Bioinspired Materials Science and Engineering E-Book

• Herausgeber: John Wiley & Sons
• Kategorie: Fachliteratur
• Sprache: Englisch

Seitenzahl: 1031

Veröffentlichungsjahr: 2018

Beschreibung

An authoritative introduction to the science and engineering of bioinspired materials Bioinspired Materials Science and Engineering offers a comprehensive view of the science and engineering of bioinspired materials and includes a discussion of biofabrication approaches and applications of bioinspired materials as they are fed back to nature in the guise of biomaterials. The authors also review some biological compounds and shows how they can be useful in the engineering of bioinspired materials. With contributions from noted experts in the field, this comprehensive resource considers biofabrication, biomacromolecules, and biomaterials. The authors illustrate the bioinspiration process from materials design and conception to application of bioinspired materials. In addition, the text presents the multidisciplinary aspect of the concept, and contains a typical example of how knowledge is acquired from nature, and how in turn this information contributes to biological sciences, with an accent on biomedical applications. This important resource: * Offers an introduction to the science and engineering principles for the development of bioinspired materials * Includes a summary of recent developments on biotemplated formation of inorganic materials using natural templates * Illustrates the fabrication of 3D-tumor invasion models and their potential application in drug assessments * Explores electroactive hydrogels based on natural polymers * Contains information on turning mechanical properties of protein hydrogels for biomedical applications Written for chemists, biologists, physicists, and engineers, Bioinspired Materials Science and Engineering contains an indispensible resource for an understanding of bioinspired materials science and engineering.

Leseprobe

Table of Contents

Cover

Foreword

Preface

Introduction to Science and Engineering Principles for the Development of Bioinspired Materials

I.1 Bioinspiration

I.2 Bioinspired Materials

I.3 Biofabrication

I.4 Biofabrication Strategies

I.5 Part II Biomacromolecules

I.6 Part III Biomaterials

I.7 Scope of the Book

Acknowledgments

References

Part I: Biofabrication

1 Biotemplating Principles

1.1 Introduction

1.2 Mineralization in Nature

1.3 Petrified Wood in Construction and Technology

1.4 Structural Description and Emulation

1.5 Characteristic Parameters

1.6 Applications

1.7 Limitations and Challenges

1.8 Conclusion and Future Topics

Acknowledgments

References

2 Tubular Tissue Engineering Based on Microfluidics

2.1 Introduction

2.2 Natural Tubular Structures

2.3 Microfluidics

2.4 Fabrication of Tubular Structures by Microfluidics

2.5 Conclusion

Acknowledgments

References

3 Construction of Three‐Dimensional Tissues with Capillary Networks by Coating of Nanometer‐ or Micrometer‐Sized Film on Cell Surfaces

3.1 Introduction

3.2 Fabrication of Nanometer‐ and Micrometer‐Sized ECM Layers on Cell Surfaces

3.3 3D‐Tissue with Various Thicknesses and Cell Densities

3.4 Fabrication of Vascularized 3D‐Tissues and Their Applications

3.5 Conclusion

Acknowledgments

References

4 Three‐dimensional Biofabrication on Nematic Ordered Cellulose Templates

4.1 Introduction

4.2 What Is Nematic Ordered Cellulose (NOC)?

4.3 Exclusive Surface Properties of NOC and Its Unique Applications

4.4 Conclusion

References

5 Preparation and Application of Biomimetic Materials Inspired by Mussel Adhesive Proteins

5.1 Introduction

5.2 Various Research Studies

5.3 Conclusion

References

6 Self‐assembly of Polylactic Acid‐based Amphiphilic Block Copolymers and Their Application in the Biomedical Field

6.1 Introduction

6.2 Micellar Structures from PLA‐based Amphiphilic Block Copolymers

6.3 Hydrogels from PLA‐based Amphiphilic Block Copolymers

6.4 Conclusion

Acknowledgments

References

Part II: Biomacromolecules

7 Electroconductive Bioscaffolds for 2D and 3D Cell Culture

7.1 Introduction

7.2 Electrical Stimulation

7.3 Electroconductive Bioscaffolds

7.4 Conclusion

Acknowledgments

References

8 Starch and Plant Storage Polysaccharides

8.1 Starch and Other Seed Polysaccharides: Availability, Molecular Structure, and Heterogeneity

8.2 Effect of the Molecular Structure of Starch and Seed Polysaccharides on the Macroscopic Properties of Derived Carbohydrate‐based Materials

8.3 Chemo‐enzymatic Modification Routes for Starch and Seed Polysaccharides

8.4 Conclusion

References

9 Conformational Properties of Polysaccharide Derivatives

9.1 Introduction

9.2 Theoretical Backbone to Determine the Chain Conformation of Linear and Cyclic Polymers from Dilute Solution Properties

9.3 Chain Conformation of Linear Polysaccharides Carbamate Derivatives in Dilute Solution

9.4 Lyotropic Liquid Crystallinity of Polysaccharide Carbamate Derivatives

9.5 Cyclic Amylose Carbamate Derivatives: An Application to Rigid Cyclic Polymers

9.6 Conclusion

Appendix: Wormlike Chain Parameters for Polysaccharide Carbamate Derivatives

References

10 Silk Proteins

10.1 Introduction

10.2 Bio‐synthesis of Silk Proteins

10.3 Extraction of Silk Proteins

10.4 Structure and Physical Properties of Silk Proteins

10.5 Properties of Silk Proteins in Biomedical Applications

10.6 Processing Silk Fibroin for the Preparation of Biomaterials

10.7 Processing Silk Sericin for Biomaterials Applications

10.8 Conclusion

Acknowledgments

Abbreviations

References

11 Polypeptides Synthesized by Ring‐opening Polymerization of N‐Carboxyanhydrides

11.1 Introduction

11.2 Living Polymerization of NCAs

11.3 Synthesis of Traditional Copolypeptides and Hybrids

11.4 New Monomers and Side‐Chain Functionalized Polypeptides

11.5 The Self‐assembly of Polypeptides

11.6 Novel Bio‐related Applications of Polypeptides

11.7 Conclusion

References

12 Preparation of Gradient Polymeric Structures and Their Biological Applications

12.1 Introduction

12.2 Gradient Polymeric Structures

12.3 Gradient Polymeric Structures Regulated Cell Behavior

12.4 Conclusion

References

Part III: Biomaterials

13 Bioinspired Materials and Structures

13.1 Introduction

13.2 Fiber‐reinforced Structures Inspired by Unbranched and Branched Plant Stems

13.3 Pomelo Peel as Inspiration for Biomimetic Impact Protectors

13.4 Self‐repair in Technical Materials Inspired by Plants’ Solutions

13.5 Elastic Architecture: Lessons Learnt from Plant Movements

13.6 Conclusions

Acknowledgments

References

14 Thermal‐ and Photo‐deformable Liquid Crystal Polymers and Bioinspired Movements

14.1 Introduction

14.2 Thermal‐responsive CLCPs

14.3 Photothermal‐responsive CLCPs

14.4 Light‐responsive CLCPs

14.4 Conclusion

References

15 Tuning Mechanical Properties of Protein Hydrogels

15.1 Introduction

15.2 What Are Different about Proteins?

15.3 Protein Cross‐linking

15.4 Strategies for Mechanical Reinforcement

15.5 Conclusion

References

16 Dendritic Polymer Micelles for Drug Delivery

16.1 Introduction

16.2 Dendrimers

16.3 Hyperbranched Polymers

16.4 Dendrigraft Polymers

16.5 Conclusion

References

17 Bone‐inspired Biomaterials

17.1 Introduction

17.2 Bone

17.3 Bone‐like Materials

17.4 Bone‐like Scaffolds

17.5 Conclusion

References

18 Research Progress in Biomimetic Materials for Human Dental Caries Restoration

18.1 Introduction

18.2 Tooth Structure

18.3 The Formation Mechanism of Dental Caries

18.4 HA‐filled Biomimetic Resin Composites

18.5 Biomimetic Synthesis of Enamel Microstructure

Acknowledgments

References

Index

End User License Agreement

List of Tables

Introduction

Table I.1 Typical biological materials with function integration.

Table I.2 Comparison of four types of bioprinting techniques.

Chapter 01

Table 1.1 Comparison of the mass increase due to multiple infiltrations with constant masses of dissolved ethanolic tetraethyl orthosilicate precursor as opposed to single infiltrations with larger amounts.

Table 1.2 Comparison of levels of hierarchy found in selected natural materials [230]. Note that plant elementary fibrils are sometimes termed micro‐ [231] or nanofibrils, and subsequently microfibrils become macrofibrils [233], or just fibrils [232].

Table 1.3 Comparison of the processing steps of two biotemplated materials and each one alternative processing route. The columns termed ‘Complexity’ are assigned to the processes to their left as a measure of the work time required.

Chapter 07

Table 7.1 Some examples of vertebrate cells’ responses to weak DC‐EFs.

Table 7.2 Conductive polymers‐based bioscaffolds used in cell culture and tissue engineering, based on the literature.

Table 7.3 CNTs‐based bioscaffolds used in cell culture and tissue engineering, based on the literature.

Table 7.4 Graphene and graphene oxide‐based bioscaffolds used in cell culture and tissue engineering.

