Research Article
Received: 8 December 2015
Revised: 27 April 2016
Accepted article published: 30 May 2016
Published online in Wiley Online Library: 25 July 2016
(wileyonlinelibrary.com) DOI 10.1002/pi.5165
Investigation of the thin film crystallization of a
DNA copolymer hybrid composed of chitosan
Ilyès Safir,a Mohamed Chami,b Thomas Buergic and Corinne Nardina*
Abstract
We describe for the first time the crystallization in thin films of a DNA copolymer composed of a low molecular weight chitosan
backbone to which a sequence of nucleic acids is grafted (chitosan-g-ssDNA). As assessed by atomic force microscopy, optical
microscopy and spectroscopy, crystallization occurs due to intermolecular hydrogen bonding in which the nucleic acid strands
engage. The morphology of the crystals depends on the affinity for the surface of the polymer segments that compose the DNA
copolymer hybrid. The nucleic acids adsorb on mica and silica on which side-branched structures are observed whereas chitosan
interacts preferentially with gold, and dendritic crystals are assembled. Attenuated total reflectance infrared spectroscopy
supports the high ordering of the structure and the establishment of strong intermolecular interactions by hydrogen bonding.
© 2016 Society of Chemical Industry
Supporting information may be found in the online version of this article.
Keywords: DNA copolymer; chitosan; self-assembly; thin film; crystallization
INTRODUCTION
Polym Int 2016; 65: 1165–1171
∗
Correspondence to: C Nardin, Université de Pau et des Pays de l’Adour (UPPA),
Institut des Sciences Analytiques et de Physico-Chimie pour l’Environnement
et les Matériaux (IPREM), Equipe Physique Chimie des Polymères (EPCP), UMR
5254, Hélioparc, 2 avenue du Président Pierre Angot, 64053 Pau Cédex 9, France.
E-mail: [email protected]
a University of Geneva, Faculty of Sciences, Department of Inorganic and Analytical Chemistry, 30 quai Ernest Ansermet, 1211 Geneva 4, Switzerland
b Centre for Cellular Imaging and NanoAnalytics (C-CINA), Biozentrum, University of Basel, Mattenstrasse 26, 4058 Basel, Switzerland
c University of Geneva, Faculty of Sciences, Department of Physical Chemistry, 30
quai Ernest Ansermet, 1211 Geneva 4, Switzerland
www.soci.org
© 2016 Society of Chemical Industry
1165
Semicrystalline block copolymers, which contain at least one
crystallizable block, might undergo crystallization in thin films
under suitable conditions. Both chemical interactions between the
copolymer and the surface and intermolecular forces direct the
thermodynamics of such self-assembling systems.1 Block copolymers, which are developed to propagate the properties of the
individual blocks to the macromolecule, self-assemble via a thermodynamically driven process in which the chemical dis-affinity
between the blocks driving them apart is counterbalanced by a
restorative force due to the chemical bond between the blocks.1
Thin film formation thus takes place to minimize surface energies
and maximize interactions with the surface. Preferential wetting
layers assemble by segregation of the preferred block to the substrate. Owing to a fine balance between microphase separation
and crystallization, thin film formation in a variety of ordered and
stable film patterns takes place, and dendrites, spherulites and
lamellae might be observed.1,2 The resulting crystals usually have
a thickness of tens of nanometres, which is comparable to the size
of the microdomains formed by microphase separation.
The effect of confinement on phase separation and
crystallization is a topic of several investigations usually
studied with blends.3 – 8 The origin of the confinement in
semicrystalline triblock copolymers of poly(vinylcyclohexane)block-poly(ethylene)-block-poly(vinylcyclohexane) (PVCH-b-PEb-PVCH) by blending with PE was investigated by a combination
of morphological investigations and simulations.8 The authors
showed that the chain length of the PE homopolymer significantly influences the crystallization behaviour, to such an extent
that phase separation might prevent crystallization when the
homopolymer is of short length. With a PE homopolymer of
larger length, morphological transitions could be induced. Chen
et al. reported on the investigation of this effect using blends of
poly(styrene)-block-poly(methyl methacrylate) (PS-b-PMMA) with
poly(ethylene oxide) (PEO).6 The PEO homopolymer is different
from the copolymer blocks yet miscible with PMMA. Low molecular weight PEO polymer dissolves in the PMMA microdomains and
morphological transitions are induced on increasing the PEO volume fraction, whereas high molecular weight PEO induces phase
separation. The mechanism of crystal thickening and kinetics
during annealing is usually studied by a combination of DSC and
temperature- and time-dependent X-ray scattering.9 Hot stage
AFM also enables the process to be followed in situ as evidenced
by Frank and coworkers.10 The formation of individual lamellae by
isothermal crystallization of PEO in thin and ultrathin films could
be shown with this technique.11 Interestingly, Carvalho et al.12
reported on the use of ellipsometry to evidence the temperature driven morphological transition from spheres to lamellae
with decreasing temperature in a diblock copolymer thin film of
poly(butadiene)-block-poly(ethylene oxide). Ellipsometry showed
that the transition is surface induced owing to the lower free
energy of the wetting layer at low temperature compared to that
of the spherical caps.
