Artificial Protein and Peptide Nanofibers: Design, Fabrication, Characterization, and Applications (Woodhead Publishing Series in Biomaterials) [1 ed.] 0081028504, 9780081028506

Table of contents :
Cover
Artificial Protein and
Peptide Nanofibers:
Design, Fabrication, Characterization,
and Applications
Copyright
Dedication
Contributors
Foreword
Section A: Fabrication and characterizations of artificial protein and peptide nanofibers
Supramolecular self-assembly: A facile way to fabricate protein and peptide nanomaterials
Introduction
Protein and peptide assembly mechanism
Protein self-assembly
Peptide self-assembly
Application of protein and peptide nanomaterials
Drug delivery, tumor therapy, and tissue engineering
Biomimetic light-harvesting nanomaterials
Semiconductive materials
Conclusions
Acknowledgment
References
Self-assembly formation of peptide and protein nanofibers on surfaces and at interfaces
Introduction
Self-assembly formation of peptide/protein nanofibers on material surface
Formation of nanofibers on inorganic material surface
Peptide nanofibers
Protein nanofibers
Formation of nanofibers on organic/biological material surfaces
Self-assembly formation of peptide and protein nanofibers at interfaces
Formation of nanofibers at solid-liquid interfaces
Formation of nanofibers at air-liquid interfaces
Formation of nanofibers at liquid-liquid interfaces
Conclusions and outlooks
Acknowledgments
References
Fabrication of amyloid nanofiber matrices by electrospinning
Introduction
Electrospinning
Basic technique
Principle
Parameter optimization
Primary factors affecting fiber morphology
Concentration
Electrical conductivity
Secondary factors affecting fiber morphology
Voltage
Feed rate
Micro- and nanofibers
Microfibers
Nanofibers
Electrospinning proteins
Silk
Collagen
Albumin
Other proteins
Conclusion
References
Novel protein and peptide nanofibrous structures via supramolecular co-assembly
Introduction
Co-assembled peptide superstructures
Co-assembly of short peptides
Co-assembly of amphiphilic peptides
Co-assembly of peptides based on protein motifs
Co-assembled peptide-protein superstructures
Co-assembled protein superstructures
Electrospinning of co-assembled protein superstructures
Extrusion of co-assembled protein superstructures
Self-assembly of co-assembled protein superstructures
Conclusion
References
Characterization techniques of protein and peptide nanofibers: Self-assembly kinetics
Introduction
Kinetic triggering for molecular self-assembly
Characterizations of self-assembly kinetics of nanofibers/nanofibrils
Spectroscopy analysis
Microscopy analysis
X-ray crystallography analysis
Other analytical assays
Summary
Conclusion and outlooks
References
Section B: Enhanced functions of nanofibers by sequence design and modification
Protein synthesis and characterization
Introduction
Types of proteins
Protein structure
Primary structure
Secondary structure
Tertiary structure
Quaternary structure
Applications of protein in medicine
Bioactive/functional peptides
Protein synthesis
Protein biosynthesis
Transcription
Translation
Initiation
Elongation
Termination
Protein folding
Chemical synthesis
Solution phase peptide synthesis
Solid-phase peptide synthesis (SPSS)
Protecting agents
N-terminal protecting groups
C-terminal protecting group
Side chain protecting groups
Scavengers
Amino acid coupling
Peptide cleavage
Enzymatic synthesis
Effect of temperature
Effect of molar ratio
Solvents
Biocatalyst engineering
Synthesis by recombinant DNA technology
Preparation of rDNA
Transformation
Nonbacterial transformation
Phage introduction
Working of rDNA
Characterization of peptides and proteins
Purity analysis
Electrophoresis
Polyacrylamide gel electrophoresis (PAGE)
Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE)
Capillary electrophoresis (CE)
Capillary zone electrophoresis (CZE)
Isoelectric focussing capillary electrophoresis
Micellar electrokinetic capillary chromatography (MECC)
Assay and purification
Reversed-phase chromatography (RPC) of proteins
Principle
Method
Ion exchange chromatography
Structural characterization of proteins/peptides
Circular dichroism
X-ray crystallography
Nuclear magnetic resonance spectroscopy
Multidimensional NMR
Mass spectrophotometry
Ionization
MALDI ionization
Electrospray ionization (ESI)
Conclusions
References
Further reading
Design of functional peptide nanofibers based on amyloid motifs
Introduction
Formation mechanism and secondary structures of functional amyloid nanofibers
β-Sheet-based self-assembled amyloid nanofibers
α-Helix-based self-assembled nanofibers
Bio-applications of function-tailored amyloid nanofibers
Peptide nanofibers for biomineralization
Peptide nanofibers for tissue regeneration
Peptide nanofibers for drug delivery
Peptide nanofibers’ other biofunctions
Conclusions and outlooks
Acknowledgments
References
Design of amphiphilic peptide nanofibers
Introduction
Amphiphilic peptide design
Self-assembled peptide nanofibers
Characterization methods for self-assembling amphiphilic peptides
Conclusions
References
Nanofiber matrices of protein mimetic bioactive peptides for biomedical applications
Introduction
Protein mimetic bioactive peptide nanofibers
Biomedical applications of protein mimetic bioactive peptide nanofibers
Nanocarrier for drug delivery
Bioactive coatings
Antimicrobial property
Tissue regeneration
Stem cell proliferation
Vascular regeneration
Neural regeneration
Antiangiogenic activity
Skin regeneration
Cartilage regeneration
Bone regeneration
Bio-imaging
Bio-sensing
Conclusion and future prospects
References
Synergetic integration of computer-aided design, experimental synthesis, and self-assembly for the rational design of pept ...
Introduction
Simulation techniques for peptide/protein designs
Peptide-specific simulation techniques
Molecular dynamics simulation techniques
Simulation strategies for peptide/protein sequence designs
De novo design
Multiscale modeling-aided peptide/protein design
Experimental methods for artificial peptide/protein synthesis
Chemical synthesis
Solution-phase synthesis
Solid-phase synthesis
Synthetic biology techniques
Fibrillation of artificial peptide/proteins
Conclusions
Acknowledgments
References
Composite nanofiber matrices for biomedical applications
Introduction
Protein-based nanofiber composites and their interactions
Protein-polymer composite
Protein-metal ion composites
Protein-nanoparticles composites
Protein‑carbon materials composites
Protein-small molecules composites
Protein-hydrogel nanofiber composites
Biomedical applications of composite nanofibers
Tissue engineering
Drug delivery
Sensors
Bioimaging
Conclusion/future perspective
References
Further reading
Nanofiber-based hydrogels and aerogels
NFHGs are of great significance for in vitro culture of spheroid tumor models
Application in nerve repair
Application in drug delivery
Application in healing the wound
Application in separating oil/organic liquid and water
Acknowledgments
References
Section C: Related applications of artificial protein and peptide nanofibers
Protein and peptide nanostructures for drug and gene delivery
Introduction
Albumin nanoparticles
Collagen nanoparticles
Gelatin nanoparticles
Elastin nanoparticles
Fibroin nanoparticles
Sericin nanoparticles
Keratin nanoparticles
Zein nanoparticles
Gliadin nanoparticles
Casein nanoparticles
Beta lactoglobulin nanoparticles
Lactoferrin nanoparticles
Legume protein nanoparticles
Soy protein-based nanoparticles
Lysozyme nanoparticles
Protein-modified nanoparticles
Role of cell penetrating peptides in drug and gene delivery
Self-assembled peptide structures
Emerging applications for protein and peptide-based delivery systems
Peptide-drug conjugates
Electrospun drug delivery systems
Concluding remarks
Acknowledgment
References
Protein and peptide nanofiber matrices for the regenerative medicine
Proteins and peptides in tissue engineering and regenerative medicine
Fundamentals of proteins and peptides structures
Interaction of protein with substrate and role in the cell-materials interaction
Biofunctionalization of nanofiber-based scaffolds with proteins
Self-assembling peptides and proteins in bioengineering
Types and structures
α -Helical peptide nanofibers
β -Sheet peptide nanofibers
Collagen-mimetic peptides
β -Hairpin-like peptide nanofibers
Supramolecular self-assembly of peptides
Design of various structural motifs
Self-assembling peptide in peptide-based hydrogels
Self-assembling proteins
Surface modification with peptides-based nanofibers for the tissue engineering applications
Bone tissue regeneration
Cartilage tissue repairs
Cardiovascular tissue regeneration
Nerve tissue regeneration
Naturally occurring and engineered proteins for the tissue regeneration applications
Summary
Acknowledgment
References
Fibrous scaffolds for bone tissue engineering
Introduction
Bone cells and their microenvironment
Design considerations for fibrous scaffolds for bone tissue engineering
Degradation
Mechanical properties
Fiber orientation
Surface chemistry and osteoinductive properties
Methods for fibrous scaffold processing
Centrifugal spinning
Wet spinning
Fiber knitting
Electrospinning
Self-assembly
Fiber reinforced scaffolds
Current findings in bone tissue engineering using nano- and micro-fibers
Synthetic polymers and polymer composites
Natural polymers
Conclusion and future trends using fibrous scaffold approaches
References
Assembled peptides for biomimetic catalysis
Introduction
Enzyme models constructed by peptide assembly
Oxidoreductase mimics
Metal ion coordination
Ferriporphyrin complexes
Nanozyme incorporation
Hydrolase mimics
Histidine as the catalytic group
Glutamic acid/aspartic acid acts as the catalytic group
Metal ion coordination
Aldolase mimics
Key issues for constructing peptide assembly enzyme mimics
Peptide assembly versus protein folding
Structure design: α-helix or β-sheet?
Catalytic groups in the active site
Catalytic microenvironment
Specificity versus diversity
Switchable activity based on a reversible supramolecular structure
Applications in the environment and healthcare
Perspective
Acknowledgment
References
New protein-based smart materials
Introduction
Smart materials based on protein/hybrid self-assembly
“Smart” hydrogel biomaterials
Smart protocell models based on assembly of protein/protein-polymer materials
Biosensors based on protein assembly
Outlook and perspective
Acknowledgments
References
Nanofibers for soft-tissue engineering
Introduction
Soft-tissue injuries
Soft-tissue engineering
Scaffold matrices
Cell sources
Soft-tissue engineering using nanofiber matrices
Nanofiber matrices fabrication
Self-assembly
Phase separation
Electrospinning
Type of materials used in nanofiber matrices fabrication
Natural nanofiber matrices
Synthetic nanofiber matrices
Blended nanofiber matrices
Bioactive nanofiber matrices
Physical incorporation
Chemical tethering
Nanofiber matrices for protein and growth factor delivery
Application of nanofiber matrices in soft-tissue engineering
Skin
Tendon and ligament
Nerve
Muscles
Conclusion
Acknowledgment
References
Index
A
B
C
D
E
F
G
H
I
K
L
M
N
O
P
Q
R
S
T
V
W
X
Z
Back Cover

Citation preview

Artificial Protein and Peptide Nanofibers

Woodhead Publishing Series in Biomaterials

Artificial Protein and Peptide Nanofibers Design, Fabrication, Characterization, and Applications

Edited by

Gang Wei Sangamesh G. Kumbar

An imprint of Elsevier

Woodhead Publishing is an imprint of Elsevier The Officers’ Mess Business Centre, Royston Road, Duxford, CB22 4QH, United Kingdom 50 Hampshire Street, 5th Floor, Cambridge, MA 02139, United States The Boulevard, Langford Lane, Kidlington, OX5 1GB, United Kingdom © 2020 Elsevier Ltd. All rights reserved No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, recording, or any information storage and retrieval system, without permission in writing from the publisher. Details on how to seek permission, further information about the Publisher’s permissions policies and our arrangements with organizations such as the Copyright Clearance Center and the Copyright Licensing Agency, can be found at our website: www.elsevier.com/ permissions . This book and the individual contributions contained in it are protected under copyright by the Publisher (other than as may be noted herein). Notices Knowledge and best practice in this field are constantly changing. As new research and experience broaden our understanding, changes in research methods, professional practices, or medical treatment may become necessary. Practitioners and researchers must always rely on their own experience and knowledge in evaluating and using any information, methods, compounds, or experiments described herein. In using such information or methods they should be mindful of their own safety and the safety of others, including parties for whom they have a professional responsibility. To the fullest extent of the law, neither the Publisher nor the authors, contributors, or editors, assume any liability for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions, or ideas contained in the material herein. Library of Congress Cataloging-in-Publication Data A catalog record for this book is available from the Library of Congress British Library Cataloguing-in-Publication Data A catalogue record for this book is available from the British Library ISBN: 978-0-08-102850-6 ISBN: 978-0-08-102851-3 For information on all Woodhead publications visit our website at https://www.elsevier.com/books-and-journals

Publisher: Matthew Dean Acquisitions Editor: Sabrina Webber Editorial Project Manager: Mariana Kuhl Production Project Manager: Vignesh Tamil Cover Designer: Matthew Limbert Typeset by SPi Global, India

Dedication

Prof. Kumbar dedicates this book to a deceased mentor and friend. Prof. Anandrao R. Kulkarni, M. Pharm., PhD. Sonia College of Pharmacy, Dharwad, India.

Contributors

Sama Abdulmalik  Department of Orthopedic Surgery, University of Connecticut Health Center, Farmington; Department of Biomedical Engineering, University of Connecticut, Storrs, CT, United States Aneela Anwar  Department of Biomedical Engineering, Stevens Institute of Technology, Hoboken, NJ, United States; Department of Basic Sciences and Humanities, University of Engineering and Technology, Lahore, Pakistan Treena Livingston Arinzeh  Department of Biomedical Engineering, New Jersey Institute of Technology, Newark, NJ, United States Michael R. Arul  Department of Orthopaedic Surgery, University of Connecticut Health Center, Farmington, CT, United States Jiana Baker  Department of Orthopedic Surgery, University of Connecticut Health Center, Farmington, CT, United States Rosalie Bordett  Department of Orthopaedic Surgery, University of Connecticut Health Center, Farmington, CT, United States Ramya Dhandapani  Tissue Engineering & Additive Manufacturing (TEAM) Lab, Centre for Nanotechnology & Advanced Biomaterials (CeNTAB), School of Chemical & Biotechnology, SASTRA Deemed University, Thanjavur, Tamil Nadu, India Ibrahim Dulijan  Department of Biomedical Engineering, Stevens Institute of Technology, Hoboken, NJ, United States Nithyadevi Duraisamy Department of Orthopaedic Surgery, University of Connecticut Health Center, Farmington, CT, United States Mustafa O. Guler The Pritzker School of Molecular Engineering, The University of Chicago, Chicago, IL, United States Christian Helbing  Chair of Materials Science, Otto Schott Institute of Materials Research, Friedrich Schiller University, Jena, Germany

xivContributors

Xin Huang MIIT Key Laboratory of Critical Materials Technology for New Energy Conversion and Storage, School of Chemistry and Chemical Engineering, Harbin Institute of Technology, Harbin, China Devina Jaiswal  Department of Biomedical Engineering, Western New England University, Springfield, MA, United States Klaus D. Jandt  Chair of Materials Science, Otto Schott Institute of Materials Research, Friedrich Schiller University, Jena, Germany Shobhit Kadakeri Burlington High School, Burlington, MA, United States Sara Katebifar Department of Biomedical Engineering, University of Connecticut, Storrs; Department of Orthopedic Surgery, University of Connecticut Health Center, Farmington, CT, United States Uma Maheswari Krishnan Centre for Nanotechnology and Advanced Biomaterials; School of Chemical and Biotechnology; School of Arts, Science and Humanities, SASTRA Deemed University, Thanjavur, India Alok Kumar Department of Biomedical Engineering, Stevens Institute of Technology, Hoboken, NJ; Department of Biochemistry & Molecular Biology, University of Maryland School of Medicine, Baltimore, MD, United States Sangamesh G. Kumbar Department of Biomedical Engineering; Department of Materials Science and Engineering, University of Connecticut, Storrs; Department of Orthopedic Surgery, University of Connecticut Health Center, Farmington, CT, United States Apurva Limaye  Department of Biomedical Engineering, New Jersey Institute of Technology, Newark, NJ, United States Shengjie Ling School of Physical Science and Technology, ShanghaiTech University, Shanghai, PR China Yawen Liu  School of Physical Science and Technology, ShanghaiTech University, Shanghai, PR China Amrutha Manigandan Tissue Engineering & Additive Manufacturing (TEAM) Lab, Centre for Nanotechnology & Advanced Biomaterials (CeNTAB), School of Chemical & Biotechnology, SASTRA Deemed University, Thanjavur, Tamil Nadu, India Seyed Mohammad Mir Department of Biomedical Engineering, Stevens Institute of Technology, Hoboken, NJ, United States