Chapter 08

Table 8.1 Composition and relative amounts (in % dry weight) of AX and mixed‐linkage β‐glucans in different cereal tissues.

Chapter 09

Table 9.1 The Kuhn segment length

λ

−1

and the helix pitch per residue

h

for ATPC in 1,4‐dioxane, CTPC in THF, and curdlan tris(phenylcarbamate) (CdTPC) in THF at 25 °C.

Table 9.2 Helix pitch per residue

h

and the Kuhn segment length

λ

−1

for polysaccharide carbamate derivatives.

Chapter 18

Table 18.1 Evaluation of dental resin composites with microscopic and nanoscopic HA particles [26].

List of Illustrations

Chapter 01

Figure 1.1 Scheme of the deposition of inorganic phases (red) in a biological material (black, adaptation of the wood structure) on several levels of hierarchy. The shown levels of hierarchy may themselves contain sublevels. For example, in the actual wood structure, the nm scale contains the hierarchical levels of the cellulose micro‐ and elementary fibrils.

Figure 1.2 Micrometer‐scale structures of coal, showing several well‐replicated features such as (a, b) the wood cells, (c) ray fibril intersections, and (d) pit holes.

Figure 1.3 Scheme of the general pathway to creating biotemplated materials starting from a living object. The boxed steps are those followed during the artificial petrifaction of structures. The surface functionalization is an optional step; the deposition may be reactive, or purely sterical, as outlined in the text.

Figure 1.4 Deconvolution of a thermal analysis curve (in air, solid line) of an artificially mineralized biological template (phenylated silicon dioxide on wood) [213] by Gaussian cumulative distribution functions (dashed lines) to give a fit of the curve with a goodness

R

2

 = 0.9999. The mass losses associated with the proposed processes are (low temperature to high): 4.4 Ma% loss of absorbed moisture, 29.8 Ma% and 27.7 Ma% stepwise devolatilization of the wood structure,

1

22.7 Ma% combustion of the carbonized material, 3.6 Ma% combustion of the temperature‐resistant phenyl substituents, 11.9 Ma% ash. The vertical dotted lines mark appropriate holding temperatures for a gentle template removal.

Figure 1.5 Structural analysis of the mineralized skeletal system of

Euplectella sp

. (A) Photograph of the entire skeleton, showing cylindrical glass cage. Scale bar, 1 cm. (B) Fragment of the cage structure showing the square‐grid lattice of vertical and horizontal struts with diagonal elements arranged in a chessboard manner. Orthogonal ridges on the cylinder surface are indicated by arrows. Scale bar, 5 mm. (C) Scanning electron micrograph (SEM) showing that each strut (enclosed by a bracket) is composed of bundled multiple spicules (the arrow indicates the long axis of the skeletal lattice). Scale bar, 100 µm. (D) SEM of a fractured and partially HF‐etched

1

single beam revealing its ceramic fiber‐composite structure. Scale bar, 20 mm. (E) SEM of the HF‐etched junction area showing that the lattice is cemented with laminated silica layers. Scale bar, 25 mm. (F) Contrast‐enhanced SEM image of a cross‐section through one of the spicular struts, revealing that they are composed of a wide range of different‐sized spicules surrounded by a laminated silica matrix. Scale bar, 10 mm. (G) SEM of a cross‐section through a typical spicule in a strut, showing its characteristic laminated architecture. Scale bar, 5 mm. (H) SEM of a fractured spicule, revealing an organic interlayer. Scale bar, 1 mm. (I) Bleaching of biosilica surface revealing its consolidated nanoparticulate nature (25). Scale bar, 500 nm.

Figure 1.6 Simulated small‐angle X‐ray scattering patterns from closest‐packed arrangements of each 500 fibrillar structures (with angular uniformity of σ

ang

:0.2 °

−1

), 2 nm in diameter, embedded in cylindrical cell walls, which were probed centrally, accounting for isotropic scattering from randomly oriented structures (10 nm) and for a Gaussian beam profile (σ

beam

:8 nm

−1

) [282]. The simulated microfibril angles (cylinder front to back) are (a) 0 °, (b) 2 °, (c) 3 ° and (d) 35 °. The separation of the directional scattering patterns originating from the fibrils can, under such practical conditions, be observed starting at 3 °.

Figure 1.7 Scheme of the free energy over particle radius for alumina, titania, zirconia and silica and their metastable, often amorphous, polymorphs, showing that the free energy barrier for nucleation is lower for metastable (often amorphous) polymorphs [183].

Figure 1.8 Comparison of the photonic structures found in

Entimus imperialis

and

granulatus

. While the scales from both animals show a similar internal D‐surface structure, they differ in their domain boundary, and outer shell cover. This leads to a different appearance of the beetle, (a)

imperialis

being brilliant and (b)

granulatus

dull.

Chapter 02

Figure 2.1 Schematic illustration of tubular tissue engineering on microfluidics by mimicking various natural tubular structures.

Figure 2.2 The principle of μCP, μFP, and LFP. (a) Schematic demonstration of the process of μCP. Step 1, pour PDMS prepolymer over master and cure it to form a PDMS stamp; Step 2, dip the PDMS stamp into patterning solution; Step 3, press PDMS stamp onto the substrate for seconds to minutes; Step 4, Lift up the stamp and leave patterns on the substrate. (b) Schematic demonstration of the process of μFP. Step 1, pour PDMS prepolymer over master and cure it to form a PDMS mold containing microchannels; Step 2, put the PDMS mold and substrate together to form sealed microchannels; Step 3, introduce different solutions into different microchannels to deposit components onto the substrate; Step 4, wash away the solution and leave components on the substrate to form patterns in confined areas.

Figure 2.3 (a) The azobenzene moiety can be converted photochemically between the E and Z configurations to either present or mask the RGD ligand and hence modulate cell adhesion. Cells adhered onto SAMs with the azobenzene group in the E configuration. Few cells adhered to the same SAMs with azobenzene in the Z configuration. Cells adhered to the SAMs again when the conformation of azobenzene was changed from Z to E. (b) Schematics of fabrication processes of the multilayered microchip and 3D reconstruction of the multilayered neurite connections. The inset represents the orthogonal neurite pathways in the brain. Copyright 2012, American Association for the Advancement of Science. (c) Schematic of the microfluidic flow‐stretch chip. The stress fibers tend to align in the direction parallel to that of the FSS, CS and the resultant force of the FSS and CS.

Figure 2.4 (a) Schematic illustration of the fabrication process of tubes, tubes‐in‐a‐tube and spirals. (b) Schematic illustration of a stress‐induced rolling membrane (SIRM) and tubes with multiple types of cells as the walls. A thin PDMS membrane is stretched as the top layer of the SIRM and covers a semi‐cured PDMS membrane. After curing the two layers to cause adhesion, a SIRM is obtained that rolled up when the ends were released. Microfluidic channels cover the surface of the SIRM, different cells are delivered via microfluidic channels to the surface of the SIRM. One end of the SIRM is released by cutting its edge. The SIRM rolls up into a tube and each type of cell is delivered to a designated position as the tubular wall. The structure of the tube is similar to that of tubular tissues, such as small‐diameter vessels. (c) Schematic diagram of cell surface modification and stepwise formation of multicellular structures. Confocal image of a 3D reconstruction of the bilayer on the SIRM after rolling.

Chapter 03

Figure 3.1 Schematic illustration of fabrication of ECM layers on cell surfaces by (left) nanometer‐sized FN/G films and (right) micrometer‐sized collagen nanofiber matrices to construct 3D‐ tissue models with higher and lower cell densities.

Figure 3.2 (a) Phase and fluorescent microscopic images of L929 mouse fibroblast cells coated with nanometer‐sized films by 9‐step assembly of Rh‐FN and FITC‐G. Scale bars in the figure are 10 mm. (b) Frequency shift of the quartz crystal microbalance (QCM) LbL assembly of FN‐G nanofilms onto a phospholipid bilayer. Closed (●) and open circles (○) represent the assembly steps of FN and G, respectively. The phospholipid bilayer was fabricated on a base layer prepared by PDDA and PSS. (c) Fluorescence intensity of the cells and 7‐step‐assembled Rh‐FN‐G nanofilms on cell surfaces observed by CLSM. (d) Schematic illustration of cells after coating for collagen nanofiber matrix once, twice, and three times with rhodamine or FITC‐labeled collagen. (e) From left to right are the coating results of once, twice, and three‐times‐ coated cells observed by CLSM. (f) line scanning results of collagen‐coated cells with different sizes of collagen microlayers on cell surfaces.

Figure 3.3 (a) Fluorescence microscopic image Rh‐FN‐FITC‐G films prepared on L929 cells after 24 h of incubation. (b) SEM image of the L929 cells with FN‐DS films after 24 h of incubation. NHDF cells (c) with and (d) without micrometer‐sized collagen fiber coating procedures were observed by LV‐SEM.

Figure 3.4 Phase contrast images of (a) the L929 fibroblasts without or with various nanofilms on the cell surfaces and (c) cells after no coating, and cells coated with 0.03 wt% once, twice, and three times after 24 h of incubation. Cell proliferation in (b) various nanofilms prepared on the L929 cell surfaces and (d) various micrometer‐sized collagen fiber matrix on the cell surfaces during 72 h of incubation were also evaluated.