www.soci.org
1166
Besides the academic interest of establishing the mechanism of
crystallization in confined environments, the resulting thin films
are of potential use in several fields and in particular in lithography or optoelectronics.2 For instance, nanoporous thin films
of about 15–100 nm thickness can be used to control the epitaxial crystallization of Si and NiSi.13 The morphology of olefin
block copolymer structures driven by epitaxy induced crystallization was investigated by Wang and coworkers.14 In that study, the
templates were of periodicities on the scale of tens of nanometres, obtained in a bottom-up self-assembly approach, a process
directed by block copolymer self-assembly using inorganic precursors. Depending on the template thickness, either arrays of
isolated nanopillars or interconnected three-dimensional nanostructures were generated. A self-epitaxy effect in block copolymers was proposed by Han and coworkers owing to the covalent bonding between two blocks, poly(3-hexylthiophene) and
poly(p-phenylene).15
Biocompatible and biodegradable polymers are of interest
for several applications in the biological and medical fields.16
Along this line, the self-assembly of copolymers constituted
by a segment composed of a polypeptide, a polysaccharide
or a nucleotide sequence has been reported.17 – 21 Structure
formation in thin films of copolymers composed of biodegradable blocks has been reported. Ree and coworkers studied thin
films of a series of poly(n-hexyl isocyanate)-poly(𝜀-caprolactone)
(PHIC-PCLm, m = 1–3) by grazing incidence X-ray scattering.22
Poly(styrene)-poly(L-lactide) block copolymers (PS-PLLA) undergo
crystallization.23 Portinha and coworkers took advantage of stereocomplex formation between enantiomeric graft copolymers composed of a partially acetylated poly(vinyl alcohol) backbone and
either oligo(L-lactide) or oligo(D-lactide) as grafts.24 Microphase
separation of cellulose triacetate-block-poly(𝛾-L-glutamate)
(CTA-b-PBLG) was clearly shown by AFM studies.25 Chiang et al.26
reported on the melt and solvent induced crystallization of
PEO-b-PCL-b-PLLA triblock copolymer thin films. Interestingly,
although melt crystallized thin films induced a single-crystalline
morphology, solvent induced crystallization led to multiple crystalline layered crystals with flat-on chain orientation, which opens
discussion on layer-by-layer single-crystal formation by epitaxial
growth.
Chitosan results from the deacetylation of chitin. This complex
linear polymer exhibits amphiphilic properties that depend on various parameters.27 At low pH and when the degree of acetylation
is below 28%, chitosan is a strong polyelectrolyte.27 At full ionization, the persistence length is maximal and the polymer is a
semi-flexible chain.22 Upon increasing the neutralization degree,
chitosan becomes hydrophobic.27 Besides, chitosan is an elicitor
of numerous biological activities when in contact with animal or
plant cells.28 This property led to the synthesis of self-assembling
chitosan-based copolymers for various potential applications such
as wood preservation,28 drug delivery29 – 37 and gene delivery.38
To advance the understanding of the molecular organization of DNA copolymers and achieve highly versatile selfassembling macromolecules, we reported recently on the solution
self-assembly of a chitosan-based DNA copolymer.39 We demonstrate here the crystallization in thin films of this comb/graft DNA
copolymer composed of a chitosan backbone on which short
single stranded nucleotide sequences (ssDNA) are grafted. As
evidenced by imaging and spectroscopy, we show for the first
time that a DNA copolymer of suitable composition undergoes
crystallization due to hydrogen bonding in which the nucleic acids
engage.
wileyonlinelibrary.com/journal/pi
I Safir et al.
