attacks the graphene
sheets on the C C bands [34], introduces defects and oxidizes the
CNTs at the defect sites leading to shorter nanotubes. To reduce
the reaction rate of acids attack, the water bath sonicator was
maintained at room temperature. The carboxylic acidic groups
263
introduced in SW- and MWCNTs were determined previously using
acid–base titrations [35,36] or the formation of a dodecylamine
zwitterions [37].
We further investigated the chemical composition of pristine
and acids oxidized CNTs using energy dispersive X-ray analysis
(EDX) [20,38]. EDX spectra of pristine SW- and MWCNTs are shown
in Fig. 1a and b, respectively, as a plot of X-ray counts vs. energy (in
keV). The analysis revealed the presence of high contents of carbon
(C) and oxygen (O), with iron (Fe) as metal catalyst in both pristine SW- and MWCNTs samples. The energy peaks correspond to
the various elements in the sample, with Fe yielding two peaks at
0.70 keV and 6.40 keV [39]. Other elements (e.g., Al, Si, Cl, S, etc.)
were also present but in very low amount. The Fe peak was larger
for the SWCNTs sample when compared to the MWCNTs one. The
difference was reflective of their pristine characteristics since SWCNTs purity was 85% while the purity of pristine MWCNTs was 95%,
per manufacturer information (see Section 2). The insets in Fig. 1
show the changes in the O and Fe contents with the acids oxidation treatment time for both SW- and MWCNTs samples. As shown,
Fe content decreased with the treatment time for both SW- and
MWCNTs samples indicating removal of the metal catalyst. The
decrease in the Fe content was more pronounced for the SWCNTs
when compared to MWCNTs samples. This is a reflection of the
different purities of the two samples chosen in these experiments.
For the O content, the change was also dependent on the sample
characteristics. The relative low purity SWCNTs samples contain
more amorphous carbon [40] than the higher purity MWCNTs [41].
Thus, the acids treatment led to a significant increase of the O content with the acids treatment time for the SWCNTs (Fig. 1a, inset)
when compared to a smaller increase for the MWCNTs samples.
Fig. 2 shows the SEM images of the pristine and acids treated
samples (both SW- and MWCNTs). As shown, user-controlled acids
treatment did not lead to significant morphological changes either
for SW- (Fig. 2a shows pristine SWCNTs while Fig. 2c shows 6 h
acids treated SWCNTs) or MWCNTs (Fig. 2c shows pristine MWCNTs
while Fig. 2d shows 6 h treated MWCNTs) samples.
The structural changes upon acids treatment of the CNTs
samples were investigated using Raman resonance spectroscopy
[42–44]. Fig. 3 shows the Raman spectra of pristine and acids
treated SW- and MWCNTs. The Raman analysis of the SWCNTs
reveals the presence of 4 bands (Fig. 3a), the so-called D (disorder mode) band around 1340 cm−1 , G− and G+ bands at around
1545 cm−1 and 1590 cm−1 respectively, and G band at 2650 cm−1
[22,45]. The Raman analysis of the MWCNTs also reveals the presence of 4 bands (Fig. 3b), with the D band around 1340 cm−1 , G
band at 1585 cm−1 , G band at 2650 cm−1 , and another band at
2920 cm−1 [46,47]. The D band around 1340 cm−1 is related to the
non-crystalline C species, i.e., defects in the CNTs [48], while the
G band observed around 1585 cm−1 is indicative of a high degree
of ordering and well-structured C-based structures [42]. The size
of the D band relative to the G band can be used as a qualitative
measurement for the formation of undesired forms of C [49]. Both
pristine and acids treated CNTs (SW- and MWCNTs) have a relatively small D band at around 1350 cm−1 , with the D band being
wider and shifted toward higher frequency in the acids treated
Scheme 1. Time-dependent incubation of pristine CNTs (SW- and MWCNTs) with a mixture of sulfuric and nitric acids leads to acids oxidized CNTs.