Contributorsxv

Jennifer Moy  Department of Biomedical Engineering, New Jersey Institute of Technology, Newark, NJ, United States Hemantkumar Naik  PPD, FSP Department, 320 Research Way, Middleton, WI, United States Jonathan Nip  Department of Orthopedic Surgery, University of Connecticut Health Center, Farmington; Department of Biomedical Engineering, University of Connecticut, Storrs, CT, United States Wei Qi  School of Chemical Engineering and Technology, Tianjin University, The ­Co-Innovation Centre of Chemistry and Chemical Engineering of Tianjin, Tianjin, PR China Jing Ren  School of Physical Science and Technology, ShanghaiTech University, Shanghai, PR China Swetha Rudraiah  Department of Pharmaceutical Sciences, School of Pharmacy, University of Saint Joseph, Hartford; Department of Orthopedic Surgery, University of Connecticut Health Center, Farmington, CT, United States Muthu Parkkavi Sekar  Tissue Engineering & Additive Manufacturing (TEAM) Lab, Centre for Nanotechnology & Advanced Biomaterials (CeNTAB), School of Chemical & Biotechnology, SASTRA Deemed University, Thanjavur, Tamil Nadu, India Swaminathan Sethuraman Tissue Engineering & Additive Manufacturing (TEAM) Lab, Centre for Nanotechnology & Advanced Biomaterials (CeNTAB), School of Chemical & Biotechnology, SASTRA Deemed University, Thanjavur, Tamil Nadu, India Zhiqiang Su State Key Laboratory of Chemical Resource Engineering; Beijing Key Laboratory of Advanced Functional Polymer Composites, Beijing University of Chemical Technology, Beijing, PR China Anuradha Subramanian Tissue Engineering & Additive Manufacturing (TEAM) Lab, Centre for Nanotechnology & Advanced Biomaterials (CeNTAB), School of Chemical & Biotechnology, SASTRA Deemed University, Thanjavur, Tamil Nadu, India Li Sun Cixi Institute of Biomedical Engineering, Chinese Academy of Science (CAS) Key Laboratory of Magnetic Materials and Devices & Zhejiang Engineering Research Center for Biomedical Materials, Ningbo Institute of Materials Technology and Engineering, CAS, Ningbo; University of Chinese Academy of Sciences, Beijing, PR China

xviContributors

Shuwei Sun  State Key Laboratory of Chemical Resource Engineering, Beijing University of Chemical Technology, Beijing, PR China Paulina Szarejko  Department of Orthopedic Surgery, University of Connecticut Health Center, Farmington, CT, United States Lei Wang MIIT Key Laboratory of Critical Materials Technology for New Energy Conversion and Storage, School of Chemistry and Chemical Engineering, Harbin Institute of Technology, Harbin, China Mengfan Wang School of Chemical Engineering and Technology, Tianjin University, The Co-Innovation Centre of Chemistry and Chemical Engineering of Tianjin, Tianjin, PR China Gang Wei College of Chemistry and Chemical Engineering, Qingdao University, Qingdao; Division of Functional Materials and Nanodevices, Ningbo Institute of Materials Technology and Engineering, Chinese Academy of Sciences, Ningbo, PR China; Faculty of Production Engineering, University of Bremen, Bremen, Germany Wenfeng Wei  State Key Laboratory of Chemical Resource Engineering; Beijing Key Laboratory of Advanced Functional Polymer Composites, Beijing University of Chemical Technology, Beijing, China Aiguo Wu  Cixi Institute of Biomedical Engineering, Chinese Academy of Science (CAS) Key Laboratory of Magnetic Materials and Devices & Zhejiang Engineering Research Center for Biomedical Materials, Ningbo Institute of Materials Technology and Engineering, CAS, Ningbo, PR China Xuehai Yan State Key Laboratory of Biochemical Engineering; Center for Mesoscience, Institute of Process Engineering, Chinese Academy of Sciences, Beijing, China Laurie Yousman  Department of Orthopedic Surgery, University of Connecticut Health Center, Farmington, CT, United States Xiaojun Yu Department of Biomedical Engineering, Stevens Institute of Technology, Hoboken, NJ, United States Allen Zennifer  Tissue Engineering & Additive Manufacturing (TEAM) Lab, Centre for Nanotechnology & Advanced Biomaterials (CeNTAB), School of Chemical & Biotechnology, SASTRA Deemed University, Thanjavur, Tamil Nadu, India

Contributorsxvii

Luyang Zhao State Key Laboratory of Biochemical Engineering, Institute of Process Engineering, Chinese Academy of Sciences, Beijing, China Ke Zheng  School of Physical Science and Technology, ShanghaiTech University, Shanghai; Biomass Molecular Engineering Center and Department of Materials Science and Engineering, School of Forestry and Landscape Architecture, Anhui Agricultural University, Hefei, Anhui, PR China

Foreword

I am delighted to write the foreword for Artificial Protein and Peptide Nanofibers edited by Drs. Wei and Kumbar. This book presents a fine summary of the present status of designer peptides and proteins, their structures and properties, and their associated nanofibers for biomedical applications. This book will be extremely useful as a reference source for all those working in the areas of proteins/peptides, polymer chemistry/physics, biomaterials, tissue engineering, drug delivery, and regenerative medicine. Proteins and peptides of both synthetic and natural origin have been widely used as biomaterials for a variety of biomedical applications and have greatly impacted the advancement of modern medicine. These polymeric materials have the ability to self-assemble as fiber matrices that closely mimic the native architecture of the extracellular matrix. Synthetic peptides are designed to mimic biological patterns either to study biological systems or to influence tissue healing responses. They offer controllable physicochemical properties needed for certain biomedical applications including sensors, imaging, drug delivery, targeted therapies, tissue engineering, and electronic devices. Particular emphasis is made here on the procedures to synthesize protein and peptide nanofibers via molecular self-assembly, supramolecular chemistry, electrospinning, templates, and enzymatic synthesis. New proteins/peptides, as well as modifications to existing peptides, are continually being developed and applied to meet new challenges of biomedical applications. Nanofiber matrices in the form of implants, coatings, scaffolds, and drug delivery devices have been created to manipulate cell-material interactions and promote tissue regeneration, as well as for biosensing, imaging, and diagnostics. This intensive literature review on these topics is covered in 18 chapters written by experts in their fields from different parts of the world who present excellent overviews that will be useful for a wide audience. In my opinion, this book will be an ideal resource for those working in materials science, polymer science, chemical engineering, nanotechnology, and biomedicine. I believe this textbook will be a part of homes, libraries, and classrooms throughout the world. Abraham (Avi) J. Domb School of Pharmacy-Faculty of Medicine, The Hebrew University of Jerusalem, Israel

Supramolecular self-assembly: A facile way to fabricate protein and peptide nanomaterials

1

Luyang Zhaoa and Xuehai Yana,b a State Key Laboratory of Biochemical Engineering, Institute of Process Engineering, Chinese Academy of Sciences, Beijing, China, bCenter for Mesoscience, Institute of Process Engineering, Chinese Academy of Sciences, Beijing, China

1.1 Introduction Supramolecular self-assembly, which has its roots in biology, plays a vital role in the construction of natural materials with various fascinating properties and functions. It bridges the starting building blocks and the final complex systems through a b­ ottom-up approach with delicately tuned intermolecular interactions [1–3]. As typical examples, light-harvesting pigments in the chlorosome self-assemble into large supramolecular systems, which efficiently transfers energy to the reaction center by pigment-pigment interactions [4]. In the calicivirus, the contiguous shell is constructed by 180 copies of a protein molecule via finely tuned hydrophobic interactions [5]. These nature-­ inspired or nature-derived self-assembly protocols are highly promising for the future development of functional supramolecular materials. However, natural materials have hierarchical architecture and complicated functions, while artificial materials are comparatively simple in supramolecular structures with confined functionalities [6]. How to construct desired supramolecular structures with abundant functions using simple self-assembly units, thus, remains one of the great challenges. Proteins and peptides are a class of amino acids-composed natural compounds with definite molecular structures and unique functions in biological processes. Proteins possess sophisticated stereo-structures, acting as ideal building blocks for organisms with a broad range of functions. In comparison, holding the abilities of easy manipulation and synthesis, peptides with specifically designed structures show superior self-assembly properties [7–12]. The self-assembling peptides as building blocks enable diverse supramolecular structures such as nanotubes [13, 14], nanofibrils [15, 16], nanobelts [17], and hydrogels [18–20], which show great functionalities for a broad range of applications. In addition to their individual self-assembly, proteins and peptides may induce the assembly of functional species as templates, resulting in not only homogeneous nanostructures (nanotubes, nanofibrils, etc.), but even more complexed hierarchical structures [21–23]. The unique advantage of protein/peptide-­ involved supramolecular assembly lies in their expanded and improved properties that their individual components do not possess [24–26]. Consequently, proteins and peptides are ideal species for constructing functional nanomaterials. Artificial Protein and Peptide Nanofibers. https://doi.org/10.1016/B978-0-08-102850-6.00001-2 © 2020 Elsevier Ltd. All rights reserved.

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Artificial Protein and Peptide Nanofibers

In this chapter, we focus on self-assembling protein- and peptide-modulated nanomaterials in functional applications, mainly including biological materials, biomimetic photo-catalytic systems for energy use, and semiconductive materials. In addition, the supramolecular assembly mechanisms are clarified at first for their substantial importance in bridging the structures and functions between starting building blocks and supramolecular products.

1.2 Protein and peptide assembly mechanism Elucidating the protein and peptide assembly mechanism, especially the complicated noncovalent intermolecular interactions, is highly important because they determine the final assembly structure, morphology, stability, and functionalities [27, 28]. The mechanism complexity lies on two aspects. Firstly, the decisive noncovalent interactions include electrostatic interaction, hydrogen-bonding, π-π interaction, van der Waals interaction, and hydrophobic interaction. They are not independent, but have latent influences with each other and usually show synergistic or competitive effects in the self-assembly [29–31]. Secondly, the self-assembled architectures are not only dependent on the chemical structure of the starting building block, but on various thermodynamic or kinetic conditions during the assembly process, such as environmental temperature, pH, ionic strength, reaction rate, external electric or magnetic field, and the existence of nucleation seed [6, 32–38]. This means one set of starting building blocks can form different nanostructures with distinctive properties [39–42].

1.2.1 Protein self-assembly The inherent complex structure of the protein itself results in its intermolecular interactions encompassing all possible forms as listed above. These weak interactions may distribute on different sites of the protein molecule surface depending on the surficial functional groups. As a consequence, the self-assembly of protein is rather a synergistic effect driven by several noncovalent interactions than merely one dominant interaction. Therefore, it is not suitable to classify the protein self-assembly by the dominant intermolecular interactions. Instead, the various protein self-assembly processes may be distinguished by different experimental strategies, because protein self-assembly is severely dependent on the environmental conditions [43–45]. These experimental strategies include host-guest interaction, ligand-receptor interaction, template-assisted assembly, covalent-conjugation, and coordination-driven assembly. While there have been several review works elucidating the mechanism of protein assembly [43, 46], here only a few typical strategies are represented to show the unique features of protein self-assembly. One of the typical and efficient biological self-assembly strategies is the symmetric fusion-based self-assembly. Inspired by many proteins that can self-assemble into larger highly symmetrical architectures [47, 48], the symmetry-based protein self-­ assembly is available by creating fusions of known protein capsids that already have intrinsic subunit-subunit interaction interfaces [49]. Yeates and coworkers reported

Supramolecular self-assembly

5

that dimeric M1 matrix protein and trimeric bromoperoxidase as two natural oligomeric domains could combine together by genetic fusion in a predetermined orientation with a semirigid helical linker. The protein building blocks self-assembled into a tetrahedral cage containing 12 subunits and holding specific symmetries of 3-fold and 2-fold rotational axis from each oligomeric species [50]. In order to acquire a more delicate self-assembled architecture that could be resolved at atomic level, a series of mutation was further conducted to avoid potential flaw that might cause heterogeneity. Guided by computational design, triple mutant was resulted, and the intermolecular interactions and steric hindrance were modified, leading to a successful crystallization of a homogeneous 12-subunit 16-nm protein cage [51]. In the symmetric fusion-based self-assembly, the driving force of the assembly lies on the strong tendency of each assembly unit to recognize other units, which could be either inherent or externally modified, and an axis that renders constructing a specific geometric pattern. In comparison, chemical assembly strategies of protein self-assembly are featured by the induction of relatively concrete noncovalent interactions as driving forces [52]. The assembly, especially, can be controlled by the strength, directionality, and selectivity of metal-ligand interactions, where the coordination between metal ion and electronegative ligand is stronger than all noncovalent interactions and has explicit directionality. Tezcan and coworkers reported that a 4-helix bundle heme protein, Cytochrome cb562 (cyt cb562), could be constructed to a 16-helix architecture through Zn2 + mediation [53]. The C2-symmetrical interface of cb562 provided a structural point on which two bis-histidine moieties were incorporated to produce the variant (MBPC-1) for selectively binding metal ions. The key role of metal ion coordination for protein self-assembly was revealed by sedimentation velocity and crystallography analysis, where Zn(II) ions stabilized the tetrameric assembly with two pair of V-shaped MBPC-1 wedged into one another (Fig. 1.1).

1.2.2 Peptide self-assembly The driven forces of peptide self-assembly can be explicitly recognized by the contributions of each intermolecular interaction compared with those of protein self-assembly, because peptides possess relatively simple molecular structures, and the intermolecular interactions are solely based on the intermolecular configurations which can be clearly revealed by both experiments and molecular simulations. For example, we have shown that diphenylalanine-mediated coassembly with tetraphenylporphinesulfonate (TPPS) leads to porous multichambered microparticles (Fig. 1.2A) [54], while coassembly of TPPS and dilysine (KK) resulted in nanorods which subsequently aligned at long range to form hierarchical supramolecular fiber bundles (Fig. 1.2B) [55]. The formation of the fiber bundles was probably driven by the hydrogen-bonding interactions of KK, which are oriented forces and lead to the ordered assembly structure. Comparably, FF is hydrophobic molecule and dominantly provides hydrophobic interaction, thereby forming less ordered supramolecular structure. The relatively simple molecular structures and intermolecular interactions of peptides also enable diversified supramolecular structures. Diphenylalanine, a short dipeptide as an extremely versatile self-assembly building block, may afford various

6

Artificial Protein and Peptide Nanofibers

(A)

Mol.1

Mol.3

Mol.4

Mol.2

Asp744 His633

His772

(B)

His732

Fig. 1.1  (A) Supramolecular assembly structure of Zn(II) cooperated cb562, where gray spheres indicate zinc ions. (B) Zoom-in view of the Zn coordination structure. Adapted with permission from E.N. Salgado, J. Faraone-mennella, F.A. Tezcan, J. Am. Chem. Soc. 129 (2007) 13374–13375. Copyright 2007, American Chemical Society.

nanostructures like nanotubes, nanowires, nanovesicles, nanotapes, nanocrystals, nanofibers, and necklaces depending on the environmental thermodynamic condition during its self-assembly process [56]. Sometimes, the resultant different architectures from the same peptide building block are caused by subtle variations, like the existence of trace water in an organic phase [57, 58]. Consequently, it is much more important to elucidate the relationship between supramolecular structure and environmental conditions during peptide self-assembly. In many cases, the formation of different morphological structures from the same self-assembly building block undergoes kinetic or thermodynamic controlling conditions [59]. For example, a simple chemically modified diphenylalanine compound, (((9H-fluoren-9-yl)methoxy)carbonyl)phenylalanylphenylalanine (Fmoc-FF), may transform its self-assembly architectures between α-helix and β-sheet under molecular charge variation (Fig. 1.3) [60]. Neutral Fmoc-FF in acidic solvent tended to form fibrous hydrogels with probably antiparallel β-sheet secondary structure. However, in basic solvent, the C-terminus of Fmoc-FF deprotonated to possess a negative charge, which tended to form helical structure that was totally different from β-sheet. The remarkable structural conversion originated from the dynamic balance between several intermolecular interactions. When deprotonated, repulsive electrostatic interaction

Fig. 1.2  Self-assembly mechanisms of (A) TPPS-FF microspheres and (B) TPPS-KK fiber bundles. Adapted with permission from Q. Zou, L. Zhang, X. Yan, A. Wang, G. Ma, J. Li, H. Möhwald, S. Mann, Angew. Chem. Int. Ed. 53 (2014) 2366–2370; K. Liu, R. Xing, C. Chen, G. Shen, L. Yan, Q. Zou, G. Ma, H. Möhwald, X. Yan, Angew. Chem. Int. Ed. 54 (2015) 500–505. Copyright 2014 and 2015, Wiley-VCH.