MATERIALS AND METHODS
Materials
Chitosan was purchased from Sigma Aldrich (Buchs, Switzerland) with a deacetylation degree above 75% and a molecular
weight (Mw ) between 50 000 and 160 000 Da. The ssDNA modified at the 5′ -end through a decyl spacer with a carboxylic acid
group (5′ -CTCTCTCTCTTT-3′ ), where C stands for cytosine and
T for thymine (5′ -(CT)5 T2 -3′ ), was purchased on controlled pore
glass (desalted, Mw 3763.9) from Microsynth (Balgach, Switzerland). The complementary sequence without any modification
(5′ -AAAGAGAGAGAG-3′ ), G stands for guanine and A for adenine (3′ -(GA)5 A2 -5′ ) (desalted, Mw 3776.2), was purchased from
Microsynth as well. Acetonitrile was purchased from Fisher Scientific (Waltham, MA, USA). Other chemicals such as salts and
organic solvents were purchased from Sigma Aldrich, Fluka
(Cressier, Switzerland), GE Healthcare (Glattbrugg, Switzerland),
Acros Organics (Geel, Belgium) and BioRad (Hercules, CA, USA)
and used without further purification. Mica was purchased as
high-grade quality V1 from Plano GmbH (Wetzlar, Germany);
silica wafers (100) were purchased from Silchem (Freiburg, Germany) and gold substrates (111) were purchased from PHASIS
(Geneva, Switzerland). The synthesis and characterization of the
chitosan-g-ssDNA copolymer has been described previously.39
DSC measurements to determine the crystallization temperature
(T c ) of the chitosan-g-ssDNA hybrid (50 ∘ C) were performed with a
DSC822e from Mettler Toledo (Schwerzenbach, Switzerland) from
0 to 110 ∘ C at a speed of 10 ∘ C min−1 .
Sample preparation
Due to the chemical dis-affinity between chitosan and the ssDNA,
solutions of the DNA copolymer were prepared in an aqueous solution of acetonitrile (pH 7.6) in which the chitosan-g-ssDNA copolymer is soluble. The solution of the copolymer was prepared by
first dissolving the chitosan-g-ssDNA copolymer in acetonitrile followed by the addition of ultra-pure water (MilliQ). Throughout the
whole study, the concentration of the solution was 1 mg mL−1 in
a 70:30 volume:volume ratio of acetonitrile and water. To perform
control experiments, a solution of chitosan at a concentration of
1 mg mL−1 in an acidic aqueous solution (pH 2) in which the polymer is fully soluble was prepared. Stock solutions of ssDNA and of
the ssDNA complementary sequence were prepared at a concentration of 1 mg mL−1 in pure water. Hybridization studies between
the ssDNA and its complementary sequence were conducted at
1:1 molar ratio in a 50 mmol L−1 NaCl aqueous solution.39
Thin films were prepared on three surfaces chosen for their
atomic flatness and their intrinsic physicochemical properties, in
particular their surface energy. Thin films were thus prepared
on surfaces differing in surface energy or atomic orientation, i.e.
hydrophobic gold (47 mJ m−2 ) and hydrophilic silica (2.3 × 10−3 mJ
m−2 ) and mica (0.12 mJ m−2 ) surfaces. Silica and gold are atomically oriented ((100) and (111) respectively), whereas mica cleaves
along the (001) plane. Cleaning of silica was performed by sonication in acetonitrile for 15 min and subsequent plasma treatment
for 15 min. The mica substrate is a phyllosilicate mineral of silica,
aluminium and potassium consisting of several parallel flat sheets.
The cleaning consists thus in removing one sheet with an adhesive tape. Gold surface cleaning was done by UV/ozone treatment
for 20 min. The thin film preparation was performed either by drop
casting or by spin coating (at a velocity of 2000 rpm for 1 min).
Samples were systematically annealed (Heraeus Materials Technology, Hanau, Germany) at 90 ∘ C for 1 h prior to slow cooling to room
temperature.
© 2016 Society of Chemical Industry
Polym Int 2016; 65: 1165–1171
Thin film crystallization of a DNA copolymer hybrid
Characterization
Cryo-transmission electron microscopy
Cryo-TEM analysis was performed with samples prepared by
adsorption of 4 μL of the sample on holey carbon-coated grids
(Quantifoil, Germany), blotted with Whatman 1 filter paper and
vitrified into liquid ethane at −178 ∘ C using a Vitrobot (FEI Company, Hillsboro, USA). Frozen grids were transferred onto a Philips
CM200-FEG electron microscope using a Gatan 626 cryo-holder.
Electron micrographs were recorded at an accelerating voltage of
200 kV and a nominal magnification of 50 000×, using a low-dose
system (10 e− Å−2 ) and keeping the sample at −175 ∘ C.
Imaging
The topography of the thin films on the different substrates was
obtained by AFM in the tapping mode (MFP-3D microscope,
Asylum Research, Santa Barbara, CA, USA) and by optical
microscopy (Olympus Corporation, Tokyo, Japan). For AFM, a
soft silicon nitride microcantilever (spring constant 1.75 N m−1 ,
resonance frequency ca 63 kHz; Olympus, Tokyo, Japan) was used
for scanning. For AFM, images were collected with an All-Digital
ARC2™ Controller and analysed with the Igor software.