264
C. Dong et al. / Applied Surface Science 264 (2013) 261–268
Fig. 1. EDX elemental analysis of pristine SWCNTs (a) and MWCNTs (b). The insets show the changes in the O and Fe contents with the acids treatment time employed under
user-control.
samples when compared with the pristine ones. The ratio of intensity of D peak relative to the G peak represents the degree of CNTs
functionalization [49]. Higher ID /IG ratio suggests higher level of
functionalization (I represents the peak’s relative intensity).
D band, G band and ID /IG ratio of the various CNTs samples
(both SW- and MWCNTs) are shown in Table 1. The ratio of ID /IG
for SWCNTs changed minimally from 0.237 for pristine to 0.263
after 6 h acid treatment. For 1 and 3 h acid oxidized SWCNTs, the
ID /IG ratio seemed to have decreased. Previous reports have shown
that for relatively low purity CNTs (in this particular example the
SWCNT’s purity is 85%; see Section 2) the ID /IG does not provide
precise overall information on the sample structure [50], and the
ID /IG ratio might be both a reflection of washing away amorphous
carbon while simultaneously inducing carboxylic acid groups [20].
Table 1
Relative intensity of representative Raman peaks of pristine and acids treated CNTs.
CNT
D band position
(cm−1 )
G band position
(cm−1 )
ID/IG intensity
ratio
Pristine SWCNTs
1 h cut SWCNTs
3 h cut SWCNTs
6 h cut SWCNTs
1328
1333
1336
1336
1590
1587
1592
1595
0.237
0.195
0.229
0.263
Pristine MWCNTs
1 h cut MWCNTs
3 h cut MWCNTs
6 h cut MWCNTs
1345
1347
1349
1351
1586
1586
1586
1589
0.457
0.783
0.788
0.796
For instance, in the initial 1 h SWCNTs acids oxidation, the effect of
washing away amorphous C (which is known to lead to decreased
ID /IG [51]) suppressed the effect of adding carboxylic acid groups
(which is known to lead to increased ID /IG [52]). However, after
6 h, most of the amorphous C was removed and the ID /IG became
indicative only of the degree of functionalization with carboxylic
groups.
ID /IG for MWCNTs increased from 0.457 for pristine to 0.788 for
3 h, and 0.796 after 6 h acids oxidation. This increase in the level
of functionalization has a similar trend to the increase in the O or
decrease in the Fe catalyst content as observed through the EDX
analyses (Fig. 1). Specifically, for the high purity MWCNTs most of
the Fe catalysts are removed during the 3 h treatment time (see
inset Fig. 1b) this leading to removal of the defects in the MWCNTs
structure. Since defects are where the promotion of the carboxylic
groups formation takes place [53], and since for the MWCNTs there
was a small decrease in the Fe and a small increase in the O content (Fig. 1b inset) from the 3 h to 6 h treatment time, the ID /IG for
MWCNTs will be minimally changed between these time points as
indicated in Table 1. Such analyses confirm that the acids oxidation
introduced CNTs chemical property changes i.e., added functional
free carboxylic acid groups, to both SW- and MWCNTs sample.
We further investigated how the degree of CNTs dispersion in
water-based environments is influenced by the acids oxidation
time. We used two solvents with different pH’s and ionic strengths,
i.e., di water (pH 6.25) and Phosphate Saline Buffer (PBS, pH 7,
100 mM). The results (Fig. 4) indicated that the solubility of CNTs
C. Dong et al. / Applied Surface Science 264 (2013) 261–268
265
Fig. 2. SEM image of (a) pristine SWCNTs, (b) pristine MWCNTs and (c) 6 h acids treated SWCNTs (d) 6 h acids treated MWCNTs; the scale bar is 1 ␮m.