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Artificial Protein and Peptide Nanofibers

Fig. 1.3  Schematic illustration of Fmoc-FF self-assembly with charge-induced secondary structures including β-sheet and α-helix. Reproduced with permission from R. Xing, C. Yuan, S. Li, J. Song, J. Li, X. Yan, Angew. Chem. Int. Ed. 57 (2018) 1537–1542. Copyright 2018, Wiley-VCH.

on the negatively charged C-terminus became the major driving force, twisting the ­β-sheet into a helical structure until a new balance is achieved between repulsive electrostatic interaction and attractive hydrogen-bonding and π-π stacking interactions. In a similar while more complicated manner, a peptide amphiphile V3A3K3 conjugated to a hexadecyl chain could self-assemble into either short or long nanofibrils with different secondary structures. The thermodynamic stability of these two states was rigorously controlled by the ionic strength, which changed the molecular charge by charge screening, and thus, remarkably changed the intermolecular interactions [40].

1.3 Application of protein and peptide nanomaterials 1.3.1 Drug delivery, tumor therapy, and tissue engineering In nature, self-assembly biomolecular building blocks have substantial influences on life process and activity. Protein and peptide as fascinating self-assembling biomolecules with unique advantages in availability and flexibility have become one of the ideal building blocks for biological applications. Drug delivery, which requires a stable, biocompatible, and efficient carrier for loading drug, can be realized by supramolecular protein- or peptide-assembling nanosystems [61–64]. One of the wide-applied supramolecular methods for drug delivery is the polymer encapsulation, in which the drug molecules are “protected” by the artificial polymer nanoparticles [65–67]. However, protein or peptide-assisted assembly of drug nanomaterials developed in recent years showed significantly enhanced efficiency in stability, drug loading,

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d­ elivery, and therapeutics, where the protein and peptide involved in the flexible interplay of interactions play the vital role that enables a better controlled and more precise assembly architecture than conventional methods [68–71]. Proteins such as human serum bovine (HSA) are natural vehicles for small molecules including drugs. Due to the unique biocompatibility and concrete structure, ­protein-loaded drug nanoparticles are highly attractive in drug delivery and phototherapy. Liu and coworkers reported HSA-bound IR825 dye nanoparticles, showing efficient imaging-guided photothermal therapeutic effects (Fig. 1.4) [72]. Individual IR825 molecule tended to self-assemble into large particles or even precipitates, while by binding with HSA via noncovalent encapsulation, the HSA-IR825 complex showed 5–10 nm of size and great stability at pH = 3–11. Compared with molecular IR825 in aqueous solution, the protein-bound complex nanoparticles showed enhanced fluorescence due to the inhibited dye aggregation by complexing with HSA and exhibited significant photothermal efficacy in vivo with complete tumor ablation upon irradiation of an 808 nm laser. Here, the protein improved phototherapeutic efficacy by forming a suitable particle size and reducing the rapid blood clearance. In addition to noncovalent encapsulation, the drug molecules can be covalently bonded with protein, resulting in a protein conjugate. The covalent bonding increases the nanoparticle stability and drug loading efficiency, provided that there are many anchors in the protein structure to bind drug molecules [73, 74]. Protein nanocages such as ferritin, apoferritin, and the capsid of cowpea chlorotic mottle virus (CCMV) also enable drug loading via their internal spaces. The drugs loaded in the protein cages can be phthalocyanine, methylene blue, cyanine derivatives, or many other kinds, where the loading stability mainly relies on the hydrophobic interactions between the drug molecules and the interior of protein cages. In some cases, there are also metal ion coordinating interactions that improve the drug loading stability [75, 76].

Fig. 1.4  Schematic illustration of the supramolecular assembly of HSA-IR825 complex and its optical performance. Adapted with permission from Q. Chen, C. Wang, Z. Zhan, W. He, Z. Cheng, Y. Li, Z. Liu, Biomaterials 35 (2014) 8206–8214. Copyright 2014, Elsevier.

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Peptide serving as a supramolecular assembly template provides an easier and more efficient route to modulate the assembly of functional molecules than protein does [76– 79]. Recently, Yan and coworkers reported that elegant fabrication of photosensitizer nanoparticles could be mediated by short peptide for enhanced photodynamic therapy [80]. Cationic diphenylalanine amide (CDP) and 9-fluorenylmethoxycarbonyll-lysine (Fmoc-K) induced the self-assembly of Chlorin e6 (Ce6) to afford stable nanoparticles (Fig. 1.5A). The nanoparticle stability mainly relied on the electrostatic

Fig. 1.5  Schematic representation of CDP- and Fmoc-K-tuned self-assembly of photosensitizers for PDT. (A) Self-assembly process and dominant intermolecular interactions. (B) FCNPs Internalization by MCF-7 cells. The red staining indicates photosensitizer, blue the nuclei, and green the cell membrane. (C) In vitro cytotoxicity of FCNPs. (D) In vivo tumor accumulation of FCNPs and free Ce6 imaged by fluorescence. (E) Tumor growth of different mouse groups. Reproduced with permission from K. Liu, R. Xing, Q. Zou, G. Ma, H. Möhwald, X. Yan, Angew. Chem. Int. Ed. 55 (2016) 3036–3039. Copyright 2016, Wiley-VCH.

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interaction between cationic CDP/Fmoc-K and anionic Ce6 in addition to the hydrophobic effect. The sizes of the resultant CDP/Ce6 and Fmoc-K/Ce6 nanoparticles (CCNPs and FCNPs) could be flexibly controlled by the assembly ratio of the starting building blocks. Due to the responsiveness to pH and specific enzymes, these nanoparticles enabled controlled drug release. FCNPs and CCNPs also showed efficient internalization to MCF-7 cells and large enhancement in photocytotoxicity compared with free Ce6 (Fig. 1.5B, C). Guided by in vivo fluorescence imaging, PDT treatment using CCNPs or FCNPs revealed enhanced tumor ablation (Fig. 1.5D, E) without causing organ damage or body weight loss. All these results proved the availability of the short peptide for finely tuning the self-assembly of functional organic species. Peptide-mediated supramolecular assembly for enhanced drug delivery and therapy can also be realized by covalent conjugation with drug molecules. Yan and coworkers further synthesized a porphyrin-dipeptide conjugate (TPP-G-FF), which individually self-assembled into highly stable photothermal nanomaterials [81]. Here, the peptide plays a dual role of both an assembly template and an amphiphilic group that tunes the hydrophobicity of photosensitizer. The conjugate self-assembly was driven by strong hydrophobic interactions, resulting in stable nanodots with a diameter of 25 ± 10 nm (Fig. 1.6A, B). The closely stacked conjugate exhibited complete fluorescence quenching of and inhibited generation of reactive oxygen species, which gave rise to 54.2% photothermal efficiency. Such a high efficiency rendered in vivo photoacoustic imaging, which further guided photothermal treatment at 24 h after intravenous injection. The temperature at the tumor region increased to 58.1°C under continuous laser treatment, and the tumors were completely ablated without recurrence, demonstrating the excellent photothermal efficacy of the porphyrin-peptide conjugate nanodots (Fig. 1.6C–E). Peptide-based hydrogels that integrated various functions are a class of attractive soft materials for tissue engineering. Different from nanoparticles, peptide-based hydrogels rely on a lot of intermolecular electrostatic or hydrogen-bonding interactions to maintain their supramolecular structure with controllable mechanistic and chemical properties. Recently, Yan and coworkers proved that poly-lysine and Fmoc-FF could form supramolecular hydrogels with switchable shear-thinning and self-healing properties suitable for injection. The hydrogel formation was driven by the strong electrostatic interactions between the poly-lysine and Fmoc-FF building blocks [19, 82]. Nilsson and coworkers found that controlling the hydrogel viscoelasticity could be realized by a stimulus-responsive manner. The Ac-(FKFE)2-NH2 peptide-based nanofibril hydrogels could be rigidified via either oligonucleotide-mediated or ­lectin-mediated cross-linking strategies, both of which were based on noncovalent interactions. These noncovalent cross-linking strategies could be readily employed to tune the peptide hydrogels for a wide range of biological applications like drug delivery and wound healing [83].

1.3.2 Biomimetic light-harvesting nanomaterials Sunlight fuels the origin and evolution of life. Mimicking natural cellular microcompartments like bacteria and chloroplasts are highly appreciated because they may provide opportunities not only to understand the complex evolutionary processes, but

Fig. 1.6  (A) Molecular structure and schematic self-assembly of the peptide-porphyrin conjugate. (B) TEM images of the afforded nanodots. (C) Thermal image of the tumor-bearing mice under continuous irradiation after intravenous injection of nanodots. (D) Tumor temperature at different irradiation time. (E) Tumor volume variations of different mice groups after photothermal treatment. Reproduced with permission from Q. Zou, M. Abbas, L. Zhao, S. Li, G. Shen, X. Yan, J. Am. Chem. Soc. 2017, 139, 1921–1927. Copyright 2017, American Chemical Society.

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also to create new biocatalytic utilization systems [84]. Rational design of complex biocatalysts that mimic the structure and functions of natural systems can be based on either proteins or peptides, where the proteins afford well-ordered supramolecular architectures as scaffolds and the peptides serve as templates to flexibly control supramolecular assembly. The most widely applied method to create new biocatalysts using protein is site-specific modification of catalytic moieties onto protein surface, where the catalytic moieties are usually metal-contained. A high catalytic activity can be obtained by this strategy using various metal complexes. Ueno and coworkers utilized protein assembly for attaching multiple catalytic sites on the protein surfaces. A triple-stranded β-helix fusion protein [(gp5bf)3]2 was dual-modified by Lys and Cys on the surface with Ru and Re complexes, which promoted a photo-catalytic CO2 reduction (Fig. 1.7.) [85]. The improvement of catalytic reactivity probably originated from the proximity effect when Re and Ru complexes were fixed at the Lys and Cys residues.

Fig. 1.7  (A) Supramolecular structure of [(gp5bf)3]2 with surface modification of Re and Ru complexes. (B) Scheme of the photo-catalytic reaction from CO2 to CO. Adapted with permission from N. Yokoi, Y. Miura, C. Y. Huang, N. Takatani, H. Inaba, T. Koshiyama, S. Kanamaru, F. Arisaka, Y. Watanabe, S. Kitagawa, T. Ueno, Chem. Commun. 47 (2011) 2074–2076. Copyright 2011, Royal Chemical Society.

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Another strategy includes the use of protein cage assemblies which can form unique reaction environments for the catalytic organic reactions via the catalysts inside their interior space. For example, within the discrete space of apoferritin in aqueous media, polymerization of phenylacetylene derivatives could be promoted by Rh complexes with narrow molecular weight distribution [86]. Compared with proteins, peptide- or even amino acid derivative-based biomimetic light-catalytic systems keep the advantages of dynamic, flexible, reversible, and controllable noncovalent interactions during supramolecular assembly [87]. Yan and coworkers employed Fmoc-K as a model template for the self-assembly of a light-­ harvesting agent TPPS [88]. The two building blocks co-assembled into u­ rchin-like light-harvesting antenna fibers, which allowed subsequent in situ growth of platinum nanoparticles on their surface triggered by photo-reduction of K2PtCl4 solution. The resulting hierarchical antenna first captured light by TPPS, then underwent charge transfer from TPPS to platinum nanoparticle, on which H+ was reduced to H2. Further, replacement of Fmoc-K to dilysine (KK) could result in a similar supramolecular system enabling mimicry of green sulfur bacteria nanoarchitecture [89]. The ­peptide-mediation is, thus, an easy and effective supramolecular route for biomimetic nanomaterials producing green energy [90]. The peptide-mediated coassembly also enables construction of complicated hierarchical architectures. Better than the Fmoc-K/TPPS/platinum photo-catalytic nanosystem, Yan and coworkers further developed a complex system for simultaneously mimicking the structure and function of chloroplasts (Fig. 1.8) [91]. In the nanosystem, photo-catalytic nicotinamide adenine dinucleotide (NADH) played the role of

Fig. 1.8  (A) Schematic mechanism of the chloroplast photosynthesis. (B) Proposed selfassembly structure. (C) Hierarchical supramolecular assembly for biomimetic photo-catalytic system. Reproduced with permission from K. Liu, C. Yuan, Q. Zou, Z. Xie, X. Yan, Angew. Chem. Int. Ed. 56 (2017) 7876–7880. Copyright 2017, John Wiley and Sons.

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driving enzymatic redox reaction, TPPS a photocatalyst, ADH a biocatalyst, and zinc and cystine the self-assembly template. The zinc ion-directed self-assembly of cystine with TPPS resulted in symmetric nanorods-consisted spherical crystals. Then, platinum nanoparticles grew in situ by photo deposition. Platinum nanoparticles efficiently reduced H+ to H2 following the electron transfer from the photo-excited TPPS, which simultaneously coupled the oxidation of NADH to NAD+. The NAD+ can be converted back by further coupling an ADH-catalyzed ethanol oxidation, well mimicking of the cylindrical grana in chloroplasts. Such a biomimetic photo-catalytic system enabled various photosensitizers like semiconductive ZnS nanoparticles [92], showing the unique advantage of peptides in modulating self-assembly architectures.

1.3.3 Semiconductive materials Highly ordered and directional protein or peptide self-assemblies have quantum confined structures which decrease the energy band gap into a semiconductive region. These ordered self-assembled architectures can work as semiconductors with flexible control, doping, and functionalization, providing a better interface between themselves and biological world than inorganic materials [93–97]. Biological systems contain a lot of well-ordered organizations such as filoviruses, annulus, and membrane proteins [98–100]. Recently, Kaminski and coworkers demonstrated that natural protein aggregates possess intrinsic semiconductive optical properties. Under photoexcitation, assemblies of misfolded proteins exhibited remarkable fluorescent emission [101]. Significantly, self-assembled short peptides may also have intriguing semiconductive properties, where intermolecular hydrogen-bonding interactions play a vital role for the ordered and robust supramolecular architecture [102, 103]. One of the intriguing properties of semiconductive peptide self-assemblies is photoluminescence. Under photoexcitation at 255 nm of wavelength, FF nanotubes exhibited fluorescence in the range of 350–500 nm. When excited at 340 to 380 nm, the FF nanotubes emitted excitation-dependent blue fluorescence, which was probably due to the quantum well confinement structures of the FF nanotubes [101]. This is an advantageous feature for light-emitting diodes. Another interesting example is the peptide nucleic acid (PNA) crystals. When the nucleic acid CG was chosen, aromatic and hydrogen-bonding interactions were introduced by Watson-Crick base-pairing, which resulted in broad emission with red-edge excitation shifting from 420 to 684 nm [95, 104]. Peptide semiconductors can also transmit photons along the axis under excitation, enabling them to serve as optical waveguides. Employed FF, hexagonal peptide microtubes were obtained via solvent thermal annealing [105]. The microtubes were hierarchically grown with a few micrometers of hollow diameter and several millimeters in length (Fig. 1.9A). By incorporating Nile Red (NR) as a guest dye compound, the microtube produced fluorescence at 515–560 nm under photoexcitation which was the intrinsic emission of NR. An optical waveguiding phenomenon was observed by the bright photo-luminescent spots appeared at both ends of the microtube and the weak luminescence from the tube body. In addition, a light on one end of the microtube caused the other end to appear bright, and the brightness of the end ­decreased

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as the propagation distance increased, further indicating an optical waveguiding ­phenomenon (Fig. 1.9B–D). Moreover, upon the introduction of an aldehyde to the oriented FF self-assembly, Yan and coworkers further fabricated three-dimensional fibrous peptide networks with inherently crystallized structure, exhibiting remarkable thermal stability and enhanced optical waveguiding performance [106].

Fig. 1.9  FF self-assembling microtubes for optical waveguiding. (A) Hierarchically selfassembled supramolecular architecture starting from FF building blocks; (B) SEM image of the hexagonal microtube; (C) Bright image of the microtube; (D) Optical waveguiding. Adapted with permission from X. Yan, J. Li, H. Möhwald, Adv. Mater. 23 (2011) 2796–2801. Copyright 2011, Wiley-VCH.

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1.4 Conclusions Supramolecular protein and peptide self-assembly is an attractive and expanding field with unique advantages such as flexible design, easy availability, and good biocompatibility. It aims to create novel functional materials through controlled organization. The controllability of the organization relies much on the realization of protein and peptide self-assembly mechanism, especially the noncovalent intermolecular interactions. While several effective strategies have been developed for the delicate protein self-assembly, peptide self-assembly is comparably easier and can be feasibly regulated by dynamic conditions. Since the protein and peptide architectures can be rationally designed and fabricated, they show broad potential applications in drug delivery and phototherapy, biomimetic photo-catalysis, semiconductive materials, and many other fields. In future, it will be highly appreciated to further develop the protein and peptide supramolecular materials in a more precise and multifunctional way. For example, (i) the de novo design of protein or peptide starting building blocks to arbitrary supramolecular architectures with desired functionalities, (ii) stimuli-responsive and dissipative systems, and (iii) system with structures over different length scales. All of these are expected to greatly promote the applications of protein and peptide nanomaterials in biology, electronic devices, and many other attractive fields.