Attenuated total reflectance infrared spectroscopy (ATR-IR)
ATR-IR spectra were measured with a Bruker VERTEX 80v Fourier
transform infrared spectrometer with a liquid-nitrogen-cooled
narrow-band mercury cadmium telluride detector. Spectra were
recorded at a resolution of 4 cm−1 . All experiments were performed at room temperature and the spectrometer was evacuated to avoid contributions from gas-phase water and CO2 . The
samples were deposited on a Ge internal reflection element (IRE)
(50 mm × 20 mm × 1 mm, Komlas). The IRE was first polished with
a 0.25 μm grain size diamond paste (Buehler, Metadi II) and afterwards rinsed copiously with Milli-Q water before the surface was
plasma cleaned under a flow of air for 2 min (Harrick Plasma instrument). The clean Ge IRE served as the reference for the ATR-IR spectra. Then spectra were measured by dropping 30 μL of solution on
the plasma cleaned Ge ATR crystal. In some experiments a ZnSe
wire grid polarizer was used.
RESULTS AND DISCUSSION
Polym Int 2016; 65: 1165–1171
(Fig. 1). 50 nm sized micellar structures assemble due to chemical
and physical dis-affinity between chitosan, which is a non-soluble
semi-flexible polymer segment, and the flexible water-soluble
ssDNA.39
Thin film properties
Morphological characterization
The only practical means of developing thin films (<100 nm) is
deposition from a solvent.1 Since the chitosan-g-ssDNA hybrid is
solvent cast on the substrate surface by dip or spin coating, solvent effects cannot be ignored. To limit these effects, the system was systematically heated at 90 ∘ C, a temperature above the
crystallization temperature (50 ∘ C), which also disrupts intermolecular hydrogen bonding. The most ideal ordering possible and
minimization of the total free energy of the system can thus be
expected.1 The nanopatterns in the resulting thin films might thus
essentially result from self-organization via microphase separation
of the block copolymers at the surface and not via micelle formation and related phenomena in solution.1
Figures 2(a) and 2(b) are respectively representative AFM and
optical images recorded upon chitosan-g-ssDNA hybrid structure
formation into single polygonal crystals assembled on gold substrates.
It is noticeable that the rate of formation of the structures is
instantaneous and congruent with solvent evaporation. The single
polygonal crystals are growing from a primary nucleus located
at the centre of the structures. Nucleation probably arises from
the formation of a small amount of crystalline material due to
fluctuations in material density or to a surface tension effect.40
From that point of primary nucleation, structures expand to six
branches, sometimes ending with a leaf-like structure. This step
is known to be the stage of secondary nucleation, referring to
continuation of crystallization at the growth front.41 The height of
the central nucleus of about 94 ± 15 nm is the highest point of
the structures. Statistical analysis on 20 different trunks enabled
the assessment of the thickness and height of the trunks forming
the single polygonal structures. The height of the trunks is 19
± 5 nm, while the width of the trunks is 484 ± 102 nm, much
larger than the micelles assembled in dilute aqueous solution (see
Fig. 1).39 This result indicates that the single polygonal structures
are thin and that the growth of the structure is homogeneous. As
the size of chitosan-g-ssDNA is about 6 nm,39 the trunks are about
two lamellae in height.
Since the growth of amphiphilic copolymer single crystals on
hydrophobic gold substrates occurs through adsorption of the
hydrophobic segment on the surface, leaving the hydrophilic
flexible segments extending outwards from the surface,42 the
chitosan thus probably adsorbs on the gold substrate whereas the
hydrophilic DNA extends outwards from the surface.
Figure 3(a) reveals the formation of crystal structures on silica as
imaged by AFM whereas Fig. 3(b) evidences the formation of comparable structures on mica, as observed by optical microscopy.