Figure 3.5 Histological staining images and schematic illustration of cell densities inside 3D tissues constructed by (a) cells without coating, (b) FN‐G coated cells, and (c) collagen‐coated cells (coated once), using the cell accumulation technique.

Figure 3.6 Thickness of constructed 3D tissues made of cells coated once, twice, or three times with collagen nanofibers with various numbers of seeded cells after 24 h of incubation.

Figure 3.7 HE staining results of 3D tissues constructed with (a) FN‐G nanofilm‐coated cells and (b) micrometer‐sized collagen fiber‐coated cells with various seeding cell numbers after a 24 h of incubation.

Figure 3.8 Images of confocal laser scanning microscopy (CLSM) of vascularized 3D ‐tissue models constructed by (a) nano‐ and (b) micro‐coatings by sandwich culture. Immunohistological staining images using anti‐CD31 antibody of the 3D tissue models by (c) nano‐ and (d) micro coating methods.

Chapter 04

Figure 4.1 Site‐specific, amphiphilic nature in cellulose chemical structure, leading to unique surface properties of cellulose films, depending on the tilted angle of anhydroglucose planes when the glucan chains are uniaxially aligned.

Figure 4.2 Schematic diagram of the hydroxymethyl conformations at the C‐6 position, namely, the orientation of the C6‐O6 bond, gauche‐trans (

gt

), trans‐gauche (

tg

) or gauche‐ gauche (

gg

) with a cellobiose unit.

Figure 4.3 Schematic images of NOC: (a) the initial state before stretching; (b) after stretching; the inserted view is the arrangement of cellulose molecules on the NOC surface.

Figure 4.4 High‐resolution TEM image of NOC, the molecular ordering template, together with the schematic cross‐section image along the perpendicular A (= lateral direction) to the stretching direction indicating the tilted arrangement of the glucan chains. The film was negatively stained by uranyl acetate and spans a region of the copper support grid. Note the individual glucan chains that are separated by an average distance of 0.66 nm (arrows). The arrangement of glucan chains is clearly resolved with 0.46 nm in width.

Figure 4.5 AFM images of (a) the NOC template, (b) the p‐NOC template and (c) the non‐ordered cellulose template as a reference for the p‐NOC before the reaction with calcium cations. Insets in the upper right indicate the fast Fourier transformed images. The cross‐section profiles along the perpendicular A–B are shown in the bottom of each figure. In figures (a) and (b), the average distances between two parallel lines were (a) ca. 0.17 µm and (b) ca. 0.14 µm. The double arrows indicate the stretching direction.

Figure 4.6 FE‐SEM images of the cellulose ribbon deposition process. (A and B) are examples of bacteria synthesizing cellulose ribbons on the oriented molecular track of NOC. Figure 4.6A demonstrates the tight association between the molecular track and the cellulose ribbon. In 4.6B, a bacterium shows a flat ribbon immediately behind its site of synthesis. (1–4) are successive images showing the motion of a bacterium on nematic ordered blended film of cellulose and cellulose acetate which has been uniaxially stretched in the same manner for NOC.

Figure 4.7 AFM images before (a) and after (b) incubation of

Gluconacetobacter xylinus

on NOC for 10 hours in SH media.

Figure 4.8 Schematic figures of self‐assembly of cellulose microfibrils secreted from

G. xylinus

in normal fashion (left) and on the tracks of NOC templates (right). The circles indicate cellulose‐synthesizing enzyme subunits linearly arranged across the cell body.

Figure 4.9 Successive images showing the motion using real‐time video analysis of a bacterium as it secretes a cellulose nanofiber on the nematic ordered chitin template. The bacterium is attached, and thereafter synthesizing the fiber on the molecular tracks in the template to show a “wavy” moving pattern.

Figure 4.10 Differential interference light microscopic images of selective attachment of bacteria (inside circles) on surfaces of honeycomb frames under slow rotation during culture.

Figure 4.11 Successive images showing motion of a bacterium as it secretes a cellulose nanofiber, using real‐time video analysis. The bacterium was selectively attached to the honeycomb frame and synthesized its cellulose fiber on the molecular tracks on the honeycomb frame. The interval of each image was set to be 1 min.

Figure 4.12 Morphological changes in the honeycomb frame on film, observed using atomic force microscopy, after culture. The square areas were focused and cross‐sectioned, and show the 3D images before and after culture. Twenty layers of bacterial cellulose nanofibers were accumulated in 10 h (one layer = 3.5 nm).

Figure 4.13 SEM images of the surface of p‐NOC templates after reactions with calcium cations for (a) 0.5, (b) 5.0 and (c) 10 min with the fast Fourier transformed images as insets on the right. The morphology of calcium phosphate deposition on the non‐ordered cellulose template after the same reaction for 10 min is also displayed as a reference for the p‐NOC template. In (a)–(c), the line widths of calcium phosphates were 0.23 ± 0.07 µm, 0.19 ± 0.04 µm and 0.30 ± 0.08 µm, respectively. The double arrows indicate the stretching direction.

Figure 4.14 Culturing epidermal cells on NOC scaffolds and a plastic plate.

Figure 4.15 Schematic image of three‐dimensional cultivation on the semipermeable NOC template. Broken arrow indicates possible proliferation direction of epidermal cells and arrows indicate the flowing tide direction of the medium into and through NOC template.

Figure 4.16 Cell proliferation on the provided medium of the NOC templates from the loop bottom to the top through the semipermeable NOC template. Culture times, (a, a’) 3 days and (b, b’) 5 days cultivation, and circle, colony. On NOC, the cells kept covering the surface after 5 days culture, indicating that the former layers could be a scaffold for the next generation coming from the bottom.

Chapter 05

Figure 5.1 Hypothetical adhesion, oxidation and cross‐linking reaction pathways between Dopa and different substrates [12].

Figure 5.2 (a) Scheme of the gecko‐ and mussel‐inspired wet/dry adhesive. (b) Adhesion forces of the adhesive with and without mussel‐mimetic polymer in wet and dry condition. (c) Performance of geckel adhesive during multiple contact cycles in water (red) and air (black) [31].

Figure 5.3 (a) The pH‐dependent stoichiometry of Fe

3+

‐catechol complexes. (b) Schematic of proposed cross‐linking mechanism of byssal thread cuticle. (c) PEG‐Dopa

4

. (d) Physical state and color of PEG‐Dopa

4

gels in different pH [40].

Figure 5.4 (a) pH‐responsive hydrogel based on cPEG and 1,3‐benzenediboronic acid [47]. (b) gel structure in DMF [48].

Figure 5.5 (a) Photos and (b) schematic diagram of the self‐healing polymer [49].

Figure 5.6 Schematic drawing of (a) water‐dispersible nanoparticles coated with multiple‐interaction ligand (MIL) [50]. (b) delivery of diamagnetic particles with magnets by the reversible, light‐controlled formation of a dynamic paramagnetic coating on their surfaces, resulting from the isomerization of azobenzene [58].

Figure 5.7 (a–d) Photos of mussel and adhesive protein mfp‐5. (e) The molecule structure of dopamine. (f) Modification process of PDA coating. (g) The thickness of PDA coating with reaction time. (h) Electroless metallization of PDA‐coated substrates [60].

Figure 5.8 Possible structure models of PDA [61, 62].

Figure 5.9 The schematic illustration of the fabrication of (a) Ag conductive film [78], (b) silica film [85], (c) superhydrophobic particles [58], and (d) functionalized yeast cell [101].

Chapter 06

Figure 6.1 Schematic illustration of the possible polymer chain arrangements in different morphologies of diblock copolymers changing from sphere (A) to cylinder (B) and to lamella (C), as the volume fraction (

f

A

) of the hydrophobic block (orange) increases. The dashed curve in each morphology represents a part of the interface between hydrophobic and hydrophilic domains.

Figure 6.2 Cryo‐TEM images of PLA

x

‐

b

‐PEG

44

micelles in aqueous solution. The concentrations of initial PLA

x

‐

b

‐PEG

44

THF solutions were 0.45 wt%. (A) PLA

56

‐

b

‐PEG

44

, (B) PLA

134

‐

b

‐PEG

44

, and (C) PLA

212

‐

b

‐PEG

44

. The scale bars in the images present 200 nm.

Figure 6.3 PLA‐

b

‐PEG micellar system for sustained intracellular drug delivery regulated by stereo complexation and host‐guest interactions.

Figure 6.4 (a) Schematic illustration of formation and triggered drug release from DOX‐loaded PEG‐BM/CDPLLA supramolecular micelles in response to the intracellular microenvironment; (b)

In vitro

DOX release profiles of DOX‐loaded micelles in PBS at 37 °C and different pH values: (a,c,e) DOX‐loaded PEG‐

b

‐ PLLA and (b,d,f) DOX‐loaded PEG‐BM/CD‐PLLA at pH (a,b) 7.4, (c,d) 6.5, and (e,f) 5.5; (c)

In vivo

antitumor efficacies after tail‐vein injection of PBS (control), free DOX, DOX‐loaded PEG‐b‐PLLA, and DOX‐loaded PEG‐BM/CD‐PLLA into male BALB/c nude mice bearing HepG2 xenografts.