Acknowledgment We acknowledge financial support from the National Natural Science Foundation of China (Project No. 21703252, 21522307, and 21473208), the National Natural Sciences Fund BRICS STI Framework Programme (51861145304), and Innovation Research Community Science Fund (No.21821005), the Key Research Program of Frontier Sciences of the Chinese Academy of Sciences (CAS, Grant No. QYZDB-SSW-JSC034).

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Self-assembly formation of peptide and protein nanofibers on surfaces and at interfaces

2

Shuwei Suna, Zhiqiang Sua, and Gang Weib,c a State Key Laboratory of Chemical Resource Engineering, Beijing University of Chemical Technology, Beijing, PR China, bCollege of Chemistry and Chemical Engineering, Qingdao University, Qingdao, PR China, cDivision of Functional Materials and Nanodevices, Ningbo Institute of Materials Technology and Engineering, Chinese Academy of Sciences, Ningbo, PR China

2.1 Introduction Many biological nanostructures and nanomaterials in nature are derived from biomolecular self-assembly and bioinspiration [1, 2]. For example, the DNA spiral structure in cells, the phospholipid membrane, and the third-level structure of proteins are formed by self-assembly of small molecular units under certain conditions [3]. Inspired by molecular self-assembly, it is possible to obtain various uniform biomaterials through the bottom-up approaches. As early as 1993, scientists have isolated an ionic short peptide containing 16 amino acid residues from a yeast protein, which exhibited excellent ability to form nanofiber structure via self-assembly [4]. Previously, a lot of one-dimensional (1D), two-dimensional (2D), and three-dimensional (3D) nanostructures and nanomaterials have been created by controllable molecular self-­ assembly through hydrogen bonding, hydrophobic/hydropholic, electrostatic, π-π, DNA hybridization, molecular recognition, and other interactions [5], and the fabricated nanomaterials have been widely used for biomedical engineering, tissue engineering, materials science, analytical science, and other material-related fields [6–8]. Self-assembled peptide and protein nanomaterials have attracted great attentions due to their wide applications and unique physical, chemical, biological, and electronic properties [9–13]. When studying the self-assembly of peptides and proteins, the surfaces of some inorganic, organic, and biological materials are usually used as the substrates to guide the self-assembly of peptide/protein molecules to form ordered nanostructures, such as nanofibers, nanowires, nanotubes, films, networks, and others [12, 14–16]. During the biomolecular self-assembly process, the formation of nanostructures is affected by pH, temperature, concentration of molecules, ionic strength, solution, and molecular sequence [17–19]. Therefore, the desired nanostructures can be obtained by adjusting appropriate conditions [20, 21]. Furthermore, since self-assembled structures are sensitive to the physical and chemical properties of the surface or interfaces, the surface/interface properties (including the hydrophobicity, hydrophilicity, and surface charge) of substrates play an important role in guiding Artificial Protein and Peptide Nanofibers. https://doi.org/10.1016/B978-0-08-102850-6.00002-4 © 2020 Elsevier Ltd. All rights reserved.

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the self-assembly process of biomolecules [22]. Therefore, understanding the interactions between peptides or proteins and the substrates/interfaces, as well as their self-­ assembly behavior, is of great significance for the design and synthesis of functional nanomaterials [23]. In this chapter, we present an overview on the self-assembly of peptides and proteins on/at various surfaces and interfaces, which guides the formation of ordered nanofibers. The effects of different surfaces of inorganic, organic, and biological materials on the formation of peptide/protein nanofibers are introduced and discussed in detail. In addition, the self-assembly and applications of peptide/protein nanofibers that are mediated by the solid-liquid, air-liquid, and liquid-liquid interfaces are demonstrated. It is expected that this work will be valuable for readers to understand the effects of various surfaces and interfaces on the formation of biological nanofibers and promote the fabrication of nanofiber-based functional nanomaterials and nanodevices for advanced applications.

2.2 Self-assembly formation of peptide/protein nanofibers on material surface In this part, we would like to introduce the self-assembly of peptide and protein molecules and the formation of nanofibers on inorganic, organic, and biological material surfaces.

2.2.1 Formation of nanofibers on inorganic material surface 2.2.1.1 Peptide nanofibers Peptides can self-assemble into nanofibers on the surface of many inorganic materials, such as mica, metal, non-metal, and glass [24–27]. It has been reported that the self-assembly of peptide on material surface was greatly influenced by the surrounding environment [28]. First, the growth and self-assembly of peptide nanofibers on mica surface were affected by pH. For instance, Yang et al. studied the growth of ionic-complementary peptide nanofibers on mica surface under different pH conditions [29]. They used atomic force microscopy (AFM) to monitor the growth and self-assembly of peptide nanofibers and found that self-assembled nanofibers were densely dispersed and short at lower pH, while the nanofibers were long in pure water (pH = 5.0). When pH increased to 9.9, no nanofibers appeared in a short time (Fig. 2.1A–C). In fact, the pH of solution changed the surface charge of peptide molecules, which in turn changed the peptide-peptide and peptide-surface interactions. With pH increasing, the charge on the surface of peptide nanofibers changed from positive charge to negative charge, while the force between nanofibers and mica surface changed from the electrostatic attraction to electrostatic repulsion. However, if the pH was too high or too low, the repulsive force between the peptide nanofibers inhibited the growth of peptide nanofibers. Based on the obtained results, they suggested that the growth of nanofibers

Self-assembly formation of peptide and protein nanofibers

500 nm

25

500 nm

1 mM HCl

Pure water

pH 3.4

pH 5.0

500 nm

1 mM NaOH pH 9.9

Fiber width - - + + - - ++ - - + + - - ++ - - + + - - ++ - - + + - - ++ - - + + - - ++ + + - - ++ - - - + + - - ++ - - + + - - ++ Nanofiber “seeds” on mica

Monomers and oligomers in solution Elongated nanofiber on mica

Fig. 2.1  Formation of peptide nanofibers on inorganic material surface. (A–D) Self-assembly of EAK16-II on mica: (A–C) AFM images of peptides on mica in (A) 1 mM HCl, (B) pure water, and (C) 1 mM NaOH solutions for 30 min. Scan area is 2000 nm × 2000 nm. (D) Corresponding self-assembly mechanism. Reproduced with the permission from H. Yang, S.Y. Fung, M. Pritzker, P. Chen, Surfaceassisted assembly of an ionic-complementary peptide: Controllable growth of nanofibers, J. Am. Chem. Soc. 129 (2007) 12200–12210. Copyright 2007, American Chemical Society.

seemed to follow a mechanism of nucleation and growth, as shown in Fig. 2.1D. Their work provides valuable experiences for understanding the self-assembly process of peptides on materials surfaces, as well as the potential applications for biomedicine, electrochemistry, and biosensing. The solvent property can also affect the self-assembly of peptides on inorganic material surface. For instance, Huang et al. investigated the self-assembly of diphenylalanine (FF) peptide on glass surface by testing pure water, methanol, and acetonitrile as solvent [27]. Compared with pure water, acetonitrile increased the self-assembly rate of peptide due to its low surface tension. In pure acetonitrile solvent, homogeneous peptide nanofibers were created, while in pure water peptide self-assembled into hollow tubes. Although both methanol and acetonitrile had high hydrogen-bond acceptor

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(HBA) ability and low surface tension, no peptide nanofibers were generated in high concentration methanol solution. The possible reason is that high hydrogen-bond donor (HBD) and HBA ability caused the solvation of peptide molecules. In addition, the surface structure and property of materials can affect the self-­ assembly of peptides to nanofibers. Yang et  al. investigated the self-assembly of ­ionic-complementary peptides EAK16-II on the surface of hydrophobic HOPG [30]. It can be found that peptides were arranged on HOPG surface in the direction of green stripes (Fig. 2.2A) in order to have more contact with hydrophobic HOPG surface. Therefore, self-assembled nanofibers were arranged in an orderly parallel or directional manner on HOPG (Fig. 2.2B). In addition, it was found that EAK16-II nanofibers were mainly formed in the terraces rather than HOPG step edge (Fig. 2.2C), where long nanofibers can actually be suspended from one platform to the next. The further control experiments proved that EAK16-II nanofibers were stable on HOPG surface under neutral (Fig. 2.2D), acidic (Fig. 2.2E), and alkaline (Fig. 2.2F) conditions. The obtained results indicated that EAK16-II interacted with HOPG through hydrophobic interaction rather than electrostatic interaction. In another study, Liao and coworkers studied the self-assembly mechanisms of nanofibers from peptide amphiphiles (NapFFKYp) and found that the self-assembled morphologies (nanofibers and nanosheets) were affected by some factors such as the peptide concentration, incubation time, and substrates [31].

Fig. 2.2  Formation of peptide nanofibers on HOPG: (A) The orientation of assembled EAK16-II strands; (B, C) AFM images of nanofibers on HOPG surface with different orientations; (D–F) AFM images of nanofibers on HOPG surfaces after immersing in (D) water, (E) 10 mM HCl, and (F) 10 mM NaOH for 10 h. Reprinted with the permission from H. Yang, S.Y. Fung, M. Pritzker, P. Chen, Modification of hydrophilic and hydrophobic surfaces using an ionic-complementary peptide, Plos One 2 (2007) e1325. Copyright 2007, PLoS One.

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2.2.1.2 Protein nanofibers Similar to peptides, proteins can also self-assemble into nanofibers on the different inorganic materials surface, which can also be affected by various interactions between protein molecules and material surface. Hwang et al. explored the effect of ionic strength on the formation of silk-elastinlike Protein (SELP) nanofibers on mica and silicon surface and revealed the importance of the electrostatic interaction between protein and mica surface to the formation of protein nanofibers [16]. They found that mica surface provides sites for promoting the self-assembly of proteins, and therefore, SELP-415K can self-assemble into nanofibers on the mica surface rather than in the bulk state at the same concentration. By adjusting the salt concentration, they observed that the length of the fibers became shorter and were more spherical as the salt concentration was increased. It was clear that the negatively charged mica surface can interact with positively charged SELP through the electrostatic attractions. Under the appropriate ion concentration, mica surface promoted the self-assembly of protein nanofibers. The self-assembly of SELP415K on hydrophilic silicon surface was further studied, which exhibited also the ability of SELP-415K to form nanofibers. Apart from mica and silicon, proteins could self-assemble into nanofibers on the surface of gold surface. Chen et al. studied the growth and self-assembly of fibrinogen (Fg) nanofibrils on mica and Au(111) substrates [32]. It was found that Au surface provided more adsorption and higher ability to guide the formation of Fg nanofibrils than mica surface. They believed that the presence of cysteine residues in Fg molecules leads to strong interactions between Au and sulfide, driving the molecular self-­ assembly. The Au-S and non-covalent interactions indicated that the self-assembly of proteins on the gold surface involved both physical processes and chemical bonding. Due to its hydrophobicity, graphite, an important inorganic material, plays important roles in guiding/mediating the self-assembly of proteins to nanofibers. For example, Reichert et al. used the hydrophobic highly oriented pyrolytic graphite (HOPG) to induce the formation and topotactical orientation of Fg nanofibrils [33]. Fg was preferentially adsorbed on the surface of HOPG, and the formed nanofibrils followed the direction of the nanostructure of HOPG. The surface energy of step regions was relatively high with a high affinity, and Fg molecules may form protofibrils on its surface. As the protein concentration increased, the thickness of the nanofibers on the surface of HOPG increased. Interestingly, the fibers can be only formed during the cleaning and drying process, while the Fg nanofibrils were absent during the adsorption process. In another study, Ling and coworkers found that silk fibroin (SF) could reduce graphene oxide (GO) into reduced graphene oxide (RGO) under certain conditions and self-assemble into nanofibers on the graphene surface [34], as indicated in Fig. 2.3A. It can be found that graphene surface was covered with nanofibers with very high density (Fig. 2.3B). From the blue curve in Fig. 2.3C, the height of SF nanofibers was measured to be about 3.5 nm. And, the red curve in the illustration showed that the height can reach up to 9 nm, indicating that SF nanofibers grew on the double-sided graphene sheet. Importantly, SF self-assembled into nanofibers only when graphene was present, suggesting the importance of graphene surface on guiding the formation of nanofibers. Through control experiments, it was concluded that the self-assembly

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Fig. 2.3  Formation of silk fibroin nanofibrils on graphene nanosheets. (A) Schematic representation of the self-assembly of SF on graphene nanosheets. (B) AFM images and (C) height profile of self-assembled SF nanofibrils on graphene nanosheets. Reproduced with the permission from S.J. Ling, C.X. Li, J. Adamcik, S.H. Wang, Z.Z. Shao, X. Chen, R. Mezzenga, Directed growth of silk nanofibrils on graphene and their hybrid nanocomposites, ACS Macro Lett. 3 (2014) 146–152. Copyright 2014, American Chemical Society.

process of nanofibers on graphene surface was a specific in situ growth based on the sp2-hybridized carbon surface. Due to multiple hydrophilic properties and large surface area, graphene materials provide high potential for creating functional hybrid nanomaterials for biomedical and biosensing applications. Based on the above cases, it is clear that the surfaces of various inorganic materials provide good platforms for guiding the self-assembly of peptide and protein to nanofibers. Meanwhile, the self-assembly of peptides and proteins on inorganic materials surface is affected by many factors. Therefore, it is possible to regulate these conditions to control the self-assembly of peptide and protein nanofibers for various applications.

2.2.2 Formation of nanofibers on organic/biological material surfaces Like inorganic materials, organic material surface can guide the formation of peptide nanofibers. For instance, Liao et al. explored the self-assembly process of a small peptide amphiphile (NapFFKYp) on the surfaces of mica, HOPG, and polystyrene (PS) film [31]. They found that the organic PS film can promote the growth of nanofibers further to nanosheets. In addition, the thickness of nanosheets formed on PS film was thicker than that on mica and HOPG surfaces. Besides organic and inorganic material surface, peptides and proteins can self-­ assemble on biological material surfaces to nanofibers. Self-assembled peptide-based

Self-assembly formation of peptide and protein nanofibers

29

materials on cell membrane surface have been widely used in cell surface engineering [9]. For example, Wang et al. found that peptides containing D-amino acids can selectively self-assemble on the surface of cell membranes to form nanofibers and hydrogels [9]. Many factors played important roles in the distribution of peptides on cell membranes surface such as chirality and conformation of amino acids, charge, conformation and sequence of peptides, and aromatic restriction groups. By the experiments, they proposed the enrichment and self-assembly mechanism of D-peptides on the cell membrane surface, as shown in Fig. 2.4A. The results indicated that the self-assembled nanofibers of peptides containing D-amino acids interacted not only with membrane proteins, but also with negatively charged phospholipids through electrostatic interactions, resulting in the membrane enrichment of polypeptides containing D-amino acids. This study developed a novel D-peptide-based biomaterial, holding great potential for tissue engineering, drug delivery, and regenerative medicine. In another study, Zhang et al. investigated the self-assembly process of Aβ26-35 peptides on the lipid membrane [35]. It was found that Aβ26-35 peptide with positive charges interacted with the negatively charged lipid membrane through electrostatic interactions, forming highly ordered parallel-oriented nanofibers on the membrane surface. The formation of nanofibers was related to the properties of lipid membrane, for instance, the good lipid fluidity facilitated the formation of parallel-oriented nanofibers. Their study provided a pathway for the formation of supramolecular structures on biomaterial surface. The aggregation of α-synuclein (αS) can form amyloid-like nanofibers, which leads to Alzheimer’s, Parkinson's, and other diseases. αS has unique structure and dynamic characteristics and can interact with membranes and biofilms [36]. Therefore, the understanding of the effects of lipid membranes on the formation of αS amyloid nanofibers will facilitate the regulation of the production of amyloid fibers [36, 37]. Fusco and coworkers studied the conformational properties of αS during its binding to lipid membrane by combining solution and solid-state nuclear magnetic resonance (NMR) techniques [36]. They found that the affinity between αS and lipid membrane was affected by the structural properties of αS, as well as the composition and physical properties of the membrane. The protein contains three distinct regions (the N-terminal, central, and C-terminal segments) with different properties and structures, which interacted with the lipid membrane in different ways. It can be clearly seen that a C-terminal fragment (green) had almost no interaction with the lipid membrane, while the N-terminal region (blue) was only rigidly combined on the membrane, and the central region (grey) played a vital role in regulating the affinity of αS to the membrane (Fig 2.4B). Like inorganic materials and organic materials, the self-assembly of αS on the biomaterials surface was also affected by the external conditions. Galvagnion et al. investigated the formation of αS nanofibers on the phospholipidsbased small unilamellar vesicles (SUVs) [37]. In the presence of high concentrations of SUVs, it is difficult for the formation of amyloid protein nanofibers. In contrast, when SUV concentration was low, amyloid fibers formed rapidly and the formation process was independent of the average size of the lipid vesicle. Besides, the amyloid fibers did not damage the lipid vesicles. It can be concluded that the existence of

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Artificial Protein and Peptide Nanofibers Cellular membrane Carbohydrate chains

Selfassembling peptides with D-amino acids Globular protein Phospholipid (negatively charged)

1 6

25

Nonpolar region of membrane protein

98

140

(A)

(B) Fig. 2.4  Formation of peptide nanofibers on biological material surfaces: (A) Self-assembly and formation of nanofibers of D-amino acids on cell membrane. (B) Self-assembly and formation of αS nanofibers on lipid bilayers. (A) Reprinted with the permission from H.M. Wang, Y.Z. Wang, A.T. Han, Y.B. Cai, N.N. Xiao, L. Wang, D. Ding, Z.M. Yang, Cellular membrane enrichment of self-assembling d-peptides for cell surface engineering. ACS Appl. Mater. Interfaces 6 (2014) 9815–9821. Copyright 2014, American Chemical Society. (B) Reprinted with the permission from G. Fusco, A. De Simone, T. Gopinath, V. Vostrikov, M. Vendruscolo, C.M. Dobson, G. Veglia, Direct observation of the three regions in alpha-synuclein that determine its membrane-bound behaviour. Nat. Commun. 5 (2014) 3827. Copyright 2014, Nature Publishing Group.