in both di water and PBS was improved upon the acids oxidation,
with increased acids oxidation times leading to increased solubility. Generally, pristine and acid oxidized SWCNTs (either 1, 3 or
6 h cut) were more dispersed in PBS when compared to di water
(Fig. 4a). MWCNTs did not show a similar trend; specifically, pristine and 1 h cut MWCNTs were more soluble in PBS, however, after
longer acids oxidation times (i.e., 3 and 6 h) the solubility was
higher in water when compared to PBS (Fig. 4b). The changes in
the solubility observed for the MWCNTs samples after longer acids
oxidation times are correlated with the changes in the functionality
of these samples and number of carboxylic acidic groups being generated. Specifically, longer acids oxidation times will lead to higher
number of carboxyl groups being generated (see Figs. 1 and 3).
When the MWCNTs acids treated samples are placed in waterbased environments, carboxylate anions groups are generated by
the deprotonation of carboxylic acid groups [54]. At high ionic
strength, the probability for these anions to form aggregates [55]
increases thus leading to the lower solubility observed for the 3
and 6 h acids oxidized MWCNTs placed in PBS when compared to
solubility of these samples placed in water.
Atomic force microscopy (AFM) and tapping mode [56] was
used to analyze the morphology and quantify the length of the
CNTs samples. Specifically, cross sectional areas from (10 × 10) to
(1 × 1) ␮m × ␮m were scanned to derive the length of at least 30
CNTs/sample (both SW- and MWCNTs; pristine, 1, 3 and 6 h cut).
Pristine and acids oxidized CNTs length distributions are shown in
Fig. 3. Raman spectra of pristine, 1, 3 and 6 h acids oxidized SWCNTs (a) and MWCNTs (b).
266
C. Dong et al. / Applied Surface Science 264 (2013) 261–268
Fig. 4. Solubility of pristine and acids oxidized SWCNTs (a) and MWCNTs (b) in deionized (di) water and phosphate buffer saline (PBS).
Fig. 5. The average length distribution and the standard deviation of SWCNTs (a) and MWCNTs (b) with the acids treatment time.
Fig. 5; a general non-linear distribution toward shorter CNTs was
observed with the increase in the acids oxidation time.
Having established that the acids oxidation influences the
chemical and physical properties of pristine CNTs (both SW- and
MWCNTs), we proceeded to examine whether user-controlled
acids oxidation would also affect CNTs biocompatibility. First, we
performed a systematic study on the cellular toxicity resulted from
the incubation of immortalized human bronchial epithelial cells
with acids oxidized SWCNTs. Previous in vivo studies have shown
that cellular exposure to SWCNTs results in macrophages without
nuclei [57,58], with SWCNTs inducing chromosome aberration [18].
However, to our knowledge, no studies that looked at the influence of the different acids oxidation times to BEAS-2B immortalized
human bronchial epithelial cells have been performed. Moreover,
to our knowledge, there is no correlation in the literature on how
cellular toxicity depends on the SWCNTs physical and chemical
properties as impaired by the acids oxidation time and how such
toxicity can be controlled. In our experiments, BEAS-2B cells were
exposed to SWCNTs for 24–72 h at Permissible Exposure Limit
for particulates not otherwise regulated (i.e., 100 ␮g/ml of SWCNTs, based on previous laboratory exposure levels [58,59]). Fig. 6
shows the percentage of apoptotic BEAS-2B cells upon exposure
to SWCNTs; our data shows that the cytotoxicity of the 6 h acids
treated SWCNTs is lower than that of pristine SWCNTs. Specifically, the percentage of apoptotic cells for pristine SWCNTs is
about 19% while the percentage of apoptotic cells for 6 h acids
treated SWCNTs is about 15% upon 72 h incubation. These results
are comparable to control cells (cells that have not been exposed
to SWCNTs) and they emphasize that user-controlled acids oxidation time can be employed to create a library of sample of SWCNTs
that have high biocompatibility with cellular system. We hypothesized that the observed trend is due to the changes in the chemical
and physical structure of the SWCNTs upon acid functionalization.