Figure 6.5 Hydrogel formation from PLA‐PEG copolymers (a) driven by hydrophobic interactions or (b) stereocomplex driven. Micelles containing either PLLA or PDLA cores are designated as orange or green, respectively. However, micelles containing both PLLA and PDLA within a single core are designated by the color brown to emphasize the co‐crystallization and formation of a new stereo‐complexed crystal.

Figure 6.6 Gel‐sol transition curves: (a) PEO‐PLLA diblock copolymers with Mr values as follows: diamonds, 5000–720; circles, 5000–1000; triangles, 5000–1730; squares, 5000–1960; (b) PEO‐PLLA‐PEO triblock copolymers with Mr values as follows: filled circles, 5000–2040–5000; filled triangles, 5000–3000–5000; filled squares, 5000–5000–5000. The gel–sol transition temperature was determined as follows.

Chapter 07

Figure 7.1 Electroconductive bioscaffold for 2D and 3D cell cultures. Following stimulation with electrical signals, vertebrate cells usually respond to EF by promoting cell alignment, division, migration, and growth parallel to the EF lines. An electroconductive bioscaffold could be used as a conductive, biomimetic scaffold for the regeneration of skin, nerves, and skeletal muscles.

Figure 7.2 Diversified electroconductive bioscaffolds based on conductive polymers.

Figure 7.3 Schematic illustration of the procedure for the fabrication of a groove ridge topography within the GelMA‐CNT hybrid gel (0.3 mg/mL CNTs) with (+ES) or without (−ES) the electrical stimulation at day 10 of the culture; (b)–(c) Immunostaining of cell nuclei/myosin heavy chain (b), and cell nuclei/F‐actin (c). Scale bar: (A) 30 µm, (B) 50 µm.

Chapter 08

Figure 8.1 Sketch of the structure of a rice grain.

Figure 8.2 The chemical structure of starch.

Figure 8.3 Starch structural levels. Level 1 is the individual bonds in a single chain: (1⟶4)‐α linkages (see Figure 8.2) between anhydroglucose units, with (1⟶6)‐α branch points. Level 2 is branched molecules. Level 3 comprises clusters of crystalline helices made of shorter amylopectin chains and of portions of longer amylopectin chains which span more than one lamella. Level 4 comprises alternating amorphous and crystalline lamellae, with clusters of shorter amylopectin chains largely confined to a single crystalline lamella, while the amorphous region largely contains amylose molecules, longer amylopectin chains (which span two or more lamellae), branch points and “dangling” ends of amylopectin chains. Level 5 is the granules, made up of growth rings of alternating lamellae.

Figure 8.4 Molecular structure of mannans: Linear mannan; carob galactomannan (redrawn after [9, 10]); konjac glucomannan (redrawn after [12]); and softwood galactoglucomannan. (redrawn after [7, 13]).

Figure 8.5 Mass profiling of the substitution pattern of tamarind XyG by enzymatic depolymerization using a XyG‐specific endo‐glucanase (XGase) and MALDI‐MS analysis. The substitutions in the XyG backbone are named following the nomenclature initially proposed by Fry et al. [16].

Figure 8.6 Molecular structure of xylans: Hardwood glucuronoxylan (redrawn after [27]); softwood arabinoglucuronoxylan (redrawn after [29]); cereal endosperm arabinoxylans and bran arabinoglucuronoxylan (redrawn after [33, 34]).

Figure 8.7 Effect of substitution pattern and molecular weight on the hydrodynamic properties and conformation in solution of cereal arabinoxylans and β‐glucans.

Figure 8.8 Use of selected carbohydrate‐active enzymes (CAZymes) for structural modification of starch and seed polysaccharides (redrawn after [140]).

Figure 8.9 Chemical modification methods for starch and seed storage polysaccharides.

Chapter 09

Figure 9.1 Chemical structures of 1. cellulose, 2. amylose, 3. pullulan, 4. cellulose tris(phenylcarbamayte) (CTPC), and 5. amylose tris(phenylcarbamayte) (ATPC).

Figure 9.2 Chemical structures of 6. cellulose tris(3,5‐dimethylphenylcarbamayte) (CDMPC) and 7. amylose tris(3,5‐dimethylphenylcarbamayte) (ADMPC).

Figure 9.3 Universal plots of

λ

〈

S

2

〉

z

1/2

vs the Kuhn segment number

n

K

(≡

λL

) for amylose tris(phenylcarbamate) (ATPC) in 4‐methyl‐2‐pentanone at 25 °C [51] (unfilled circles), in 1,4‐dioxane at 25 °C [50] (filled circles), in 2‐butanone at 25 °C [52] (unfilled triangles), in ethyl acetate at 33 °C [51] (filled triangles), in 2‐ethoxyethanol at 25 °C [50] (unfilled squares), and in methyl acetate at 25 °C [51] (filled squares).

Figure 9.4 Holtzer plots [

qP

(

q

) vs

q

] for ATPC50K (

M

w

 = 5.5 × 10

4

) and ATPC20K (

M

w

 = 1.9 × 10

4

) in 1,4‐dioxane at 25 °C [50]. Solid curves, theoretical values for the unperturbed wormlike cylinder. Dashed curves, theoretical values for the rigid cylinder limit (

λ

−1

 = ∞).

Figure 9.5 Infrared absorption spectra (molar extinction coefficient

ε

vs wavenumber) for an ATPC sample (

M

w

 = 2.8 × 10

5

) in 1,4‐dioxane, 2‐ethoxyethanol, and their mixtures of which volume fraction of 2‐ethoxyethanol is shown as

x

[50].

Figure 9.6 Chemical structures of 8. amylose‐2‐acetyl‐3,6‐bis(phenylcarbamate) (AAPC), 9. curdlan, and 10. curdlan tris(phenylcarbamate) (CdTPC).

Figure 9.7 Infrared absorption spectra for CdTPC (

M

w

 = 1.5 × 10

6

) in THF, ATPC (

M

w

 = 3.3 × 10

6

) in 1,4‐dioxane, and CTPC (

M

w

 = 2.6 × 10

6

) in THF [66].

Figure 9.8 Chemical structures of 11. amylose tris(ethylcarbamate) (ATEC), 12. amylose tris(

n

‐butylcarbamate) (ATBC), and 13. amylose tris(

n

‐hexylcarbamate) (ATHC).

Figure 9.9 Infrared absorption spectra for ATBC (

M

w

 = 4.6 × 10

5

) in methanol (MeOH), 2‐propanol (2PrOH), 1‐propanol (1PrOH), 2‐ethoxyethanol (2EE), 2‐butanol (2BuOH), and THF [73].

Figure 9.10 Dependence of − [

α

]

280

(filled symbols) and

f

hyd

(unfilled symbols) on the volume fraction of methanol (

ϕ

m

) for ATBC900K (

M

w

 = 8.9 × 10

5

, circles), ATBC460K (

M

w

 = 4.6 × 10

5

, triangles), ATBC55K (

M

w

 = 5.5 × 10

4

, squares), and ATBC17K (

M

w

 = 1.7 × 10

4

, inverted triangles) [72].

Figure 9.11 A possible 3D structure of rigid helical ATBC (6‐fold helix and

h

 = 0.25 nm): (a) side view; (b) top view [73].

Figure 9.12 Schematic representation of the two‐state wormlike chain (TSWC) model.

Figure 9.13 Plots of (a)

h

vs

f

hyd

, (b)

λh

vs

f

hyd

, and (c)

h

–1

vs

λ

for ATEC (circles), ATBC (triangles), and ATHC (squares) [71].

Figure 9.14 (a) Schematic representation of the amylosic main chain for

h

 = 0.36, 0.29, and 0.26 nm; (b)

h

and

λ

–1

values for ATEC, ATBC, ATHC, and ATPC in THF or 1,4‐dioxane [71].

Figure 9.15 Isothermal titration calorimetric data for ATBC53K (a,

M

w

 = 5.3 × 10

4

) and ATBC460K (b,

M

w

 = 4.6 × 10

5

) in ethyl lactates at 25 °C. 2 μL solution was dropped at every 2 min. [76].

Figure 9.16 Plots of

h

–1

vs

λ

for ATBC (circles) and ATPC in ketones and esters (filled triangles) and in 1,4‐dioxane and 2‐ethoxyethanol (unfilled triangles) [50, 51, 72, 73, 76].

Figure 9.17 Dependences of (a)

λ

−1

and (b)

h

on the molar volume of the solvent (

v

M

) for ADMPC (unfilled circles) and ATPC (filled circles) in ketones and esters [51, 52].

Figure 9.18 Comparison between experimental and theoretical phase boundary concentrations for ATBC (circles), ATEC (triangles), ATHC (squares), and CTPC (inverse triangles) all in THF at 25 °C. Filled and unfilled symbols denote experimental

c

I

(phase boundary concentration between the isotropic and biphasic region) and

c

A

(phase boundary concentration between the biphasic and anisotropic region), and solid and dashed curves are theoretical

c

I

and

c

A

, respectively [84].