SUVs can improve the primary nucleation rate of αS. It is better to understand the self-assembly of proteins and the treating of diseases through characterizing the dynamic structure and properties of the combination of αS and membrane. Importantly, compared with inorganic and organic materials, self-assembled biological nanofibers of peptides and proteins on biomaterial surfaces have better biocompatibility and wider bio-related applications.

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2.3 Self-assembly formation of peptide and protein nanofibers at interfaces In this section, the self-assembly formation of peptide and protein nanofibers and nanofiber-based nanostructures at the solid-liquid, air-liquid, and liquid-liquid interfaces will be introduced and discussed.

2.3.1 Formation of nanofibers at solid-liquid interfaces Surface-mediated self-assembly of biomolecules to various nanostructures at ­solid-liquid interfaces has attracted much attention in the last years [38]. The study on the self-assembly formation of protein and peptide nanofibers at solid-liquid surfaces is helpful for understanding the action mechanisms of amyloid-based diseases like Alzheimer's, Parkinson's, type II diabetes, and others. Niu et al. investigated the formation of molecularly tuned peptide lamella structure at the HOPG-terephthalic acid (TPA) interface by scanning tunneling microscopy (STM) [39]. The short pentapeptide (NH2-Ala-Ala-Ala-Ala-COOH) (5Ala) and the terpyridine derivative (BT-O-C16) were co-assembled to nanoscale lamella structure at the HOPG-TPA interface. Organic molecules had the advantages of being designable and functional, which can facilitate and regulate the self-assembly process at interfaces. They observed that the peptide self-assembled into lamella structure at the solid-liquid interface, and the dimer peptides were encapsulated in the nanopore of the BT-O-C16 network. The hydrogen bonds were formed between 5Ala and BT-O-C16. This method of using organic molecules to regulate the self-assembly of biological molecules at the solid-liquid interface has potential significance in biomedical applications. In another case, Matmour and coworkers reported the self-assembly of oligo (p-phenylenevinylene) (OPV)-peptide conjugates at the solid-liquid interface [40]. The OPV-peptide conjugates were combined with 1-octanoic acid through hydrogen bonds, making it difficult to polymerize. However, HOPG provides the electrical conductivity and chemical inertness, providing an excellent platform for the self-­assembly of OPV-peptide conjugates to 1D structure. In fact, the backbone of peptides was transferred to β-strand on HOPG, and the successive strands through hydrogen bonds formed antiparallel β-sheet. This synthetic strategy was very flexible and can be applied to the synthesis of whole family of π-conjugated peptide conjugates. External stimulation can also promote the self-assembly and formation of peptide nanofibers at solid-liquid interfaces. Previously, Yang and coworkers demonstrated the mechanical force-induced nucleation and growth of EAK-16II peptide (Fig. 2.5A) nanofibers at mica-water (Fig. 2.5B) and HOPG-mica interfaces (Fig. 2.5C) by using mica and HOPG as hydrophilic and hydrophobic interfaces [41]. They found that this peptide can interact with mica and HOPG via electrostatic and hydrophobic interactions, respectively, thereby resulting in different morphologies of peptide assemblies. In the water system, the force applied by the AFM tip leads to the truncation and displacement of peptide nanofiber segments, which indicates that the mechanical force exceeded the bonding strength or cohesive energy in peptide nanofibers. However, in

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Artificial Protein and Peptide Nanofibers

Fig. 2.5  Mechanical force-induced formation of peptide nanofibers at solid-liquid interfaces: (A) Peptide molecular model; (B, C) Mechanical force-induced formation of nanofibers on (B) mica-water and (C) HOPG-water interfaces; (D, E) Nanofibers formed on HOPG before (D) and after (E) applying mechanical force in the absence of peptide; (F, G) Nanofibers on HOPG after applying mechanical force and in the presence of peptide. Reproduced with the permission from H. Yang, S.Y. Fung, M. Pritzker, P. Chen, Mechanicalforce-induced nucleation and growth of peptide nanofibers at liquid/solid interfaces, Angew. Chem. Int. Ed. 47 (2008) 4397–4400. Copyright 2008, Wiley.

the presence of additional peptides, the applied mechanical force induced the growth of peptide segments to long nanofibers. Similar to mica surface, mechanical forceinduced peptide assembly and nanofiber growth were also found on HOPG surface (Fig. 2.5D–G). In the absence of peptides, the continuous long nanofibers (Fig. 2.5D)) were found to form on HOPG surface first, and then broken to short peptide segments under the mechanical force applied by AFM tip (Fig.  2.5E). When peptides were present in solution, mechanical force induced the formation of a lot of short nanofibers (Fig. 2.5F), which were tended to pack in hexagonal patterns on HOPG surface (Fig. 2.5G). This strategy to fabricate peptide nanostructure at solid-liquid interfaces exhibited high potential in the fields of DNA nanotechnology, biosensing, and other biomedical applications. Protein nanofibers can also be created by biomolecular self-assembly at the ­solid-liquid interfaces via hydrophilic and hydrophobic interactions between proteins and interfaces. For instance, Marsh et al. compared the adsorption and conformation changes of globular protein β-lactoglobuli at hydrophilic silicon-water and hydrophobic octadeyltrichiorosilane (OTS)-modified silicon-water interfaces [42]. At the hydrophilic silicon-water interface, the conformation of adsorption protein changed slowly after adsorption, resulting in irreversible adsorption and strong interaction with the surface. When adsorbed on the surface of hydrophobic OTS-silicon, however, globular proteins were bound more strongly, and the binding force was proportional to the time of binding between protein and hydrophobic surface. Besides, the exchange

Self-assembly formation of peptide and protein nanofibers

33

of nonionic surfactants on the pre-adsorbed layers of globular proteins on both surfaces also proves that the binding strength is related to time. In another study, Krishnan et al. studied the scaled interfacial activity of proteins at the hydrophobic solid-liquid interface [43]. The experiments showed that the energy of protein at hydrophobic solid-liquid interface was similar to vapor-liquid interface. It was found that the main driving force of protein adsorption and self-assembly was not related to protein bipolarity, but to solution concentration. This study showed a facile way to enhance the compatibility between biomolecules and the materials for fabricating novel functional biomaterials via protein self-assembly at solid-liquid interface.

2.3.2 Formation of nanofibers at air-liquid interfaces Amphiphilic peptides are easy to self-assemble at the air-liquid interface to form different nanostructures due to their hydrophilic and hydrophobic groups. For example, Engin et al. studied the adsorption and assembly of amphiphilic peptides on the air-water interface [44]. According to the rule of alternate arrangement of hydrophobic and hydrophilic residues, they designed the peptides with amphiphilic properties, which spontaneously formed an ordered monolayer at the air-water interface through the interactions of the β-hairpin-like units. By comparing the peptide structures at the air-water interface and in bulk water under the same conditions, it can be found that the interface was the boundary between hydrophobic and hydrophilic residues. Therefore, peptides exhibited an orderly structure at the air-water interface, while it was difficult for peptides to form uniform secondary structure in bulk water due to the shielding effect of hydrophobic residues in water. Segman-Magidovich and coworkers demonstrated the self-assembly and formation of peptide nanofibers and 2D nanosheet at the air-water interface [45]. The amphiphilic α/β-peptides were designed with 1:1 alternation of α- and β-amino acid residues along the peptide backbone. Two designed zwitterionic α/β-peptides were mixed with a 1:1 molar ration to form parallel and antiparallel strands (Fig. 2.6). The βEβK peptide assembled at the interface into nanofiber-based nanofilm, which was transferred by the Langmuir-Blodgett technique to a mica surface for AFM phase (Fig. 2.6A) and imaging (Fig. 2.6B and C) characterizations. It can be clearly seen that nanofibers with μm length are formed and aligned along the same direction. This study pointed at the way for creating new nanomaterials by designing peptides with ionizable side chains and their positions within the α/β-peptides. Xie et al. studied the self-assembly behavior of GAV-9a peptide at the air/­ethanolwater interface with mica as the substrate [46]. The introduction of ethanol-­containing atmospheres changed the water nanofilm on mica surface. They proved that the self-assembly of GAV-9a at the air-water interface tends to form flat nanofibers, while bent nanofibers at the ethanol-containing air-water interface. The thickness of self-­ assembled nanofibers was proportional to the proportion of ethanol in the atmosphere. Besides, they found that the formation of GAV-9a nanofibers induced by the substrate was determined by electrostatic interactions, and the main driving force of the epitaxial self-assembly process of GAV-9a was hydrophobic interactions. The introduction

Fig. 2.6  Formation of peptide nanofibers at air-water interface: AFM (A) phase image and (B, C) height image and section analysis of selfassembled nanofiber-based nanofilm at air-water interface, which was transferred onto a mica surface with Langmuir-Blodgett technique. Reproduced with the permission from S. Segman-Magidovich, M.R. Lee, V. Vaiser, B. Struth, S.H. Gellman, H. Rapaport, Sheet-like assemblies of charged amphiphilic alpha/beta-peptides at the air-water interface. Chem. A Eur. J. 17 (2011) 14857–14866. Copyright 2008, Wiley.

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of ethanol atmosphere can be used to adjust the properties of the interface and change the self-assembled peptide nanostructures. Furthermore, peptides can also be combined with other materials to self-assemble at the air-liquid interface. For instance, Koga et  al. synthesized a kind of peptide-­ diacetylene hybrid (PDH) by using β-sheet peptide and diacetylene [47], in which Leu and Lys sequences were arranged alternately. The designed peptide-diacetylene hybrid exhibited anisotropic self-assembly at the air-water interface, forming parallel β-sheet conformation and stable nanofiber-based Langmuir monolayer. Similarly, proteins were capable of self-assembling at the air-water interface to form nanofibers and other nanostructures. Ta et al. explored the self-assembly process of nontoxic wild-type (WT) and toxic (M8) amyloid proteins on the air-water interface [48]. They demonstrated that WT protein formed an antiparallel β-sheet structuration at the air-water interface and a parallel β-sheet structure in solution. In contrast, M8 could rapidly form antiparallel β-sheet structure at the air-water interface and in the solution. Besides, M8 revealed faster formation of nanofibrous film than WT at low concentration. In another study, Kisko et al. explored and compared the self-assembly process of two hydrophobins (HFBI and HFBII) at the air-water interface [49]. At the air-water interface, hydrophobins self-assembled into highly ordered nanofibrous films by specific protein-protein interactions in the lateral direction.

2.3.3 Formation of nanofibers at liquid-liquid interfaces Surface-active molecules tend to accumulate at interfaces between water and nonpolar liquids [50], which can also affect the self-assembly of peptides at the liquid-liquid interfaces. Previously, Nichols et al. demonstrated the rapid assembly of peptide (Aβ140) to β-sheet nanofibers at a chloroform-aqueous buffer interface [51]. They found that this two-phase system with chloroform promoted the aggregation of peptides with 1-2 orders of magnitude faster than in the buffer alone. In another study, Wang and coworkers reported the jet flow-directed self-assembly of peptide (Fmoc-FF) into nanofibers at the water-cationic polyacrylamide (CPAM) interface [52]. By controlling the jet flow of CPAM, it was possible to fabricate FF nanofiber-based macroscopic sac membranes or microfibers at the interface between two solutions. Using liquid-liquid interfaces for mediating the self-assembly of biomolecules will be an alternative way to fabricate highly ordered and functional bionanomaterials for drug delivery, enzymatic immobilization, biosensors, and tissue engineering.

2.4 Conclusions and outlooks In this chapter, we summarized the self-assembly of peptides and proteins into nanofibers by the guidance of various materials surfaces and solid-liquid/air-liquid interfaces. We mainly described the self-assembly of peptides and proteins on the surface of mica, glass, HOPG, and lipid membranes, as well as at the solid-liquid and air-­ liquid interfaces. The specific interactions between peptides and proteins with surfaces and interfaces, including electrostatic, hydrophilic, and hydrophobic interactions

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for affecting biomolecular self-assembly, were introduced and discussed. It can be concluded that the morphology and structure of self-assembled nanofibers are not only related to the properties of surface or interface, but also to the sequence and properties of peptides or proteins. Although great advances in the design, synthesis, and applications of self-­assembled peptide and protein nanofibers on surfaces and at interfaces have been made in the past few decades, there are many challenges to be faced. Optimal adjusting of the properties of materials (such as hydrophobicity and hydrophilicity) and the interfaces (such as the type and concentration of liquid) will be crucial to regulate the structure of nanofibers, but an optimized and accurate scheme has not been proposed. The controllable design and fabrication of peptide and protein nanofibers on cell membranes could be done, which will be helpful for us to understand the potential inhibition methods for amyloid-related diseases. In addition, the formation/inhabitation of biological nanofibers on the surface of nanoparticles is also suggested, which is important to develop functional nanofiber-based hybrid nanomaterials for drug delivery, regenerated medicine, tissue engineering, energy, environmental, and biosensing applications.

Acknowledgments Shuwei Sun and Zhiqiang Su acknowledged the financial support from the National Natural Science Foundation of China (No. 51573013 and 51873016). Gang Wei expressed thanks for the financial support from the National Natural Science Foundation of China (No. 51873225).