The structures formed on both hydrophilic surfaces exhibit comparable well-ordered side-branched structures homogeneously
distributed on the surface. As imaged in Fig. 3, there are two
sizes of side-branches, larger and secondary smaller trunks. Statistical analyses were performed on 20 different trunks to assess
the thickness and height of the side-branches. The larger trunks
are 1322 ± 341 nm wide whereas the height is 70 ± 17 nm. The
smaller lateral trunks are 762 ± 148 nm wide and 67 ± 15 nm in
height. Side-branches are thus homogeneous in height but differ
in width. The thickness of the trunks is therefore formed by almost
© 2016 Society of Chemical Industry
wileyonlinelibrary.com/journal/pi
1167
Solution properties
The chitosan − DNA hybrid copolymer was synthesized according to chemical routes published previously.39 Briefly, the size of
commercially available high molecular weight chitosan is reduced
by enzymatic digestion to 4425 Da (polydispersity index 1.3) prior
to coupling by solid phase synthesis to the 12 nucleotide long
5′ -CTCTCTCTCTTT-3′ sequence (C and T stand for cytosine and
thymine respectively) modified at the 5′ -end by a carboxylic acid
group spaced by a decyl spacer. Conventional chemical characterizations evidence the synthesis of a chitosan-g-ssDNA copolymer of
7908 Da according to the macromolecular configuration described
in Scheme 1; on average, one nucleic acid sequence is grafted
along the chitosan backbone.39
Chitosan is soluble in aqueous solutions of pH at which the primary amine along the backbone is positively charged, i.e. below
pH 5. At this pH, the phosphate groups along the nucleic acid acids
are negatively charged. The resulting chitosan-g-ssDNA hybrid is
therefore soluble in either aqueous solutions of low pH or an aqueous solution of acetonitrile. Structure formation in dilute aqueous solution takes place at neutral pH as evidenced by cryo-TEM
www.soci.org
www.soci.org
I Safir et al.
OH
O
*
HO
O
O P O
O
O
O
O
N
O
O P
O
O
O
O
O
N
N
O
N
O
NH2
O
5
C10H20
O
HOOC
O
O
N
N
O
O
N
O
O P
O
O P O
O
HO HN
x
O
COCH3
O
OH
O
yO
OH
O
(i) EDC/NHS
Chitosan-NH2
MES buffer, pH=5
RT, 24h
x = 0.046
y = 0.82
z = 0.125
m = 25
(ii) 34% NH4OH, 24 h
40 oC
N
N
*
NH z m
HO
O
O P O
= CPG
O
N
N
O
N
O
O
O P O
O
O
N
O
N
O
O
5
O P O
O
O
N
N
O
O
O
O P O
O
O
N
N
O
O
OH
Scheme 1. Synthesis scheme of the chitosan-grafted ssDNA copolymer hybrid.
1168
six layers of chitosan-g-ssDNA lamellae in the direction perpendicular to the substrate. The width of the trunks, which is much
larger than the chitosan-g-ssDNA hybrid, reflects the interplay
between the transport process of still molten molecules toward
the crystal surface and the probability that these molecules get
attached to the crystal and stay there.43 The chains must thus
fold back and forth into stems with chains oriented toward the
surface.44 Therefore, the height of chitosan-g-ssDNA structures
consists in several layers of lamellae, whereas in width a larger
distribution of the number of lamellae occurs. Structures of similar
morphology, i.e. side-branched structures, have been described
previously in the literature.45 These are formed from dendritic
trunks and side-branches as revealed by AFM. The branches are
initially formed at an angle of near 45∘ and turn back toward the
preferred 90∘ direction,46 as observed in Fig. 3 as well.
Single polygonal structures thus grow specifically on the
hydrophobic gold substrate, while side-branched structures grow
on hydrophilic surfaces. The orientation of the structures on
the surfaces is thus drastically influenced by the tribology of
the surface, especially its surface energy. As reported by Sutton
et al.,40 the substrate plays an important role in the final morphology of lamellar crystals. In general, semicrystalline polymers
can crystallize on a substrate, whereby molecular chains fold
wileyonlinelibrary.com/journal/pi
back and forth into stems to form crystal lamellae. However,
two preferred polymer orientations are commonly found: (1)
edge-on lamellae, in which the fold surface is perpendicular to
the substrate and the molecular chain axis is parallel to the substrate, and (2) flat-on lamellae, in which the fold surface is parallel
to the substrate and the molecular chain axis is normal to the
substrate. Usually single polygonal crystals and side-branched
structures are perpendicular to the surface (flat-on) according to
chain orientation.47
The difference in crystal morphology observed upon crystallization of chitosan-g-ssDNA on either gold or mica and silica is thus
clearly due to the specific interaction of chitosan-g-ssDNA segments with the surface. Chitosan interacts preferentially with gold
whereas ssDNA extends outwards from the surface. On silica and
mica, the ssDNA is adsorbed on the surface, and the hydrophobic
segments probably associate with each other.48
Crystallization versus phase separation: investigation of the blends
of chitosan and ssDNA
Since dendritic structures have been reported previously by Okerberg et al.46 by crystallization of PEO blended with PMMA, chitosan was blended with ssDNA and thin film formation was
induced. As can be observed in Fig. S1, an amorphous area can be
© 2016 Society of Chemical Industry
Polym Int 2016; 65: 1165–1171
Thin film crystallization of a DNA copolymer hybrid
www.soci.org
Figure 1. Cryo-TEM imaging of micellar structures assembled by chitosan-g-ssDNA.