Figure 9.19 Circular dichroism spectra for an ATBC sample (

M

w

 = 2.6 × 10

5

) [84].

Figure 9.20 Phase diagram of ATBC in

L

‐ethyl lactate (triangles),

L

‐ethyl lactate (squares), and in THF (circles) at 25 °C. Unfilled and filled symbols denote

c

A

and

c

I

, respectively [84].

Figure 9.21 (a) Polarized light micrograph of cATBC in THF; (b) schematic representation of the liquid crystal consisting of rigid ring polymers [96].

Figure 9.22 Plots of

λ

〈

S

2

〉

z

1/2

vs

n

K

for linear (gray symbols) and cyclic (black symbols) ATBC in THF at 25 °C (unfilled circles), in 2‐propanol at 35 °C (filled circles), and in methanol at 25 °C (unfilled triangles) and for linear and cyclic ATPC in 1,4‐dioxane at 25 °C (filled triangles) and in 2‐ethoxyethanol at 25 °C (unfilled squares). Solid gray and black curves indicate the theoretical values calculated by Eq. 9.2 for linear wormlike chain and by Eq. 9.3 for cyclic wormlike chain, respectively [50, 72, 73, 95, 96].

Figure 9.23 The Kuhn segment number

n

K

dependence of the reduced second virial coefficient (

A

2

M

L

2

/4

λ

−1

N

A

) for cATBC in 2PrOH (unfilled circles) at the theta temperature (35 °C), where

M

L

is the molar mass per unit contour length and calculated from

h

and the molar mass of the repeat unit [96]. Solid curve, results from Monte Carlo simulation by Ida et al. [97]. Dashed line, theoretical values for rigid ring [98].

Figure 9.24 Molar mass dependence of 〈

S

2

〉

z

1/2

for cATPC (unfilled circles) and linear ATPC (filled circles) (a) in MEA at 25 °C, (b) in EA at 33 °C, and (c) in MIBK at 25 °C. Solid black and gray curves, theoretical curves for cylindrical wormlike ring and linear chains, respectively, calculated with the parameters listed in each figure. Dashed lines, theoretical values for wormlike ring with the parameters for linear ATPC [99].

Figure 9.25 Dependences of

h

(a) and

λ

–1

(b) on

v

M

for cATPC (unfilled circles) and ATPC (filled circles) in ketones and esters [99].

Figure 9.26 The Kuhn segment number

n

K

dependence of the reduced second virial coefficient (

A

2

M

L

2

/4

λ

−1

N

A

) for cATPC in methyl acetate (MEA, unfilled triangles), ethyl acetate (EA, unfilled squares), and 4‐methyl‐2‐pentanone (MIBK, inverted triangles) at the corresponding Θ temperatures along with those for cATBC in 2‐propanol (unfilled circles) at the Θ temperature (35 °C) [99]. Solid curve, results from Monte Carlo simulation by Ida et al. [97].

Chapter 10

Figure 10.1 Schematic illustration of

Bombyx mori

’s silk gland and silk structure.

Figure 10.2 Physical properties of sericin (SS) upon ethanol treatment [6].

Figure 10.3 Properties of silk sericin (SS) and its biomedical applications. (a) SS reduces oxidative stress caused by free radicals and prevents food browning; (b) SS prevents UV‐induced apoptosis and sunburn‐related effects on skin; (c) SS is used to develop biomimetic biomaterials and stem cell‐based therapeutics; (d) SS possess attractive pharmaceutical effects and is used for targeted drug and cell delivery.

Figure 10.4 Applications of silk fibroin (SF) in tissue engineering and regenerative medicine.

Chapter 11

Figure 11.1 The mechanism of transition metal‐initiated NCA polymerization. (a) oxidative addition of NCAs to zero valent cobalt and nickel complexes; (b) metallacycle ring contraction mediated by NCA addition; (c) chain propagation in transition metal‐mediated NCA polymerization.

Figure 11.2 Organosilicon amines‐mediated NCA polymerization.

Figure 11.3 Synthesis of PEI‐PLL copolymers.

Figure 11.4 (a) Schematic illustration of the elongated hydrophobic side‐chains leading to the formation of polypeptide helixes with unprecedented stability. (b) Structure of the polypeptides with 11 and 17 σ‐bonds of side chain charge‐backbone distances.

Figure 11.5 (a) Synthetic routes to poly(L‐EGxGlu) homopolypeptides’ (b) plots of transmittance as a function of temperature for aqueous solutions (2 mg mL

−1

) of poly(L‐EG

2

Glu) and poly(L‐EG

3

Glu). Solid line: heating; dashed line: cooling. (c) LCST of poly(L‐EG

2

Glu‐

co

‐EG

3

Glu) copolypeptides as a function of sample composition.

Figure 11.6 Schematic showing structure, redox properties, and proposed self‐assembly of M

O

65

(L

0

.5

/F

0

.5

)

20

copolypeptides into vesicles.

Figure 11.7 (a) Supramolecular polymerization of polypeptide‐grafted comb‐like polymers into tubular superstructures; (b) nucleation‐controlled supramolecular polymerization of polypeptide‐grafted gold nanoparticles.

Figure 11.8 Illustration of the mechanism of formation for PLys‐silica complex with a 2D square structure.

Figure 11.9 Bright‐field optical micrographs showing the liquid coacervates or solid precipitates resulting from the stoichiometric electrostatic complexation of L, D, or racemic (D,L) poly(lysine) (K) with L, D or racemic (D,L) poly(glutamic acid) (E) at a total residue concentration of 6 and 100 mM NaCl. Complexes are formed from (a) pLK + pLE, (b) pDK + pLE, (c) p(D,L)K + pLE, (d) pLK + pDE, (e) pDK + pDE, (f) p(D,L)K + pDE, (g) pLK + p(D,L)E, (h) pDK + p(D,L)E, (i) p(D,L)K + p(D,L)E. Liquid coacervate droplets are only observed during complexation involving a racemic polymer. Scale bars, 25 µm.

Figure 11.10 Intracellular kinetics of PPABLG/PAOBLG‐MPA/siRNA and PPABDLG/PAOBLG‐MPA/siRNA HNPs, highlighting the helicity‐dependent membrane disruption capabilities of PPABLG toward effective cellular internalization as well as endosomal escape.

Chapter 12

Figure 12.1 Gradient structures in biological systems. (a) The native ACL‐bone interface (left) and Fourier Transform Infrared Spectroscopic (FTIR) image (right). (b) Composite image of a fiber cap cross‐section. The 2D UV absorbance scans illustrate the degree of cell wall lignification. (red‐to‐blue color represents high to low levels of lignin).

Figure 12.2 Examples of gradient hydrogels fabricated using the microfluidic methods. (a) Schematic of the microchannels used in the experiment. The monomer solution 1 contained rhodamine to make the fluorescent images illustrating the obvious gradient concentration at the inlet and outlet. (b) Light micrographs of HUVECs attached to the surface of PEG hydrogel without RGDS (A), with RGDS (B) and with a gradient of tethered RGDS (D). And quantification of HUVECs attachment on corresponding hydrogels (c) Schematic of gradient generation in a microfluidic channel. (d) The gradient PEG‐DA hydrogels fabricated in (c) procedure. scale bar, 50 µm.

Figure 12.3 Molecular diffusion method to form gradient hydrogels. (a) Schematic representation of spatial localization of PLG microspheres within PEG hydrogels. The bright field image show distinct PLG microspheres‐loaded depots, localized at left side of the PEG hydrogel (scale bar, 1 mm). (b) Schematic of preparation process of hyaluronic acid (HA) hydrogels with the gradient of chemically immobilized bisphosphonate (BP) groups involving two consecutive chemo‐selective reactions.

Figure 12.4 Gradient stiffness in hydrogels. (a) Schematic diagrams showing the stiffness gradient PVA hydrogel generated by gradual freezing‐thawing method. The red dots represent cross‐linking points in PVA hydrogel networks. (b) Immunofluorescent staining (Phalloidin) images of hBMSCs cultured on the stiffness gradient PVA hydrogel after 28 days. (c) Schematic diagrams of the process to prepare an HA‐based hydrogel with orthogonal gradients. (d) The orthogonal gradients of both stiffness and ligand density. (e) U373‐MG human GBM cells remained rounded and rarely exhibited lamellipodia in soft regions with low fibronectin. In contrast, cells spread extensively and developed large lamillipodia in high stiffness regions with high fibronectin.

Figure 12.5 Grafting density gradient polymer brushes. (a) Schematic of the vapor diffusion method for preparation of grafting density gradient PAA brushes. (i) The OTS evaporated and diffused in the vapor phase to generate a concentration gradient along the substrate, then reacted with the ‐OH functional group bonded onto the substrate to form the gradient OTS SAM layer. (ii) The regions of the silica substrate which were not covered with the inactive OTS were backfilled with ATRP initiators. (iii)–(iv) PtBA brushes were grafted onto the substrates by the SI‐ATRP technique, and subsequently converted into PAA brushes. (b) Schematic of the temperature gradient method for preparation of PS brushes with grafting density gradient. (c) Schematic of the plasma copolymerization method for preparation of PHEMA brushes with grafting density gradient. (d) Schematic of PHEMA brushes with gradient grafting density. The conformation of brushes changed from “mushroom” to “brush” regime. The adsorptive capacity of bovine fibronectin was inversely proportional to the grafting density of PHMEA brushes. (e) Contour plots of dry thicknesses of PHEMA brushes (left) and dry fibronectin thickness (right) in a MW‐σ orthogonal gradient. The fluorescence images of MC3T3‐E1 cells which are taken along the diagonal (marked with the numbers in the right contour plot) of the orthogonal substrate.