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[27] R.L.  Huang, W.  Qi, R.X.  Su, J.  Zhao, Z.M.  He, Solvent and surface controlled self-­ assembly of diphenylalanine peptide: From microtubes to nanofibers, Soft Matter 7 (2011) 6418–6421. [28] H. Li, F. Zhang, Y. Zhang, M. Ye, B. Zhou, Y.Z. Tang, H.J. Yang, M.Y. Xie, S.F. Chen, J.H. He, H.P. Fang, J. Hu, Peptide diffusion and self-assembly in ambient water nanofilm on mica surface, J. Phys. Chem. B 113 (2009) 8795–8799. [29] H.  Yang, S.Y.  Fung, M.  Pritzker, P.  Chen, Surface-assisted assembly of an ionic-­ complementary peptide: controllable growth of nanofibers, J. Am. Chem. Soc. 129 (2007) 12200–12210. [30] H. Yang, S.Y. Fung, M. Pritzker, P. Chen, Modification of hydrophilic and hydrophobic surfaces using an ionic-complementary peptide, Plos One 2 (2007) e1325. [31] H.S. Liao, J. Lin, Y. Liu, P. Huang, A. Jin, X.Y. Chen, Self-assembly mechanisms of nanofibers from peptide amphiphiles in solution and on substrate surfaces, Nanoscale 8 (2016) 14814–14820. [32] G.J. Chen, N.T. Ni, B.H. Wang, B.Q. Xu, Fibrinogen nanofibril growth and self-assembly on au (1,1,1) surface in the absence of thrombin, ChemPhysChem 11 (2010) 565–568. [33] J.  Reichert, G.  Wei, K.D.  Jandt, Formation and topotactical orientation of fibrinogen nanofibrils on graphite nanostructures, Adv. Eng. Mater. 11 (2009) B177–B181. [34] S.J. Ling, C.X. Li, J. Adamcik, S.H. Wang, Z.Z. Shao, X. Chen, R. Mezzenga, Directed growth of silk nanofibrils on graphene and their hybrid nanocomposites, ACS Macro Lett. 3 (2014) 146–152. [35] L. Zhang, J. Zhong, L.X. Huang, L.J. Wang, Y.K. Hong, Y.L. Sha, Parallel-oriented fibrogenesis of a beta-sheet forming peptide on supported lipid bilayers, J. Phys. Chem. B 112 (2008) 8950–8954. [36] G.  Fusco, A.  De Simone, T.  Gopinath, V.  Vostrikov, M.  Vendruscolo, C.M.  Dobson, G. Veglia, Direct observation of the three regions in alpha-synuclein that determine its membrane-bound behaviour, Nat. Commun. 5 (2014) 3827. [37] C. Galvagnion, A.K. Buell, G. Meisl, T.C.T. Michaels, M. Vendruscolo, T.P.J. Knowles, C.M. Dobson, Lipid vesicles trigger alpha-synuclein aggregation by stimulating primary nucleation, Nat. Chem. Biol. 11 (2015) 229–234. [38] Y. Yu, J.F. Hou, L.L. Yu, Y.L. Yang, C. Wang, Lattice modulation effect of liquid-solid interface on peptide assemblies, Surf. Sci. 649 (2016) 34–38. [39] L. Niu, X.J. Ma, L. Liu, X.B. Mao, D.X. Wu, Y.L. Yang, Q.D. Zeng, C. Wang, Molecularly tuned peptide assemblies at the liquid-solid interface studied by scanning tunneling microscopy, Phys. Chem. Chem. Phys. 12 (2010) 11683–11687. [40] R.  Matmour, I.  De Cat, S.J.  George, W.  Adriaens, P.  Leclere, P.H.H.  Bomans, N.A.J.M.  Sommerdijk, J.C.  Gielen, P.C.M.  Christianen, J.T.  Heldens, J.C.M.  van Hest, D.W.P.M.  Lowik, S.  De Feyter, E.W.  Meijer, A.P.H.J.  Schenning, Oligo(pphenylenevinylene)-peptide conjugates: Synthesis and self-assembly in solution and at the solid-liquid interface, J. Am. Chem. Soc. 130 (2008) 14576–14583. [41] H.  Yang, S.Y.  Fung, M.  Pritzker, P.  Chen, Mechanical-force-induced nucleation and growth of peptide nanofibers at liquid/solid interfaces, Angew. Chem. Int. Ed. 47 (2008) 4397–4400. [42] R.J. Marsh, R.A.L. Jones, M. Sferrazza, Adsorption and displacement of a globular protein on hydrophilic and hydrophobic surfaces, Colloids Surf. B. Biointerfaces 23 (2002) 31–42. [43] A. Krishnan, Y.H. Liu, P. Cha, D. Allara, E.A. Vogler, Scaled interfacial activity of proteins at a hydrophobic solid/aqueous-buffer interface, J. Biomed. Mater. Res. A 75a (2005) 445–457.

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[44] O. Engin, M. Sayar, Adsorption, folding, and packing of an amphiphilic peptide at the air/ water interface, J. Phys. Chem. B 116 (2012) 2198–2207. [45] S.  Segman-Magidovich, M.R.  Lee, V.  Vaiser, B.  Struth, S.H.  Gellman, H.  Rapaport, Sheet-like assemblies of charged amphiphilic alpha/beta-peptides at the air-water interface, Chem. A Eur. J. 17 (2011) 14857–14866. [46] M.Y.  Xie, H.  Li, M.  Ye, Y.  Zhang, J.  Hu, Peptide self-assembly on mica under ­ethanol-containing atmospheres: effects of ethanol on epitaxial growth of peptide nanofilaments, J. Phys. Chem. B 116 (2012) 2927–2933. [47] T. Koga, T. Taguchi, N. Higashi, Beta-sheet peptide-assisted polymerization of diacetylene at the air-water interface and thermochromic property, Polym. J. 44 (2012) 195–199. [48] H.P.  Ta, K.  Berthelot, B.  Coulary-Salin, B.  Desbat, J.  Gean, L.  Servant, C.  Cullin, S. Lecomte, Comparative studies of nontoxic and toxic amyloids interacting with membrane models at the air-water interface, Langmuir 27 (2011) 4797–4807. [49] K. Kisko, G.R. Szilvay, E. Vuorimaa, H. Lemmetyinen, M.B. Linder, M. Torkkeli, Serimaa, R. Self-assembled films of hydrophobin proteins hfbi and hfbii studied in situ at the air/water interface. Langmuir 2009, 25, 1612-1619. [50] L.R. Pratt, A. Pohorille, Hydrophobic effects and modeling of biophysical aqueous solution interfaces, Chem. Rev. 102 (2002) 2671–2691. [51] M.R.  Nichols, M.A.  Moss, D.K.  Reed, J.H.  Hoh, T.L.  Rosenberry, Rapid assembly of amyloid-beta peptide at a liquid/liquid interface produces unstable beta-sheet fibers, Biochemistry 44 (2005) 165–173. [52] Y.F.  Wang, W.  Qi, R.L.  Huang, R.X.  Su, Z.M.  He, Jet flow directed supramolecular self-assembly at aqueous liquid-liquid interface, RSC Adv. 4 (2014) 15340–15347.

Fabrication of amyloid nanofiber matrices by electrospinning

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Devina Jaiswala, Sara Katebifarb, Swetha Rudraiahc, and Sangamesh G. Kumbarb,d a Department of Biomedical Engineering, Western New England University, Springfield, MA, United States, bDepartment of Biomedical Engineering, University of Connecticut, Storrs, CT, United States, cDepartment of Pharmaceutical Sciences, School of Pharmacy, University of Saint Joseph, Hartford, CT, United States, dDepartment of Orthopedic Surgery, University of Connecticut Health Center, Farmington, CT, United States

3.1 Introduction Amyloid fibrils form spontaneously from a soluble protein, which undergoes a conformational change to form insoluble fibrils [1]. These are formed extracellularly, and for decades, they have been associated with diseases such as Alzheimer’s disease, Diabetes type 2, and spongiform encephalopathies [2]. However, amyloid fibers also have functional application such as the formation of biofilms or protective surface on eggs [3]. Due to the exceptional mechanical properties and resistance to degradation, amyloid fibers and amyloidogenic peptides are being recognized as nanomaterials [3]. Structurally, it is a quaternary protein formed from peptides and “misfolding” of globular proteins [4]. The X-ray diffraction pattern revealed stacked β-sheets running perpendicular to the fiber axis with an interstrand gap of 4.7 Å and an intersheet gap of ~ 10 Å [5]. A typical fibril is 7–10 nm in diameter as seen under the scanning electron microscope. An amyloid fiber is formed by winding of two protofilament subunits whose tensile strength (0.6 ± 0.4 GPa) is comparable to steel (0.4–0.55 GPa) [6, 7]. As per a report, a typical amyloid fiber has a Young’s modulus of 3.3 ± 0.4 GPa [3]. The remarkably high mechanical properties of amyloid fibers can be attributed to the hydrogen-bonded β-sheets and hierarchical fiber arrangement [8]. This type of hierarchical structure of fiber formation can also be seen in collagen and silk where peptides form polypeptide chains that organize themselves into an ordered structure, thus making them amyloid-like fibers [9]. In vitro, a polypeptide can be triggered to self-assemble into amyloid-like fibrils using variations in pH, temperature, or ionic concentration. Addition of pH-sensitive peptides or charged groups can aid in self-assembly of fibrils [10]. Self-assembly can be explained by two accepted models: template assistance and nucleation polymerization [11, 12]. Since self-assembly can be a slow process, short segments of peptide fibrils can act as a template or nucleus to promote faster fibril formation and bring about a conformational change in the soluble peptides present in the solution to form amyloid-like fibers [13]. The seed helps the existing peptides to overcome the high activation energy needed to bring about the conformational change. Artificial Protein and Peptide Nanofibers. https://doi.org/10.1016/B978-0-08-102850-6.00003-6 © 2020 Elsevier Ltd. All rights reserved.

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Fig. 3.1  Schematic representation of the wet-spinning process.

Besides self-assembly, amyloid-like nano- and microfibers can be produced using techniques such as (1) wet-spinning [14] or (2) electrospinning [15], producing fibers that closely mimic the fibers formed in vivo. (1) In wet-spinning, a solution of protein nanofibrils, formed spontaneously under ambient conditions, is poured over negatively charged support solution. The film formed at the interface of the two solutions passes through small holes in a spinneret to form microfibers (Fig. 3.1). The morphology of the fibers formed from this technique is dependent on the interfacial film thickness. The polyionic self-assembly at the interface and the strength of electrostatic interaction of the oppositely charged ions determine the length of the spun fiber [14]. Fibers in the diameter range of 10–60 μm have been wet spun to mimic natural spider and silkworm silk. (2) Electrospinning is widely used in textiles and in medical applications for making fibrous matrices from polymers. The technique has been used to form nano and microfibers from proteins which can be cross-linked to mimic morphology and physical strength of native polypeptide fibers. Some common scaffolding proteins found in nature are collagen, gelatin, silk, fibronectin, elastin, and bovine serum albumin (BSA) [16]. They can be easily electrospun to form matrices with fiber diameter and overall morphology matching extracellular matrix (ECM). The operating parameters of electrospinning can be optimized to produce nano or microfibers to produce engineered results [17].

This chapter will focus on the basics of electrospinning and how various operational parameters can affect fiber morphology and diameter. We will also discuss the electrospinning of amyloid-like fibers from proteins such as silk, collagen, and BSA and optimized parameters that can be used to make biomimicking fibrous matrices for biomedical applications.

3.2 Electrospinning Electrospinning is a technique to produce nanofibers by applying an electric field to a polymer jet. The basic theory of electrostatic attraction of liquids can be traced back to the seventeenth century when William Gilbert reported deformation of a water droplet on a dry surface when exposed to a piece of rubbed ambe [18].

Fabrication of amyloid nanofiber matrices by electrospinning43

Fig. 3.2  Growing popularity of electrospinning (Web of Science).

In the 1930s, Formhals patented the process of production of artificial filaments of a natural polymer such as cellulose acetate [19]. The threads produced were received on a moving thread collecting device enabling the production of a spool of thread which could be unwound. The mechanism behind the formation of threads from the tip of the fluid source was later explained by Taylor in 1969 [20]. Though the technique was being used for industrial applications such as the production of nonwoven plastics, the use of this technology was revived in the 1990s at University of Akron by Reneker’s group who studied fabrication of 0.05 to 5-μm diameter fibers from various polymer solutions [21]. After this study, the application of electrospinning escalated in academia from nanotechnology and drug delivery to serving as a biomimicry scaffold for tissue engineering (Fig. 3.2) [22].

3.2.1 Basic technique Electrospinning is based on the application of an electric field between polymer source and collector target. The setup, shown in Fig. 3.3, needs three basic components: conducting capillary connected to a syringe, high voltage source, and collector [23]. The polymer is ejected at a steady flow rate to develop a spherical bead at the tip of the capillary source due to the surface tension of the polymer solution. The positive terminal of the voltage source is applied to the capillary and negative terminal is connected to the target. When an electric field is applied, the positive charge collects at the spherical polymer bead which causes its deformation. The polymer fluid forms a cone, also known as “Taylor Cone,” which remains stable as long as the surface tension and electrostatic forces are balanced [24]. The droplet experiences two types of electrostatic forces: electrostatic repulsion (caused between surface charges) and Coulombic force, produced due to the external electric field [25, 26]. With the steady increase in an electric field, the electrostatic forces overcome the surface tension and a stable string of polymer pulls out of the Taylor cone and is accelerated towards the target. As the polymer travels towards the target, the solvent evaporates and the fiber undergoes elongation and whipping to deposit a mat of nonwoven solid fibers at the target [22, 27].

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Artificial Protein and Peptide Nanofibers

Fig. 3.3  Schematic of electrospinning process. Electrically charged polymer solution ejected at a controlled rate form nanofibers at the target. The polymer solution and target are connected to high voltage power supply.

3.2.2 Principle The term electrospinning is derived from “electrostatic spinning.” The principle behind the formation of the jet from an electrified liquid has been used for mass spectroscopy, metallic powder production, inkjet printing, micro-spraying, and spray painting [28]. When an evaporating liquid droplet is placed in an electric field, the droplet becomes unstable as the electrostatic forces overcome the attractive surface tension. This leads to the formation of daughter droplets, thereby leaving behind a stable drop. The instability in droplet was first explained by Lord Rayleigh [29] for conducting droplet of radius a and charge q [30, 31]. q 2 = 64π 2 0γ a 3

(3.1)

where ϵ0 is the permittivity of free space and γ is the surface tension of the droplet. This is also called Rayleigh limit. With the application of electric potential, the pendant formed at the tip of the capillary undergoes deformation. The forces exerted by the surface tension pull the droplet inside the nozzle, whereas the electrostatic forces pull the droplet towards the oppositely charged electrode. The ellipsoidal shape of the droplet forms a pointed cone called the Taylor cone (Fig. 3.4) [32]. Theoretical derivation by Taylor (1964) states that the forces at the surface of a cone are under equilibrium at a semi-vertical angle of 49.3 degree, called Taylor angle (Fig. 3.5). Electrospinning is initiated when the threshold voltage is reached and the balance between the forces becomes independent of the radius of curvature of the apex of the fluid. The electric field strength (E) at the apex of the droplet is given by [21] E=

4γ 0 R

(3.2)

where γ is surface tension of the droplet, ϵ0 is the permittivity of free space, R is the radius of curvature of the apex of the fluid.

Fabrication of amyloid nanofiber matrices by electrospinning45

Dripping

Coning

Stable Cone-jet

Multi-jet

Increased voltage

Fig. 3.4  Taylor cone formation under different voltage conditions. As voltage increases, the jet formed at the tip of the needle stabilizes which shows balance in electrical charges.

Needle

Needle

Fig. 3.5  Taylor cone charge distribution when the needle carrying polymer solution is positively charged. The positive charge distributes along the Taylor cone formed at the tip of the needle.

To explain the relationship between surface tension and electric field, Santos et al. conducted a series of experiments on water droplet suspended from a nozzle. Time lapse images of water droplet subjected to varying voltages were analyzed to predict the formation of water jet at a critical voltage of − 9.5 kV. Beyond this voltage, the fluid showed an excess of surface charge (same charge sign as the electrified needle bias voltage sign) leading to increase in surface area, stretching into threads, and further dividing into finer threads [28].

3.2.3 Parameter optimization Electrospun fiber morphology is mainly dependent on three broad parameters: solution properties such as viscosity, surface tension, polymer concentration, the ­molecular

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Artificial Protein and Peptide Nanofibers

weight of the polymer, and electrical conductivity; operational conditions such as fluid flow rate, applied voltage, the distance between the spinneret and target; and ambient conditions such as humidity and temperature [33, 34]. These parameters can be optimized to obtain fiber diameters ranging from 10 nm to 10 μm, beaded or bead-free fibers, and ribbons or cylindrical fibers. Fiber uniformity and diameter are influenced by various factors, but primarily by polymer solution concentration and solution conductivity. Other secondary factors include applied voltage, feed rate, and temperature.

3.2.3.1 Primary factors affecting fiber morphology Fiber morphology is an important feature of the electrospun matrices for cell-based studies. There are studies that have shown the influence of fiber diameter on cell response [35]. Cellular mechanosensors detect the fiber curvature which is signaled to the nucleus to upregulate or downregulate transcription factors that influence cell proliferation and differentiation [36, 37]. Primarily, solution concentration and conductivity have maximum impact on the fiber morphology compared to other factors influencing electrospun fibers [25, 38].