Figure 2. Single polygonal crystals assembled by chitosan-g-ssDNA hybrids on a gold surface as observed by (a) AFM and (b) optical microscopy (10×/0.25,
magnification/numerical aperture).
distinguished, which besieged the ssDNA cross-like structures. No
side-branched structures were observed, which is a first indication
that crystallization occurs owing to the covalent binding between
the chitosan and ssDNA. No intermolecular electrostatic interaction or hydrogen bonding occurs. As can be seen from the images
in Fig. S2, structure formation of (a) ssDNA with a decyl spacer, (b)
ssDNA without the spacer and (c) chitosan does not occur. ssDNA
with and without a spacer forms cross-like structures. These results
further indicate that ssDNA might drive the structure formation of
chitosan-g-ssDNA in thin films.
Polym Int 2016; 65: 1165–1171
Chemical interactions as studied by ATR-IR spectroscopy
To assess the role of hydrogen bonding in chitosan-g-ssDNA crystallization in thin films, ATR-IR spectroscopy was performed with
chitosan, DNA, a blend of both and the chitosan − DNA hybrid
(with parallel and perpendicular polarized light). Crystallization
occurs on the IRE (Fig. S5).
© 2016 Society of Chemical Industry
wileyonlinelibrary.com/journal/pi
1169
Chemical interactions as studied by blending the chitosan-g-ssDNA
with the ssDNA complementary nucleotide sequence
by hybridization
To evidence the role of DNA in the thin film crystallization of
chitosan-g-ssDNA, hybridization with the ssDNA complementary
sequence of the graft (3′ -(GA)5 A2 -5′ ) was performed in solution
prior to thin film formation. As displayed in Fig. 4, hybridization
disables the formation of the side-branched structures.
Short-range specific hydrogen bonding therefore plays a crucial role in the formation of chitosan-g-ssDNA crystals in thin
films. Since hybridization occurs at a given ionic strength to
screen the electrostatic repulsion between like charged nucleic
acid strands, the effect of the long-range electrostatic force on
chitosan-g-ssDNA crystallization was further investigated. As
displayed in Fig. S3, dendrite-like structures are formed on a mica
surface but no side-branched structures were observed upon
hybridization in the absence of salt. Hybridization of molecularly
dissolved chitosan-g-ssDNA with the complementary sequence
of ssDNA without NaCl does not occur. However, dendritic structures are observed. A morphological transition is observed as
expected by blending with the ssDNA complementary sequence.
This result further confirms the dependence of the formation
of side-branched crystal structures on hydrogen bonding. Since
chitosan-g-ssDNA copolymer contains positive and negative
charges along the chitosan and the ssDNA segments, thin film
formation was induced in 50 mmol L−1 NaCl. As displayed in Fig.
S4, discontinuous side-branched structures are formed. This result
indicates that crystallization is sensitive to electrostatic interaction
but not hindered. Hydrogen bonding plays the crucial role at short
distances.
www.soci.org
I Safir et al.
Figure 3. Imaging reveals side-branched crystal structures assembled by chitosan-g-ssDNA on (a) silica, as assessed by AFM, and (b) mica, as observed by
optical microscopy (50×/0.50, magnification/numerical aperture).
Figure 4. AFM imaging of crystallization induced on mica reveals destruction of chitosan-g-ssDNA side-branched crystal structures upon hybridization of the chitosan-g-ssDNA hybrid with the ssDNA complementary
sequence.
1170
The comparison in Fig. 5 shows that the IR spectrum of the chitosan − DNA hybrid is completely different from those of its constituents (chitosan and DNA). In particular most of the bands of
the hybrid are very sharp in contrast to the spectra of the DNA
and chitosan alone. This reflects the very ordered structure within
the chitosan-g-ssDNA hybrid film. In addition, band shifts indicate
strong interactions within the film. The O − H and N − H stretching
region is particularly affected with a strong red-shift of intensity
for the hybrid film, which indicates prominent hydrogen bonding as deduced from the results observed when crystallization was
induced with the hybridized hybrid. Moreover, the polarized measurements lead to the conclusion that at least some of the functional groups are oriented with respect to the IRE surface. This is
easily appreciated when the spectra measured with parallel and
perpendicular polarized light are scaled to approximately the same
intensity (Fig. 6).