Figure 12.6 Molecular weight gradient polymer brushes. (a) Schematic of the apparatus for preparation of PMMA brushes with molecular weight gradient by gradually draining the reaction solutions. (b) The thickness of dry PMMA brushes vs position of the substrate. The inset is the thickness of dry PMMA brushes vs the polymerization time. (c) Schematic of the microchannel confined method for preparation of PHEMA brushes with molecular weight gradient. (d) Schematic of movable photomask method for preparation of PHEMA brushes with molecular weight gradient by photo‐induced iniferter polymerization. (A), (B), and (C) show the procedure of gradient PHEMA brushes growth via the movable photomask.

Figure 12.7 Gradient polymer brushes by surface‐initiated electrochemically mediated ATRP (SI‐eATRP) method. (a) Mechanism of SI‐eATRP at a gold working electrode. (b) Schematic of SI‐eATRP for non‐conducting substrate through catalyst diffusion. (c) Schematic of the electrochemical apparatus used for SI‐eATRP. The initiator‐modified substrate with 10 mm length was placed against the BPE, leaving a gap with D µm. (d) The thickness of gradient PNIPAM brushes as a function of the position. (e) Schematic of the apparatus used for pattern polymer brushes.

Figure 12.8 Gradient polymer brushes by light‐mediated living radical polymerization. (a) Mechanism of the light‐mediated living radical polymerization using a photoredox catalyst

fac

‐[Ir(ppy)

3

] (ppy = 2‐pyridylphenyl) in (b). Schematic of the light‐mediated CRP method for preparation of (c) patterned PMMA brushes by using a rectangular patterned photomask, (d) gradient PMMA brushes by using a photomask with a linear gradient neutral density filter and (e) gradient comb brush architecture. (f) Optical micrograph and (g) three‐dimensional AFM image of nanoscale‐inclined plane of gradient PMMA brushes. (h) Brushes’ thickness as a function of position of gradient PMMA brushes.

Figure 12.9 Gradient polymer brushes by surface‐initiated sacrificial anode ATRP (SI‐saATRP) and Cu(0)‐mediated controlled radical polymerization (SI‐CuCRP). (a) Experimental se‐up and digital photo of the sandwiched architecture of SI‐saATRP. A brown‐red color appeared at the surface of Zn sheet. (b) The proposed mechanism of SI‐saATRP. The “sacrificial anode” (Zn) reduced Cu

II

/L to Cu

I

/L, which diffuses to the surface of the initiator‐modified silicon substrate. (c) Schematic of the preparation for gradient polymer brushes by tilting the Zn sheet in a certain angle (θ = 11.5° or 18.5°). Digital photos and 3D optical images of gradient PSPMA brushes in different tilting angles. (d) A ridge‐like polymer brush structure was obtained by placing a Zn wire above the substrate. Plots of the brush thickness as a function of position with corresponding digital photo of ridge‐like PSPMA brushes. (e) Schematic and experimental set‐up of SI‐CuCRP. The reaction solution consisted of monomer and ligand without copper halide catalysts. (f) Plots of the brush thickness vs position of gradient poly(methacryloyloxy)ethyl trimethylammonium chloride) (PMETAC) brushes. Inset is a digital photo of the substrate with macroscopic gradient PMETAC brushes.

Figure 12.10 Gradient cell adhesion. (a) Schematic of the PEG concentration gradient prepared by click chemistry. 3T3 cells were cultured on this surface to show cell density gradient. (b) Schematic of the countercurrent gradients of PEG and RGD concentration. (c) Quantification of cell attachment on the substrate in (a) and (b). (d) Schematic of the sedimentation method for fabricating the cell density gradient. (e) Fluorescence micrographs showing cell density gradients generated on glass substrate at different tilt angles. (f) Schematic of the rapid generation of a cell gradient. Top view and side view SEM images showing the increased surface roughness and thickness of the nanodendritic gradient, respectively. The fluorescence micrographs showing the gradual increasing cell density along the direction with gradient nanodendrites. (g) The influence of chemical gradients on nanodendrites for generation of cell gradients.

Figure 12.11 Gradient polymeric structures‐regulated cell migration. (a) Schematic of the process flow for generation of dynamic ligand surface gradients. (b) Fluorescent micrographs show cells at

μ

CP regions migrated toward unveiled cell‐adhesive photo‐patterned regions. (c) Time‐lapse micrographs showing the different migration rates of patterned cells up or down the RGD peptide gradients. (d) Different gradients of radially aligned fibers with varied collection time of electrospinning. Shallow (2 min), steep (8 min) and moderate (15 min) continuous gradient were generated with the increasing collection time. (e) Fluorescence micrographs showing the migration of NSCs on the CBD‐SDF1α immobilized radially aligned fibers. (f) NSCs distribution on radially aligned fibers immobilized with CBD‐SDF1α. NAT‐SDF1α, PBS, and randomly oriented fibers immobilized with CBD‐SDF1α were as the control groups. The dashed lines in all images indicate the border of NSCs seeding. (g) Schematic of the countercurrent density gradients of PHEMA brushes and YIGSR for selective guidance of ECs migration. (h) Migration traces of ECs and SMCs on countercurrent density gradient of PHEMA/YIGSR, single YIGSR gradient and PHEMA gradient. (i) Optical images of ECs and SMCs sheets cultured on PHEMA/YIGSR, YIGSR and PHEMA density gradient surfaces, respectively.

Chapter 13

Figure 13.1 Biomimetic fiber‐reinforced composites. (a) The linear (unbranched) ‘Technical Plant Stem’. (b) Carbon fiber‐reinforced polymer (CFRP) branching developed by the ITV Denkendorf that can be filled with concrete (Cc).

Figure 13.2 Technical and natural branchings. (a) Roof‐bearing truss structure as example of technical implementation. (b)

Semiarundinaria

sp. as example of a biological concept generator, longitudinal section of stem‐branch attachment. (c) Detail of longitudinal section in (b). Scale bar: 10mm.

Figure 13.3 Stress distribution (first principal stress) in a simulation of a loaded model of a threefold branching. The thick multiple arrows indicate the direction and position of loading. The loose contours indicate the initial form of the branching. Tensile stresses are shown exemplarily by the double‐headed arrows, and the compressive stresses by the ellipsis.

Figure 13.4 In case of the pomelo (

Citrus maxima

) at least seven hierarchical levels can be distinguished that range from the cm scale (whole fruit) to the µm scale (multilayered cell wall). The pomelo fruit – hierarchy level 1 – in this figure has a diameter of about 20 cm. The second hierarchy level distinguishes between the fruit pulp and the fruit peel. The latter is subdivided into flavedo, albedo, and vascular bundles (hierarchy level 3). The fourth hierarchical level is represented by the density gradient within the albedo, which – on the fifth hierarchy level – appears as an open‐pored, foam‐like structure consisting of struts (living biological cells) and void space (intercellulars). The afore‐mentioned struts consist of the cell wall and the cell lumen (hierarchy level 6). The smallest‐scaled hierarchical level considered in this approach is the cell wall itself, consisting of a fiber matrix compound.

Figure 13.5 Typical idealized 2D‐unit cells. (a) Open pored foam. (b) Re‐entrant 2D‐unit cell used to represent the mechanical principles on which auxetic foams are based.

Figure 13.6 Latex of the Weeping Fig (

Ficus benjamina

) at various times after injury. When the cortex is freshly injured, the milky white latex oozes out quickly (0 min). As the latex coagulates, it turns more and more transparent (10–30 min).

Figure 13.7 Restoration of tensile strength due to the coagulation of plant latex. When the bark of the Weeping Fig is injured (0 min), its tensile strength is reduced compared to an uninjured bark sample by more than 50%. A significant increase in tensile strength of the bark can be observed until only 30 min after injury due to the coagulation of the plant latex. A further increase in tensile strength cannot be observed after this time period (at least until several hours to few days after injury, when other mechanisms such as cell growth and cell proliferation cause an additional self‐healing effect).

Figure 13.8 Top‐down process in biomimetics, represented by the successful transfer of the self‐sealing system of the Dutchmen’s pipe to a biomimetic self‐sealing foam coating for membranes of technical pneumatic structures. (1) After the question arises, whether biology might offer a solution for a technical challenge: can a technical pneumatic structure be equipped with an efficient light‐weight self‐sealing system?, a screening for suitable biological role models follows. (2) Self‐sealing of internal fissures in the Dutchmen’s pipe. (3) After a detailed investigation of the functional principle of the biological role model (rapid sealing of internal damage by swelling of pressurized, non‐lignified parenchymatous cortex cells into the fissure), an abstraction of this operating principle is conducted. (4) The abstraction from pressurized, non‐lignified biological cells to similar technical structures can be described by mathematical formulas expressing the interaction of material properties and geometric parameters. This functional principle can then be applied to the technical material. (5) Closed cell polyurethane foam coating under compression on the inside of the membrane, resulting in an enhanced product. (6) Self‐sealing technical membrane which can be incorporated into a multitude of pneumatic systems.