Concentration One of the major factors that affects fiber diameter is solution concentration. Formation of continuous fibers is dependent on upper and lower concentration cut-offs for each polymer solution [39]. Concentrations higher than the upper limit would not result in fiber formation since the viscosity of the solution increases and resists fluid flow from the needle tip. At concentrations below the lower cut-off, the surface tension of the droplet has a more dominant effect on the fiber morphology. Generally, very low concentration solutions form droplets rather than continuous fibers. As per power law, the fiber diameter of polymer fibers can be related to solution concentration as in equation (3.3) [40]. Average fiber diameter = ( Concentration )

3

(3.3)

Following power law, increase in fiber diameter with concentration was observed for polycaprolactone (PCL) fibers, though the uniformity of fibers was compromised at high solution concentrations [33]. A bimodal diameter distribution was observed for PEO/water fibers at higher concentrations [39]. Reason being that, at higher concentration, the polymer chains entangle more compared to lower concentrations which resists the stretching effect from the electric charge. A threshold concentration of 1% poly(l-lactic acid) PLLA dissolved in dichloromethane (DCM)/pyridine was observed to produce ultrafine fibers (19 ± 6 nm). Concentrations below the threshold produced beaded fibers due to decreased solution viscosity [41]. Likewise, the molecular weight of the polymer also affects the fiber morphology since it affects solution viscosity [42]. At constant solution concentration, viscosity of a solution increases with molecular weight of the polymer. Due to polymer chain entanglement, the solution maintains its continuity while stretching and continuous fibers can be obtained for a polymer with optimum molecular weight. Inadequate

Fabrication of amyloid nanofiber matrices by electrospinning47

p­ olymer stretching can result in beaded fibers. The morphology of the beads can change with polymer molecular weight leading to spherical- or spindle-shaped beads [43]. To control fiber diameter and produce bead-free fibers, solution concentration and polymer molecular weight should be balanced. Both molecular weight and polymer concentration can be linked to polymer chain entanglement [44]. The critical entanglement number (ne, number of entanglement/ chain) has been calculated at ne ~ 2 [45]. For dilute solutions, ne  cc), the entanglement number can reach up to 3 which produces bead-free fibers. Entanglement number is affected by concentration or volume fraction and molecular weight of the polymer. Higher molecular weight polymers can achieve critical entanglement number of 2 at lower concentrations than the same polymer with lower molecular weight [44].

Electrical conductivity The electrical conductivity of the solution is another major factor that determines fiber formation and diameter. Solvents with low electrical conductivity can lead to the formation of beaded or discontinuous fibers [46, 47]. Higher electrical conductivity means more charge density on the jet which favors elongation of fibers under the high electric field [48]. Solvents such as methylene chloride can result in beaded fibers due to its inherently low electrical conductivity of 4.30 × 10− 11 μS/cm. Bead-free fibers can be produced by adding a small percentage of solvents with high electrical ­conductivity such as ethanol, N,N-dimethylformamide (DMF), tetrahydrofuran (THF), and pyridine. Tan et  al. compared solvent mixtures with different electrical conductivity, such as DCM, DCM/DMF, and DCM/pyridine [41]. Ultrathin fibers ­(diameter ~ 100 nm) were produced from a solvent mixture with the highest electrical conductivity. Electrical conductivity of DCM mixed with different percentage of pyridine revealed a critical percentage of mixture ratio (50:50); beyond this threshold, there was no increase in electrical conductivity of the solvent system [41]. A small portion of ethanol or DMF can be added to DCM to spin bead-free fibers since it improves solvent’s electrical conductivity [49]. Solvents from different suppliers and grades can have varying conductivity. Uyar et al. compared different grades of DMF and correlated it with polystyrene fiber morphology and diameter [50]. Solvent with lower conductivity produced beaded fibers due to lower charge density. Table 3.1 lists the electrical conductivity of commonly used organic solvents for electrospinning. Electrical conductivity can also be improved by adding a trace amount of salt to the polymer solution [51]. Addition of small percentage ionic salt such as sodium chloride (NaCl) resulted in bead-free PCL [33] and poly(d,l-lactic acid) (PDLA) [51] fibers, while beaded fibers were obtained for a solution with no salt. Salt increases the overall charge density of the jet which increases the elongation forces experienced by the jet during electrospinning [52]. The pulling force and jet tension cause reduction in beads’ size and produce spindle-shaped beads or bead-free fibers. Thus, under same

48

Artificial Protein and Peptide Nanofibers

Table 3.1  Electrical conductivity of commonly used organic solvents for electrospinning. Solvent

Electrical conductivity

Acetic acid Acetonitrile Dimethylformamide Nitrobenzene Pyridine 2-Nitropropane Acetonitrile Ethanol Methylene chloride Chloroform Acetone Tetrahydrofuran Dimethylformamide

6.00 × 10− 9 6.00 × 10− 10 6.00 × 10− 8 2.00 × 10− 10 4.00 × 10− 8 5.00 × 10− 7 6.00 × 10− 10 1.40 × 10− 9 4.30 × 10− 11  6 kV) is sufficient for initiation of jet formation from Taylor cone [20]. Defect-free, cylindrical fibers were formed at 5.5 kV applied voltage for poly ethylene oxide (PEO)/water system [39]. Lower applied voltage can be beneficial in producing finer fibers since low voltage would increase the time of flight [54]. This would cause additional stretching of the fibers before it reaches the target. With the increase in voltage at a constant feed rate, the shape of the droplet formed at the tip of the needle deforms which affects fiber diameter and bead formation. High applied voltage can result in more beads due to an imbalance in forces. Since the Coulomb repulsive force and external electric field cause more polymer solution to leave the needle tip, the droplet at the tip decreases in size [39, 43]. The fiber initiates from the liquid surface inside the needle tip which causes increased bead formation at higher applied

Fabrication of amyloid nanofiber matrices by electrospinning49

voltage. Since the voltage is responsible for stretching and ­acceleration of polymer solution, Krishnappa et  al. observed change in bead morphology with change in applied voltage. They reported formation of spherical/spindle-shaped beads at low voltage, pea pod-shaped beads at midrange applied voltage, and flat, broad fibers at very high voltage. The flat, large diameter fibers were formed due to the extensive increase in the formation of bead which eventually leads to beads merging to form large diameter fibers [55].

Feed rate Polymer solution is fed at a constant rate such that a droplet is formed at the tip of the conducting capillary. Depending on the feed rate, the volume of the polymer available for stretching can vary. Increasing feed rate can increase fiber diameter due to the availability of more polymer at the needle tip [41]. This is limited by the increase in voltage which stretches the polymer, thereby counteracting the increase in diameter due to a larger volume of solution. Tan et al. did not measure significant change in fiber diameter with an increase in feed rate for the same polymer at different concentrations [41]. Higher feed rate can also affect interconnectivity of fibers due to fiber fusion at the target. Larger polymer solution volume takes more time for solvent evaporation. With fixed travel distance, the solution might not dry completely and deposit as a partly dried string of solution. This can affect the overall quality of the fiber mat [24].

3.2.4 Micro- and nanofibers Electrospun fibers primary aim at mimicking the extracellular matrix (ECM) for applications such as wound healing, tissue regeneration, and drug delivery [56]. The morphology, surface properties, material property, and mechanical properties of fibers promote cell adhesion, proliferation, and differentiation [57]. The main component of ECM is collagen that is made from procollagen (diameter: 1.5 nm) secreted from cells. Typical collagen fibril can be between 10 and 300 nm [58]; these form bundles of collagen fibers that are 1–20 μm in diameter [59]. On an average, collagen fibers from skin tissue can measure 56–62 nm in diameter [60]. Inspired by the architectural structure of collagen, electrospun fibers can be characterized as micro- and nanofibers [61, 62]. Electrospun nanofibers can be defined as fibers with a diameter less than a few hundreds of nanometer, whereas fibers with a diameter more than few hundreds of nanometer are called microfibers.

3.2.4.1 Microfibers As discussed in Section 3.2.3, fiber diameter can be modulated by changing the operating parameters during electrospinning. Synthetic polymers, such as poly(l-lactic acid) (PLLA), poly(lactic-co-glycolic acid) (PLGA), and (polyvinylidene fluoride) (PVDF), have been electrospun into microscale fibers for tissue engineering, drug delivery, and filtration [63] applications. Mostly, microfibers can be fabricated using high polymer concentration below the maximum concentration threshold. Yang et al. demonstrated fabrication of fibers with diameter ranging from 150 to 500 nm for low polymer concentration, whereas the fiber diameter ranging from 800 to

50

Artificial Protein and Peptide Nanofibers

3000 nm for higher polymer concentration [64]. At cellular level, microfibers are seen as flat surface by the cells, similar to petri dishes. For drug delivery application, paclitaxel-encapsulated PLGA fibers ranging from 30 nm to 10 μm were successfully fabricated with 90% encapsulation efficiency [65]. Likewise, natural biopolymers have also been electrospun to form microscale fibers from 300 to 1000 nm in diameter [66].

3.2.4.2 Nanofibers Even though microfibers can be used for various applications, there are continued efforts to reduce fiber diameter to nanometer range by modulating the processing parameters or material properties. Synthetic polymers such as Nylon 66, PCL [67], and PVA (polyvinyl alcohol) [68] have been electrospun to produce fiber diameters ranging between 400 and 200 nm. Nylon 66 fiber showed controlled pore size for filtration application [69]. ECM mimicking fiber matrices have been produced from collagen with fiber diameters ranging from 100 to 600 nm [70]. At structural level, collagen in the fiber was denatured during electrospinning, but due to axial alignment of the molecules in a single fiber, the bending modulus was comparable to the native collagen fiber. Naturally occurring fibers such as silk and collagen exhibit high mechanical properties. In order to match those, the diameter of electrospun fibers can be modulated. A study by Yang et al. showed a decrease in bending modulus of the fibers with an increase in fiber diameter. With a fiber diameter of less than 100 nm, nano-effects dominate fiber properties such as high strength, surface energy, and high thermal and electrical conductivity [71]. As per Hall-Petch relationship, the nanofiber strength (τ) can be given as follows:

τ = τ0 +

kτ dα

(3.4)

where τ0 is the strength of the bulk material, kτ is a fitting parameter (material constant), d is a fiber diameter, 0 < d  100 mg/mL) and dissolves in organic solvents such as 2,2,2-­trifluoroethanol (TFE). Due to its globular structure, electrospinning can be a challenge since electrospinning relies on the viscoelasticity of the polymer. Unlike fibrous proteins such as silk fibroin and synthetic polymers, native BSA is not able to withstand the stretching under high electric field during electrospinning. There have been efforts to modify the globular structure of native BSA to make it spinnable. One technique involves unfolding the globular protein to more fibrous protein by reducing the disulfide bonds, preserved both in bovine and human albumin, and altering the electrostatic interactions. Use of organic solvents such as TFE and a combination of TFE/β-mercaptoethanol (β-ME) can unfold the protein and reduce the disulfide bonds to produce fibrous proteins that can retain viscoelasticity for continuous electrospun fiber formation [131]. In contrast, native BSA solution in water makes colloidal lattice arrangement of globular proteins which cannot be electrospun. BSA spinnability can also be improved by blending it with synthetic polymers such as PEO, PLGA, PVA, and PCL. These polymers also improve thermochemical properties of the scaffold. PEO, a water-soluble polymer, has been used previously to improve spinnability of many proteins such as silk and collagen. After cross-linking the protein scaffold, PEO can be easily washed away from the scaffold. BSA and PEO can dissolve within minutes in water if BSA is not chemically cross-linked. Alternatively, albumin can be subjected to thermal hardening to avoid the use of chemicals, such as glutaraldehyde, for cross-linking, which can later affect cell viability. Even though it is a chemical-free process, the slow albumin hardening takes 3 weeks at 37°C, making it a time-consuming process. Electrospinning parameters have been optimized for producing BSA/PEO fibers in the range of few hundred nanometers. The optimized parameters are given in Table 3.4 [132]. BSA electrospun fibers have been used for in vitro and in vivo tissue engineering studies due to their mechanical properties, low toxicity, and minimum foreign body response. Young’s modulus of denatured BSA fibrous mats has been reported to be 1500–2000 MPa with a tensile strength of 30–60 MPa [133]. As a scaffold for blood vessels, albumin tubular structures supported cell adhesion and guided blood vessel formation in  vivo [134]. BSA mats showed properties similar to soft biological fibers such as elastin, with high extensibility and low Young’s Modulus. Electrospun amyloid-­like BSA fiber mats (132–180 nm diameter) also find application in drug delivery due to high porosity, biocompatibility, and controlled drug release property [135]. BSA can also be entrapped as a core in a shell polymer system via coaxial Table 3.4  Optimized parameters for BSA electrospinning. BSA/PEO weight ratio Final polymer concentration Applied voltage Distance between electrode and target Flow rate Humidity Temperature

85:15 8.7% 15 kV 150 mm 0.2 mL/h 30% 22°C

Fabrication of amyloid nanofiber matrices by electrospinning59

Fig. 3.8  Application of BSA electrospun fibers in biosensors.

e­ lectrospinning. BSA entrapped in the water-in-oil emulsion as a core reduced burst release of BSA from PLGA shell electrospun fibers [136]. Conversely, BSA can also be spun as a shell with a synthetic polymer such as PVA as the core at a high voltage of 22 kV that phase-separates BSA and PVA to arrange them as core and shell during electrospinning [137]. Electrospun BSA fibrous matrices can also be used in biosensors. One such example is their use in quartz crystal microbalance, as shown in Fig. 3.8. Kabay et al. used amyloid-like BSA fibers instead of a chemical layer to reduce the complicated steps involved with chemical procedure. BSA, being a self-­ functional protein, was used to immobilize antibodies for protein detection [138].

3.3.4 Other proteins In 2003, Wnek et al. conducted the first study of electrospinning human and bovine fibrinogen dissolved in 1,1,3,3-hexafluropropanol and minimum essential media [139]. Fibrinogen is a globular protein present in blood and participates in wound healing. As a result, it is a good candidate for homeostatic and wound healing bandages. McManus et  al. conducted mechanical characterization of fibrinogen fibers and found a maximum of 0.7 MPa modulus of elasticity for dry electrospun matrices, while glutaraldehyde-treated fibers did not improve the mechanical properties of the fibers [140]. Fibrinogen blended with gelatin has been electrospun for myocardial regeneration. The average fiber diameter ranged from 150 to 300 nm, while the Young’s Modulus was 0.46 MPa [141]. Cardiomyocytes seeded on these composite scaffolds expressed cardiac markers. Elastin is an ECM protein that imparts the characteristic elasticity to the skin. This protein has been electrospun for dermal scaffolds [142] and small vascular grafts [143]. The native elastin blended with polydioxanone can enhance cell response and mechanical properties of the vascular conduits [143] as well as dermal electrospun matrices. Xie et al. have electrospun natural milk protein such as casein and enzyme proteins blended with PEO for immobilizing enzyme molecules [144]. Similar to silk

60

Artificial Protein and Peptide Nanofibers

fibroin, regenerated spider silk was electrospun by Yu et al. to study its morphology, mechanical properties, and effect of solvent on its structure [145]. In 2018, Xiang et al. electrospun recombinant spider silk blend with gelatin and PCL for small vascular grafts [146].

3.4 Conclusion Natural amyloid fibers have high structural and mechanical integrity. The hierarchical architecture of amyloid fibers made from peptide subunits can be mimicked in vitro by electrospinning proteins and small peptide chains. Nonwoven electrospun fiber matrices can be produced by varying the primary and secondary factors affecting the fiber morphology, resulting in beaded, beadless, micro-, or nano-diameter fibers. Optimized parameters can be used to produce amyloid-like fibers with the mechanical properties and diameters similar to natural fibers such as collagen. Self-functional proteins such as silk, collagen, and albumin have been electrospun for applications such as tissue engineering, wound healing, drug delivery, and biosensors. Being naturally derived, these protein matrices are biocompatible and biodegradable which makes them favorable for biomedical applications. In future, structural and self-functional proteins can be used to enhance mechanical and biological properties of synthetic and natural polymers. The fibers electrospun from these composite materials would have better material properties such as tensile strength, biodegradability, and biocompatibility with diameter that could mimic the extracellular matrix. Since these materials have active amino group, various antibodies or drugs can be conjugated to fiber matrices to be used for drug delivery and biosensor applications.

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[143] S.  Sell, M.J.  McClure, C.P.  Barnes, D.C.  Knapp, B.H.  Walpoth, D.G.  Simpson, G.L.  Bowlin, Electrospun polydioxanone–elastin blends: potential for bioresorbable vascular grafts, Biomed. Mater. 1 (2) (2006) 72. [144] J. Xie, Y.-L. Hsieh, Ultra-high surface fibrous membranes from electrospinning of natural proteins: casein and lipase enzyme, J. Mater. Sci. 38 (10) (2003) 2125–2133. [145] Q.  Yu, S.  Xu, H.  Zhang, L.  Gu, Y.  Xu, F.  Ko, Structure–property relationship of regenerated spider silk protein nano/microfibrous scaffold fabricated by electrospinning, J. Biomed. Mater. Res. A 102 (11) (2014) 3828–3837. [146] P.  Xiang, S.-S.  Wang, M.  He, Y.-H.  Han, Z.-H.  Zhou, D.-L.  Chen, M.  Li, L.Q.  Ma, The in vitro and in vivo biocompatibility evaluation of electrospun recombinant spider silk protein/PCL/gelatin for small caliber vascular tissue engineering scaffolds, Colloids Surf. B: Biointerfaces 163 (2018) 19–28.