Note that a quantitative analysis of orientation is complicated
since the hybrid sample can be treated neither as a thin film
nor as a bulk medium. In addition, the film does not have a
homogeneous thickness. Therefore the analysis remains qualitative. When comparing the two spectra it is evident that the relative intensities of the vibrational bands are not identical for the
two polarizations. Identical relative intensities of all bands would
be observed for a completely isotropic film. This is clearly not the
case as can be verified for example for the bands at 1230 cm−1 ,
wileyonlinelibrary.com/journal/pi
Figure 5. ATR-IR spectra of chitosan in brown, DNA in green, a mixture of
both in blue and chitosan − DNA hybrid (red, parallel polarized light; black,
perpendicular polarized light; these two were artificially shifted up the y
axis for clarity).
1206 cm−1 , 1172 cm−1 and 1080 cm−1 . A more detailed analysis
requires assignment of vibrational bands and knowledge of the
direction of the corresponding transition dipole moments within
the molecule, which is not easy for such a complicated structure. The band associated with C = O stretching vibrations around
1700 cm−1 is relatively strong in the parallel polarized spectrum
indicating that the corresponding transition dipole moment is
preferentially oriented perpendicular to the IRE surface. The strong
band around 1230 cm−1 can be assigned to an antisymmetric PO2 −
vibration. Obviously this band consists of several components with
different polarization dependence. The high wavenumber component is more pronounced for parallel polarized light and therefore
has a stronger component of the dynamic dipole moment perpendicular to the IRE surface compared to the low wavenumber
component.
CONCLUSIONS
Crystallization of chitosan-g-ssDNA induces the formation of two
different types of crystal structures according to the surface
energy of the substrate. On gold, chitosan-g-ssDNA crystallizes
into single polygonal structures while on hydrophilic silica and
mica substrates it crystallizes into side-branched crystal structures.
© 2016 Society of Chemical Industry
Polym Int 2016; 65: 1165–1171
Thin film crystallization of a DNA copolymer hybrid
Figure 6. Polarized ATR-IR spectra of chitosan − DNA hybrid film on Ge IRE.
Red, parallel polarized light; black, perpendicular polarized light.
This result depends on the interaction of either the hydrophilic
or the amphiphilic segments of the copolymer with either the
hydrophilic or hydrophobic surfaces.
With help of ATR-IR measurements, the role of hydrogen bonding
on the morphology of the crystals was further evidenced.
ACKNOWLEDGEMENTS
The Swiss National Science Foundation (SNSF PPOP2-153025) and
the University of Geneva are greatly acknowledged for financial support. We are very grateful for the experimental support of Professor Michal Borkovec and Dr Plinio Maroni for AFM
measurements.
SUPPORTING INFORMATION
Supporting information may be found in the online version of this
article.
REFERENCES
1 Farrell RA, Fitzgerald TG, Borah D, Holmes JD and Morris MA, Int J Mol
Sci 10:3671–3712 (2009).
2 He W-N and Xu J-T, Prog Polym Sci 37:1350–1400 (2012).
3 Arbe A and Colmenero J. Macromolecules 45:491–502 (2012).
4 Kim YY, Ahn B, Sa S, Jeon M, Roth SV, Kim SY et al., Macromolecules
46:8235–8244 (2013).
5 Jiang L, Wu J, Nedolisa C, Saiani A and Assender HE, Macromolecules
48:5358–5366 (2015).
6 Chen N, Yan LT and Xie XM, Macromolecules 46:3544–3553 (2013).
7 Michell RM, Blaszczyk-Lezak I, Mijangos C and Mueller AJ. Polymer
(Guildf) 54:4059–4077 (2013).
8 Yu PQ, Yan LT, Chen N and Xie XM. Polymer (Guildf) 53:4727–4736
(2012).
9 Hölzer S, Büttner TN, Schulze R, Arras MML, Schacher FH, Jandt KD et al.,
Eur Polym J 68:10–20 (2015).
10 Schönherr H, Waymouth RM and Frank CW, Macromolecules
36:2412–2418 (2003).
11 Schönherr H and Frank C, Macromolecules 36:1199–1208 (2003).
12 Carvalho JL, Massa MV, Cormier SL, Matsen MW, Dalnoki-Veress K, Eur
Phys J E34 (2011).
www.soci.org
13 Arora H, Du P, Tan KW, KHJ, Grazul J, Xin HL et al., Science 330:214–219
(2012).
14 Wen T, Liu G, Zhou Y, Zhang X, Wang F, Chen H et al., Macromolecules
45:5979–5985 (2012).
15 Yang H, Zhang R, Wang L, Zhang J, Yu X, Liu J et al., Macromolecules
48:7557–7566 (2015).
16 Huang S and Jiang S, RSC Adv 4:24566 (2014).
17 Novoa-Carballal R and Muller AHE, Chem Commun 48:3781–3783
(2012).