Figure 13.9 (a) Bird of paradise flower with perch which is bent down after landing of pollinating birds and served as concept generator for development of the biomimetic façade shading system Flectofin

®

. (b) Perch bent down by hand; due to torsional bending movement the perch opens and makes the previously protected stamen and ovary accessible for pollinators.

Figure 13.10 Force‐deflection‐curves of 3000 repetitive bending tests with a perch of a bird of paradise flower. The analysis proves that even after the perch was released 3000 times, no evidence of fatigue is visible. With the exception of the first 5–10 bending events, which are slightly higher than the subsequent ones, the force‐deflection curves are very similar.

Figure 13.11 Demonstrator of the biomimetic façade shading system Flectofin

®

in different positions of closure caused by bending of the “backbone” due to a 25 millimeter displacement of hydraulic pistons in the basal support.

Chapter 14

Figure 14.1 Schematic illustration of thermal‐induced anisotropic contraction and expansion of the CLCPs.

Figure 14.2 Schematic illustration of two‐step method to prepare a thermal‐responsive CLCP.

Figure 14.3 (a) Monoacrylate mesogenic monomer A‐6OCB and chial dopant S‐811. (b) Schematics of the director configuration of TNE ribbons. (c) Top view of the L‐ and S‐geometry. Nematic director twists left‐handedly by 90 ° between the top and bottom surfaces with the director at the midplane parallel to the long or short axis of the ribbon. (d) L‐geometry helicoids formed by the narrow twist‐nematic CLCP films: right‐handed at 330 K; almost flat at 353 K; left‐handed at 370 K. (e) Shape of the S‐geometry ribbons: left‐handed at 334 K; right‐handed at 378 K.

Figure 14.4 (a) A scheme of reversible transesterification. (b) The synthesis of xLCE, showing the mesogen and the spacer used at 1:1 ratio, and the triazabicyclodecene catalyst. (c) An xCLCP film bonded with a non‐liquid‐crystal epoxy elastomer could be joined together at a sufficiently high temperature T > T

v

. (d) Reversible cycle of a dome‐shaped molded CLCP actuator becoming completely flat on heating to the isotropic phase.

Figure 14.5 (a) Schematic of chemical structure of the main‐chain CLCP that can be surface‐aligned. (b) Representative photograph of CLCP film with nine +1 topological defects between crossed polarizers. As indicated in the inset schematic, the director orientation varies azimuthally around the defect. (c) Upon heating, nine cones arise from the CLCP film that reversibly flatten upon cooling.

Figure 14.6 (a) The triple co‐flowing geometry of the microfluidic set‐up allows the fabrication of nematic shell particles filled with a glycerol core. Dispersed in the surrounding continuous‐phase silicone oil, the monomer droplets are polymerized by ultraviolet light. (b) Chemical structures of the reactive mesogen (1) and cross‐linker (2) in the monomer mixture. (c) Schematic drawing of the micromanipulator to penetrate the particle shell with a thin glass capillary. (d) When heated to the isotropic phase at 130 °C, the phase transition results in a deformation of the CLCP shell. The force exerted by the actuating shell pushes the liquid glycerol core into the capillary. Once the particle resumes its former shape in the nematic phase at 90 °C, the glycerol sinks to its original level (scale bar, 100 mm).

Figure 14.7 (a) Schematic diagram of the CLCP iris with integrated polyimide‐based platinum heaters. (b) Chemical structures of the monomer and cross‐linker. (c) Actuation of the iris at different voltages showing the controllability of discrete contraction states. The standard deviation of contraction (n = 55) is shown by the error bars. The power consumption of the iris in the different contraction states is indicated on the right axis. (d) A series of images showing different tuning states of the radially oriented CLCP iris with integrated heaters.

Figure 14.8 (a) Chemical structures of monomer VC4 and cross‐linker DVSA used to prepare microsized CLCP pillars. (b–d) The shape of an isolated pillar with the diameter of 20 µm changes with temperature under polarizing optical microscope (from left to right, T = 30, 96, and 48 °C). Scale bar: 100 µm. A: Analyzer; P: Polarizer. (e) SEM images of structured surface with hexagonally‐arranged micro‐pillars. (f) Hydrophobicity changes for the responsive surface with micro‐pillars at different temperatures. Red line (structured surface); blue line (flat surface).

Figure 14.9 (a) Schematic illustration of the mechanism for the change of inverse opaline film based on CLCPs with different temperatures: (I) CLCP‐based inverse opaline film in the nematic state of T < T

NI

(voltage off); (II) CLCP‐based inverse opaline film in the isotropic state of T > T

NI

(voltage on). (b) SEM images of the inverse opaline polymer film. The bars in the images stand for 200 nm. (c) The photonic band‐gap shifts and the induced structural colour changes reversible of the CLCP inverse opaline film driven by voltage.

Figure 14.10 (a) Reflection spectra‐reversible shifts and structural color‐reversible changes of the inverse opal film as a function of temperature. The size of the CLCP inverse opal film in the image is about 12 × 10 mm

2

. (b) Schematic for the mechanism of the lattice space reversible change of CLCP inverse opal films induced by temperature variation.

Figure 14.11 (a) Preparation of a patterned alignment cell. (b) Deformation of CLCP films with a complex order of mesogens: an azimuthal polymer film and a radial polymer film. The arrows along the radius and the azimuth indicate the direction of deformation.

Figure 14.12 (a) Concept of artificial heliotropism. 3D schematic of the system (left) and 3D schematic of the heliotropic behavior (right). The actuator(s) facing the sun contracts, tilting the solar cell toward the sunlight. (b) Before irradiation. Intensity of the white light source is 100 mW/cm

2

. (c) 30 s after irradiation was on. The solar cells tilt toward the sunlight leading to the increased photocurrent output.

Figure 14.13 (a) Scheme of an origami structure of four polycarbonate (PC) films that are adhesively connected by three SWNT‐LCP/silicone bilayer hinges. (b) Reversible folding and unfolding of the origami structure in response to continuous‐wave (CW) NIR light (11.0 mW/mm

2

). (c) Reversible closing and opening of a Venus flytrap‐inspired gripper in response to CW NIR light (28.2 mW/mm

2

). The structure has two curved PC films that are connected by three SWNT‐LCP/silicone bilayer hinges. (d) The inchworm walker crawling up the wood substrate at a 50° incline in response to on and off cycles of CW NIR light (28.2 mW/mm

2

).

Figure 14.14 Schematic illustration of photoisomerization of the azobenzene chromophores and photochemical phase transition of the azobenzene‐containing LCPs.

Figure 14.15 (a) Chemical structure of the LC monomer (1) and cross‐linker (2) used for preparation of the polydomain CLCP film. (b) Images of the film bending in different directions in response to linearly polarized UV light with different polarization directions at λ = 366 nm. The film is subsequently flattened again by visible light at λ = 540 nm. Size of the film: 4.5 mm × 3 mm × 7 mm.

Figure 14.16 (a) Preparation of an oriented CLCP/CNT nanocomposite film in four steps: 1) growth of the CNT array; 2) formation and stabilization of the CNT sheet on a glass substrate; 3) preparation of the LC cell; 4) injection of a mixture containing monomers. (b) Bending behavior of a CLCP/CNT composite film upon exposure to UV (365 nm, 100 mW/cm

2

, 50 s) and visible light (530 nm, 35 mW/cm

2

, 140 s).

Figure 14.17 Schematic illustration of the mechanism of CW NIR‐light‐induced deformation of the azotolane CLCP/UCNP composite film, and photographic frames of the composite film bending in response to the NIR light at CW 980 nm and recovery after removing the light source.

Figure 14.18 (a) Schematic illustration of TTA‐UCL emission of PtTPBP (sensitizer) and BDPPA (annihilator) under excitation with 635 nm laser. (b) Red‐light‐induced deformation of the two layer assembly film. The power density of the 635 nm laser: 200 mW/cm

2

. Thickness of upconverting film: 15 µm; thickness of of CLCP: 27 µm. (c) Mechanism for the photoinduced deformation of the as‐prepared assembly films. The top layer represented the upconverting film; the bottom layer represented the CLCP film.

Figure 14.19 (a) The section of assembled prototype. (b) Photo of experimental prototype (1, inlet; 2, press plate; 3, photodeformable material; 4, outlet; 5, pump membrane; 6, pump chamber)

Figure 14.20 Oscillation of a CLCP cantilever induced by an argon ion laser. (a) Experimental set‐up. (b) The photographs of the azo‐CLCP cantilever oscillation “on” and “off.” Size of the film: 2.7 mm × 0.8 mm × 50 mm.