Specifically, shorter and more hydrophilic SWCNTs (see our previous EDX and AFM results) would be predominantly taken up by the
cells through endocytosis [60], while for the longer SWCNTs the
uptake mechanism is predominantly through piercing [61]. Further, the longer SWCNTs once taken up by the cells can localize at
the cell nucleus and interfere with the normal progression of cells to
Fig. 6. Cytotoxicity of pristine and 6 h acids treated SWCNTs to BEAS-2B human
epithelial cells after 24, 48 and 72 h, respectively.
C. Dong et al. / Applied Surface Science 264 (2013) 261–268
Table 2
Loading and retained specific activity of immobilized SBP onto acids treated
MWCNTs.
Sample
Loading (mg
SBP/mg MWCNTs)
Retained specific
activity (%)
1 h cut covalent
3 h cut covalent
6 h cut covalent
0.254 ± 0.05
0.282 ± 0.06
0.265 ± 0.15
9.40 ± 1.68
28.18 ± 6.52
33.97 ± 9.82
267
Acknowledgements
This work is support by the NSF/CBET 1033266 and NSF/EPS1003907. The authors acknowledge Adrienne McGraw, Chemical
Engineering/WVU for her help with EDX/SEM analysis and Dr.
Weiqiang Ding/WVNano for his help with Raman analysis. Authors
acknowledge use of the WVU Shared Research Facilities.
References
division [58,60] thus leading to the observed results. In the future,
such library can be utilized for instance for the cellular delivery of
drugs or molecules of interest [11].
Secondly, we tested the biocompatibility of the CNTs in relation to enzyme immobilization. Enzyme immobilization provides
enzyme reutilization and eliminates costly enzyme recovery
and purification processes. CNTs have high surface area [62]
that facilitates the preparation of enzyme–CNTs conjugates with
high enzyme loadings per unit weight of material [63,64] and
promote protein activity and stability in strongly denaturing environments [63–67]. A test enzyme, namely soybean peroxidase
(SBP) was immobilized through covalent binding onto MWCNTs [64,65,68–73]. Table 2 shows the loading (defined as the
amount of the enzyme immobilized onto the MWCNTs) and the
retained specific activity of the enzyme after immobilization. Our
results show that the physical and chemical properties of the
CNTs influence the enzyme loading and retained specific activity. The lowest activity was observed for the SBP immobilized
onto the 1 h acids treated MWCNTs, while the activity of SBP
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solubility of these CNTs (see Fig. 4b). Specifically, lower solubility of the MWCNTs leads to larger conglomerate formation (due
to predominant van der Waals interactions between the MWCNTs hydrophobic walls) thus resulting in a lower surface area
exposed for immobilization of SBP. Further, SBP (a 40 kDa molecular weight enzyme) has an isoelectric point of 3.9 [74]; thus, at the
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hydrophobic 3 and 6 h acids oxidized samples. Stronger binding
of the SBP to the substrate will further lead to a reduction in the
protein activity [32,74]. Such example demonstrated the utility
of creating biocompatible MWCNTs nanosupports for biosensors
applications [10]; such enzyme-nanosupport-based application
can further be employed for decontamination of bacteria and spores
[32].
4. Conclusion
Our results have shown that user-controlled acid oxidation of
CNTs led to the formation of a library of samples with different
physical and chemical properties. Specifically, we have shown that
CNTs oxidation with a nitric and sulfuric acids mixture results in
removal of metal catalyst, an increase in the number of functional
groups having electron accepting ability, and generation of shorter
CNTs with higher solubility in aqueous environments. Our results
were confirmed by Raman spectroscopy, SEM, AFM, EDX and solubility tests. Further, we have shown that CNTs acids oxidation
improves nanotube biocompatibility as tested by direct incubation
with human epithelial cells or with test enzymes. User-controlled
design of CNTs biocompatibility can lead to new types of analytical
tools for life science and biotechnology [75–77].
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