18 Engler AC, Bonner DK, Buss HG, Cheung EY and Hammond PT, Soft
Matter 7:5627–5637 (2011).
19 Kempe K, Neuwirth T, Czaplewska J, Gottschaldt M, Hoogenboom R
and Schubert US, Polym Chem 2:1737–1743 (2011).
20 Kwak M, Herrmann A, Nucleic acid polymers and DNA synthetic
polymer hybrid materials generated by molecular biology techniques, in Synthesis of Polymers: New Structures and Methods, ed. by
Schlueter AD, Hawker CJ and Sakamoto J. Wiley-VCH, Weinheim
(2012).
21 Robinson JW, Secker C, Weidner S and Schlaad H, Macromolecules
46:580–587 (2013).
22 Kim YY, Jung S, Kim C, Ree BJ, Kawato D, Nishikawa N et al., Macromolecules 47:7510–7524 (2014).
23 Malek A, Dingenouts N, Beskers TF, Fehrenbacher U, Barner L and
Wilhelm M, Eur Polym J 49:2704–2720 (2013).
24 Bahloul M, Chamignon C, Pruvost S, Fleury E, Charlot A and Portinha
D. Polymer (Guildf) 79:195–204 (2015).
25 Kamitakahara H, Baba A, Yoshinaga A, Suhara R and Takano T, Cellulose
21:3323–3338 (2014).
26 Chiang Y-W, Hu Y-Y, Li J-N, Huang S-H, Kuo S-W, Macromolecules
48:8526–8533 (2015).
27 Domard A, Carbohydr Polym 84:696–703 (2011).
28 Ding X, Richter DL, Matuana LM and Heiden PA, Carbohydr Polym
86:58–64 (2011).
29 Cao J, Huang S, Chen Y, Li S, Li X, Deng D et al., Biomaterials
34:6272–6283 (2013).
30 Dao-Lu TFS, Cheng C , Xiu-Li W and Yu-Zhong W, Nanotechnology
24:145101 (2013).
31 Elsaid N, Jackson TL, Gunic M and Somavarapu S, Invest Ophthalmol Vis
Sci 53:8105–8111 (2012).
32 Lian H, Sun J, Yu YP, Liu YH, Cao W, Wang YJ et al., Int J Nanomed
6:3323–3334 (2011).
33 Lu HW, He H, Zhang B, Liu GQ, Li MY and Nie QL, J Appl Polym Sci
130:908–915 (2013).
34 Moyuan C, Haixia J, Weijuan Y, Peng L, Liqun W and Hongliang J, J Appl
Polym Sci 123:3137–3144 (2012).
35 X-L Wang, Y-L Z, DL Tang, G-Y Liu and Y-Z Wang, J Polym Res 19: (2012).
36 Zhang J, Li M, Fan T, Xu Q, Wu Y, Chen C et al., J Polym Res 20:1–11
(2013).
37 Zhou W, Wang Y, Jian J and Song S, Int J Nanomed 8:3715–3728 (2013).
38 Liu C, Zhu Q, Wu W, Xu X, Wang X, Gao S et al., Int J Nanomed
7:5339–5350 (2012).
39 Safir I, Ngo K-X, Nixon Abraham J, Ghahraman Afshar M, Pavlova E and
Nardin C. Polymer (Guildf) 79:29–36 (2015).
40 Sutton SJ, Izumi K, Miyaji H, Miyamoto Y and Miyashita S, J Mater Sci
32:5621–5627 (1997).
41 Zachmann H, Pure Appl Chem 38:79 (1974).
42 Freij-Larsson C, Nylander T, Jannasch P and Wesslén B, Biomaterials
17:2199 (1996).
43 Reiter G, J Polym Sci B Polym Phys 41:1869 (2003).
44 Muthukumar M, Adv Chem Phys 123:1–63 (2004).
45 Organ SJ, Keller A, J Mater Sci Mater Electron 20:1571–1585 (1985).
46 Okerberg BC, Marand H and Douglas JF. Polymer (Guildf) 49:579 (2008).
47 Ophelia KC and TPR Tsui (eds), Polymer Thin Films, Series in Soft
Condensed Matter, eds. Tsui OKC and Russell TP, World Scientific,
Singapore. vol. 1, 312 pp. (2008).
48 Liu X, Song J, Wu D, Genzer J, Theyson T and Rojas OJ, Ind Eng Chem
Res 49:8550 (2010).
1171
Polym Int 2016; 65: 1165–1171
© 2016 Society of Chemical Industry
wileyonlinelibrary.com/journal/pi