Figure 14.21 Three‐dimensional motions of photomobile materials composed of bilayer structures of azobenzene CLCP and polyethylene layers. (a) Rotation of a light‐driven plastic motor induced by simultaneous irradiation of UV and visible light. Size of the film: 36 mm × 5.5 mm. Thickness of the layers: PE, 50 mm; CLCP, 18 mm. (b) Series of photographs showing time profiles of the photoinduced inchworm walk of the CLCP‐laminated film by alternate irradiation with UV (366 nm, 240mW/cm

2

) and visible light (>540 nm, 120mW/cm

2

) at room temperature. The film moved on the plate with 1 cm × 1 cm grid.

Figure 14.22 Schematic illustrations of the states of the microrobot during the process of manipulating the object. Photographs showing the microrobot picking, lifting, moving, and placing the object to a nearby container by turning on and off the light (470 nm, 30 mW/cm

2

). Length of the match in the pictures: 30 mm. Thickness of PE and CLCP films: 12 mm. Object weight: 10 mg.

Figure 14.23 (a) An example of paramecia (scale bar 20 µm). (b) A paramecium uses the beating motion of the cilia, characterized by different forward and backward strokes, for self‐propulsion. (c) Side view of the actuation of polymer cilia with ultraviolet light (1 W/cm

2

) in water. (d) Response of a 10‐µm‐thick, 3‐mm‐wide, 10‐mm‐long polymer sample oriented in the splay‐bend molecular organization through the thickness of the film to different colours of light (scale bar 5 mm). (e) Schematic illustration of an asymmetric motion of artificial cilia produced by controlling the wavelength of light. The sample is built with polymer‐containing azobenzene 1 at the top (absorbing at < 390 nm) and polymer‐containing azobenzene 2 at the bottom (absorbing in the 455–550 nm range).

Figure 14.24 (a) SEM image of a microwalker lying upside down. Scale bar: 10 µm. (b) Side view of the microwalker with 500 nm leg tip shown in the inset. Scale bar: 10 µm. (c) Top row shows the initial state of microwalkers on different surfaces. Bottom row shows the microwalker randomly walking on the polyimidecoated glass surface, rotating with one leg stuck onto the polyimide‐coated surface, walking with self‐reorientation on the clean glass surface, walking in the direction determined by the grating groove pattern (vertical). Insets of the top row show the schematics of the surface.

Figure 14.25 (a) The CLCP ribbons display a variety of shapes that depend on the direction in which they are cut. (Ribbon A: flat; Ribbon B: left‐handed; Ribbon C: open ring; Ribbon D: right‐handed). Ribbon B and Ribbon D irradiated for two minutes with UV light display winding and unwinding. (b) A coiled tendril of the wild cucumber plant. (c) A CLCP spring that displays a cucumber tendril‐like shape, composed of two opposite‐handed helices. On irradiation, the right‐handed helix unwinds and the left‐handed helix winds.

Figure 14.26 (a) Schematic representation of the dynamics of the fingerprints. (b) Polarized optical microscopy images of a fingerprint texture as observed between crossed polarizers. Bright regions correspond to planar and black areas to homeotropic orientation. (c) Confocal microscopic 3D image of the initial flat state of fingerprints. (d) 3D image of surface topographies under UV exposure. (e–h) Upon UV illumination, snapshots of a gripper that releases an object by changing friction of fingerprints.

Figure 14.27 (a) Schematic illustration of the PDMS‐soft‐template‐based secondary replication process to prepare the micro‐arrayed azobenzene LCP film. (b) Optical photo of MA‐F0208 with two patterned areas named D15 and D5. Large‐area optical microscopic image and local amplified image (inset) of MA‐F0208‐D15. The patterns of D15 and D5 are all square‐arrayed square posts with the post width of 10 mm. The spacings between two nearest posts for D15 and D5 are 15 mm and 5 mm, respectively. (c) Microscopic profiles of the rolling and pinned 2 mL water droplets.

Figure 14.28 (a) Schematic procedure of the fabrication of microarrayed CLCP films. (b) SEM image of submicropillar arrayed CLCP film. (b, inset) The shape of a water droplet on the CLCP film when it is turned upside down, indicating its high water adhesion. (c) SEM image of the surface of submicrocone arrayed CLCP film.

Figure 14.29 (a) SEM images of the azobenzene CLCP microarray. After irradiation of UV light, the diameter of the pillars increased. (b) Reflection spectra of the azobenzene CLCP microarray under the UV light irradiation (365 nm, 20 mW/cm

2

, 15 min) and the following visible light irradiation (530 nm, 20 mW/cm

2

, 5 min) with the angle of incidence of 60 °.

Figure 14.30 SEM images of the inverse opal film (a) before and (b) after UV light irradiation. (c) The maximum intensity of the reflection peak of the inverse opal film under alternating irradiation with UV and visible light at different times. (d) Reflection spectra of the inverse opal film as a function of temperature.

Chapter 15

Figure 15.1 Schematic presentation of hydrogels based on synthetic polymers (a) and recombinant proteins (b).

Figure 15.2 The unique features of protein (a) structure and (b) synthesis.

Figure 15.3 Chemical cross‐linking of proteins. (a) The schematic of GA‐cross‐linked BSA hydrogel. (b) Artificial proteins EPE consist of terminal cysteine residues (‐SH), elastin‐like endblocks E, and the P midblock domain, and EPE requires covalent cross‐linking with 4‐arm PEG vinyl sulfone to form gels. (c) Spy network hydrogels. Top) Genetic constructs for the four protein precursors (A and B). Construct B contains an internal integrin binding RGD sequence to facilitate cell adhesion and an MMP‐1 cleavage site at the C‐terminus to allow matrix remodeling by encapsulated cells. Bottom) Schematic illustration of the covalently cross‐linked gel formed by mixing protein precursors A and B. (d) Scheme of PPTase‐mediated cross‐linking of PEG hydrogels.

Figure 15.4 Physical cross‐linking of proteins. (a) Proposed physical gelation of monodisperse copolymer. The chains are drawn as disulfide‐linked dimers, joined through their COOH‐terminal cysteine residues. (b) Schematic of the mixing‐induced, two‐component hydrogel. (Top left) Modular association domains assemble via molecular recognition. Two WW domains (CC43 and a Nedd4.3 variant) bind the same proline peptide (PPxY). (Top right) Hydrophilic spacers. (Bottom) Mixing components at constant physiological conditions results in hydrogel formation. (c) Schematic illustration of the hydrogel formed by the biospecific association between PDZ domain in TIP1 protein (CutA‐TIP1) and PDZ domain recognizing peptide (PDZ‐peptide‐PEG). (d) Schematic showing hydrogelation based on protein fragment reconstitution. Top) Formation of reconstituted GL5 from GN and GC. Bottom) Schematic of fragment reconstitution induced two‐component hydrogelation. The two tandem polyproteins, containing either GN or GC fragments, serve as multiple functional precursors for hydrogelation.

Figure 15.5 Strategies for mechanical reinforcement

Figure 15.6 The gel containing the notch was stretched to 17 times its initial length, and the schematics of the hydrogel.

Figure 15.7 Biosynthesis of protein catenanes.

Figure 15.8 (a–c) Schematic of the micellar copolymerization of the UPyHCBA and acrylamide. (a) The structure of the UPyHCBA monomer; (b) SDS micelles loaded with UPyHCBA in an aqueous acrylamide solution; and (c) micellar copolymerization of acrylamide and UPyHCBA loaded in SDS micelles. (d) Image of the hydrogel materials before and after stretching.

Figure 15.9 Schematic diagram of α‐helix ↔ β‐sheet transition during straining.

Chapter 16

Figure 16.1 Schematic representation of dendritic polymers: (a) dendrimer; (b) hyperbranched polymer; and (c) dendrigraft polymer.

Figure 16.2 Divergent

vs

. convergent growth schemes for the synthesis of dendrimers.

Figure 16.3 Divergent PAMAM dendrimer synthesis from an ethylenediamine (EDA) core.

Figure 16.4 Synthesis of a Fréchet‐type benzyl ether dendrimer by the convergent approach.

Figure 16.5 Convergent approach toward triazole dendrimers. (a) CuSO

4

(5 mol %), sodium ascorbate (10 mol %), H

2

O/tBuOH (1:1); (b) 1.5 equiv NaN

3

, CH

3

COCH

3

/H

2

O (4:1).

Figure 16.6 Synthesis of a dendrimer containing 24 galactoside groups.

Figure 16.7 Synthesis of PEGylated dendrimer‐GFLG‐DOX conjugate.

Figure 16.8 Core (a) and non‐core (b) methods for the synthesis of hyperbranched polymers.

Figure 16.9 Hyperbranched polyphenylene synthesis by the single‐monomer polycondensation method.

Figure 16.10 Synthesis of hyperbranched PAMAM (HYPAM).

Figure 16.11 Synthesis of hyperbranched dendritic‐linear polymers by sequential SCV(C)P‐ATRP and ATRP techniques.

Figure 16.12 Synthesis of HPAH–DOX and autophagy inhibitor LY‐loaded HPAH‐DOX micelles.

Figure 16.13 Synthetic route to poly(VBPT‐

co

‐PEGMA)‐S‐S‐MP.

Figure 16.14 General grafting onto scheme for the synthesis of arborescent polymers.

Figure 16.15 Arborescent polystyrene synthesis by grafting onto chloromethylated polystyrene substrates.