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Christian Helbing and Klaus D. Jandt Chair of Materials Science, Otto Schott Institute of Materials Research, Friedrich Schiller University, Jena, Germany

4.1 Introduction The assembly of small molecules into larger structures is a crucial mechanism in nature. Several processes begin with the assembly of small molecules into larger ones or structures, such as the formation of crystal or colloid structures, the formation of lipid bilayers, and the phase separation of copolymers or small aptamers that assemble into RNA and DNA. They are responsible for peptide synthesis, which leads, in combination with peptide folding, to protein formation. Proteins are essential for several functions in the human body, such as initiation of cell differentiation or transport of smaller molecules. Albumin, for example, transports substances such as steroids, fatty acids, or hormones, while hemoglobin transports oxygen. The function of other proteins is related to their arrangement into supramolecular structures like fibers or networks. Collagen forms a network, the basic structure of biological tissue, and determines its mechanical properties and morphology. Fibrinogen, the main component of blood coagulation, assembles into a fibrous fibrin network and plays an important role in healing processes and tissue repair [1]. Structures in living nature often show a hierarchical structure assembled by a ­bottom-up formation. The structural principle leads to outstanding properties of natural materials, such as being tough and strong at the same time. Thus, there is a growing interest in the creation of biomolecules-based nanostructures and materials and further to investigate their assembly mechanisms. Understanding self-assembly mechanisms is a requirement to fabricate natural biological templates based on proteins, biopolymers, and designed peptides forming fibrous materials. These fibrous materials that span the nano-to-meso scales have broad applications in nanotechnological and biomedical fields, e.g., scaffolds in three-dimensional (3D) cell cultures and tissue engineering, biosensors, or functional templates for the assembly of other polymeric and inorganic materials. In addition to these desired properties, it is hypothesized that the construction and formation of self-assembled amyloid structures based on peptides and proteins are correlated with various diseases such as Alzheimer’s, type II diabetes, and Parkinson’s syndrome. The deposition of insoluble amyloid assemblies, so-called plaques, in the human body is assumed to be causal for these diseases [2]. For example, misfolding Artificial Protein and Peptide Nanofibers. https://doi.org/10.1016/B978-0-08-102850-6.00004-8 © 2020 Elsevier Ltd. All rights reserved.

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of the proteins amyloid-β and Tau causes plaque formation in the human brain which is toxic for oligodendrocytes [3]. This is one of the main causes of neurodegenerative Alzheimer’s. In order to treat such diseases, it is important to first understand the co-assembly mechanisms of peptides and proteins. A comprehensive understanding of the underlying mechanism will not only allow to identify strategies to reduce or inhibit the cytotoxic properties of the amyloid structures, but also to create biocompatible structures from different proteins with a synergistic effect of their properties. In this chapter, we focus on the co-assembly of peptides and proteins into mostly nanofibrous supramolecular structures. We will outline the different assembly mechanisms like π-π stacking, amphiphilic peptides, β-sheets, and coiled-coil-forming peptides that are used to build heterogeneous structures. Due to a large number of studies on the co-assembly of peptides, only the most important factors and strategies are presented. Furthermore, methods to combine these peptide-based approaches with proteins are shown. To this end, we will give examples for the part of the co-­ assembled protein nanofibers that has not yet been investigated thoroughly. In addition to self-assembly, other techniques for the fabrication of heterogeneous protein fibers that have been developed in recent decades will also be presented. Our aim is to give an overview of this fascinating topic and to demonstrate the great potential of this material class.

4.2 Co-assembled peptide superstructures Peptides are suitable molecules for the preparation of self-assembled structures, and moreover, for identification of interactions responsible for superstructure formation. The flexibility in designing peptide structures allows to tailor the molecule properties and thus their interaction [4–9]. There are 20 natural proteinogenic amino acids common in the synthesis of peptides and proteins in biological cells. Depending on the properties of the R-group, amino acids are categorized as hydrophobic, hydrophilic, charged, and others [9]. The hydrophobic residues include the aliphatic residues (A, I, L, M, V) and aromatic residues (F, W, Y). The hydrophilic, uncharged residues can either be involved in hydrogen bonds via OH (S, T) or CONH (N, Q) groups [9]. The charged residues can be positively (H, K, R) or negatively charged (D, E), and these residues can facilitate the creation of electrostatic interactions that may be conducive to the assembly of peptides into nanofibers [9]. The other amino acids (C, P, G) may allow structural modifications, such as bends in peptide chains or sites for chemical modification which are essential for the formation of tube-like peptide nanofibers by the aromatic-directed assembly.

4.2.1 Co-assembly of short peptides Several groups synthesized simple mono- or dipeptides (peptides consisting of one or two amino acids) to reduce the number of possible interactions or peptides containing protein motifs intended to induce the formation of the superstructure. This peptide flexibility renders it as a viable system for the investigation of co-assembly in order to

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increase versatility and further tailor the properties of the superstructure. Non-covalent interactions play an essential role in biology and especially in superstructure formation. The stabilization of different biomolecules such as DNA or proteins is based on H-bonds or van der Waals forces. The arrangement of such small biomolecules, e.g. peptides, is usually induced by intermolecular interactions, e.g., electrostatic interactions or London dispersion forces leading to molecular stacking. The stacking mechanism is divided into two main categories, π-π or CH-π stacking. The CH-π stacking is characteristic of the interaction of aromatic residues and sugars as observed in lectin with carbohydrates that can inhibit amyloidogenesis [10]. The π-π stackings are typical interactions between aromatic residues in proteins that stabilize their structure or in peptides that lead to the formation of supramolecular structures [11]. In peptides, π-π stacking is generally associated with the formation of amyloids. For example, Reches et al. showed that amyloid peptides with a core sequence of diphenylalanine (FF) can assemble themselves into stable peptide nanotubes [12], and Yang et al. reported that the addition of aromatic groups (carbobenzyloxy and naphthalene) to the N-terminus of some peptides enables them to form stable hydrogels [13]. FF can also be used to form composite peptide superstructures. Adler-Abramovich et al. reported the creation of nanotubes by mixing FF with N-(tert-butoxycarbonyl)modified diphenylalanine (Boc-FF) [14]. Both peptides were co-assembled by aromatic interactions in a tube which was shown by time of flight secondary ion mass spectrometry. Furthermore, the length of the nanotubes and the length distribution depended on the FF/Boc-FF ratio. The increase of the Boc-FF content reduced the tube length and led to a narrow length distribution (Fig.  4.1A–E) [14]. This is explained by the differences in formation kinetics. Heterogeneous tubes had reduced assembly kinetics and an extended lag phase compared to homogeneous tubes because heterogeneous molecules were integrated slower than homogeneous molecules into the tube. This led to a higher amount of formed seeds and a more homogeneous tube growth, showing that the short dipeptide FF is an interesting molecule with an N- and C-termini which can be easily modified and further induce the formation of co-­assembled modified and unmodified FF into supramolecular structures. The flexibility of the co-assembly based on peptides with aromatic residues was shown with the creation of hydrogels based on fluorenylmethoxycarbonyl-FF (Fmoc-FF) and Fmocarginine-glycine-aspartic acid (Fmoc-RGD) [15, 16]. RGD is one of the smallest moieties which forms self-assembled structures. The Fmoc-RGD and Fmoc-FF structures were formed by β-sheets and both structures were interlocked via π-π stacking of the Fmoc-groups (Fig. 4.1F–I). Based on the integration of the RGD-sequence, these hydrogels showed good biocompatibility [15, 16]. Tena-Solsona et  al. used various tetrapeptides based on F, aspartic acid (D) and lysine (K) to produce hydrogels based on co-assembled nanofibers [17]. They showed a pH-sensitive coaggregation of the tetrapeptides Z-FDFD with Z-FKFK and Z-KFKF, where Z denotes a benzyloxycarbonyl group. Both peptides coaggregated at a pH of about 7, whereas the peptides alone were soluble. The co-assembly took place through the electrostatic interaction between oppositely charged groups in the peptides and through π-π stacking. The resulting secondary structure within the nanofibers was dependent on the amino acid sequence used [17]. While Z-FDFD with Z-FKFK formed

Fig. 4.1  See legend on opposite page

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β-sheets which were also stable at pH changes, the secondary structure of Z-FDFD with Z-KFKF changed with respect to the pH value. This showed a better packing of the first combination compared to the second. Furthermore, they investigated the tendency of the tetrapeptide family consisting of F and D to first self-assemble to hydrogels and then co-assemble with the Aβ1–40 fragment of the β-amyloid (Aβ) peptide [18]. The primary sequence of the tetrapeptides showed a major influence on the supramolecular formation behavior [18]. The resulting hydrophobic and stacking interactions were crucial for hydrogel formation. In addition, a dependence between co-assembly and amino acid sequences was observed. ZFDFD and ZFDDF showed a co-assembly with Aβ, evidenced by fiber morphology after fibril formation, while ZDFFD and ZDFDF did not interact with Aβ. For ZFFDD and ZDDFF, no information about the co-assembly has been obtained. Interestingly, after mixing with ZFDFD, ZFDDF, and ZDDFF, a disappearance of the characteristic amyloid β-sheet band in the secondary structure of Aβ was observed. ZFDFD showed the strongest effect, indicating a remarkable preference for interaction [18]. In the other combinations, the secondary structure remained unaffected.

4.2.2 Co-assembly of amphiphilic peptides Another approach to create self-assembled peptide nanofibers is based on the peptide derivatives, for instance, peptide amphiphile (PA). PA consists of oligo-peptides that are modified with a hydrophobic alkyl tail to form molecules with distinctly hydrophobic and hydrophilic ends, similar to lipids. This kind of PAs tends to self-organize in aqueous solution, so those hydrophobic domains are buried away from water while hydrophilic regions are exposed to water. In this way, the PA can self-assemble into various types of micelles, vesicles, and of course, nanofibers and scaffolds [4–6]. In 2001, Fig. 4.1  Co-assembly of FF and Boc-FF peptides. (A) Illustration of a co-assembled nanotube. FF building blocks and their chemical structure are represented in red, and Boc-FF building blocks and their chemical structure are represented in blue. SEM images of the peptide assemblies for different FF:Boc-FF molar ratios, (B) FF alone, (C) 20:1, (D) 10:1, (E) 5:1, are shown. Scale bar = 2 μm. (F) The chemical structures of the hydrogel building blocks: Fmoc-FF and Fmoc-RGD. (G) The mixture of Fmoc-FF and Fmoc-RGD selfassembles into a translucent hydrogel at 37°C. (H) AFM height image of the hydrogel which shows an overlapping mesh of nanofibers, with bundles and entanglements. (I) The proposed supramolecular model demonstrates the formation of the 3 nm fibrils and their further lateral assembly into larger ribbons. RGD sequences are presented on the fiber surface enhancing their accessibility and bio-availability. Panels (A–E) reprinted with permission from L. Adler-Abramovich, P. Marco, Z.A. Arnon, R.C.G. Creasey, T.C.T. Michaels, A. Levin, D.J. Scurr, C. J. Roberts, T.P.J. Knowles, S.J.B. Tendler, E. Gazit, Controlling the physical dimensions of peptide nanotubes by supramolecular polymer coassembly. ACS Nano 10 (8) (2016) 7436–7442, Copyright 2016, American Chemical Society and panels (F–I) from M. Zhou, A.M. Smith, A.K. Das, N.W. Hodson, R.F. Collins, R.V. Ulijn, J.E. Gough, Self-assembled peptide-based hydrogels as scaffolds for anchorage-dependent cells. Biomaterials 30(13) (2009) 2523–30, Copyright 2009, Elsevier.

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Fig. 4.2  Design of PAs for articular cartilage regeneration. Chemical structure of (A) TGFbinding PA and (B) nonbioactive filler PA. (C) Illustration of co-assembly of the TGF-binding PA and the filler PA showing binding epitopes exposed on the surface of the nanofiber. (D) The catanionic mixture of qCPT-Sup35 (mixing ratio 1:3) results in the almost exclusive formation of tubular structures in 1:1 MeCN/H2O. The tubular size measured from cryo-TEM imaging is 123 ± 28 nm. Panels (A–C) reprinted with permission from R.N. Shah, N.A. Shah, M.M. Del Rosario Lim, C. Hsieh, G. Nuber, S.I. Stupp, Supramolecular design of self-assembling nanofibers for cartilage regeneration. Proc. Natl. Acad. Sci. U. S. A. 107(8) (2010) 3293–3298 and panel (D) from Y.-A. Lin, A.G. Cheetham, P. Zhang, Y.-C. Ou, Y. Li, G. Liu, D. Hermida-Merino, I.W. Hamley, H. Cui, Multiwalled nanotubes formed by catanionic mixtures of drug amphiphiles. ACS Nano 8(12) (2014) 12690–12700 (https://pubs.acs.org/doi/10.1021/nn505688b). Copyright 2014, American Chemical Society.

Stupp’s group first reported the preparation of PA with mono-alkyl chains attached via the N-termini [19]. Based on their structure, PAs are suitable peptides for the formation of composite structures by functionalization of the hydrophilic part, and thus, for the adjustment of the fiber properties (Fig. 4.2A–C) [9, 20, 21]. Niece et al. reported the co-assembly of PAs functionalized with either the RGD sequence known for cell attachment, the IKVAV or the YIGSR sequence, both derived from laminin and known to interact with mammalian neurons [22]. In addition to their amphiphilic character, the modified PAs also had different charges, which further facilitated co-assembly through electrostatic interactions. The formation of nanofibers, which occurred over a broad

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concentration range ( 10% by weight), cannot be explained by the hydrophobic interactions between the alkyl tails; if so, micelles should also occur [22]. A second possible driving force is the β-sheet hydrogen bonding between the peptide sequences in the outer area, as observed in Fourier transform infrared spectroscopy [22]. In addition, the Stupp group synthesized “reverse” PA with a free N-terminus, which enables easier functionalization and makes it biologically active [23]. Again, the addition of different charged groups, one PA with a triple lysine and the other with a triple glutamic acid, at the end of these PAs led to a co-assembly with β-sheet formation within the fibers. This was also the proof for co-assembly because β-sheets did not appear for the assembly of one type of PAs or two PAs with a similar charge [23]. The heating of individual PA systems to 85°C resulted in disassembly into individual molecules and cooling, in turn, led to the formation of individual composite nanofibers. In contrast, circular dichroism spectroscopy of co-assembled fibers at different temperatures showed a temperature-dependent increase in peak intensity for β-sheets up to a temperature of 80°C. This demonstrates that these fibers were not in an equilibrium state at room temperature. It is noteworthy that the fibers at 85°C had a higher melting/disassembly temperature than expected due to the hydrogen bonding network (40–45°C) or the aliphatic end of the PAs (60°C), which shows the high stability of these supramolecular structures [23]. A similar approach was followed by Lin et al. who produced drug-loaded multiwall amyloid nanotubes by mixing two oppositely charged PAs [24]. They used the β-sheet forming part of the yeast prion Sup35 modified with the anticancer drug camptothecin (CPT). Furthermore, the drug-containing peptides were functionalized with either two lysine or two glutamic acids to achieve amphiphilicity and the opposite charge status. Both peptides were able to co-assemble, but the resulting morphology depended on the amount of CPT molecules added. Four CPT molecules bound to the peptide led to the formation of nanotubes (Fig. 4.2D), while only two CPT led to the formation of nanofibers. Based on the wall thickness of 25 nm for the uniform nanotubes, the presence of several double layers in this structure has been suggested [24].

4.2.3 Co-assembly of peptides based on protein motifs Another design approach which allows peptides to be used as building blocks for self-assembly is by utilizing protein motifs, the basic conformational units of naturally existing proteins, β-sheets and turns, α-helices, and coiled-coils. As mentioned above, β-sheets are well-known for their ability to assemble into long fibrous structures, as is seen in amyloid linked diseases, such as Alzheimer’s and Parkinson’s diseases. Some peptide sequences were thought to be critical for the formation of amyloid nanofibers and cause of diseases. The most studied protein motifs are the peptide sequences of the Aβ peptide with 39–43 amino acids, and many fibrous structures have been created by controlled assembly [9, 25–27]. Ray et al. reported that a water-soluble tripeptide (VIA) that originated from the Aβ(40–42) can self-assemble to form amyloid-like nanofibers [28]. One very thoroughly investigated peptide is the Aβ which is connected to Alzheimer dementia, in particular the co-assembly of the two derivatives, Aβ40 and Aβ42, which

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typically produced and formed aggregates with neurotoxic properties [29]. To gain more information about the arrangement of both peptides in aggregates and if co-­ assembly is relevant for Alzheimer’s disease, Baldassarre et al. used isotope infrared spectroscopy to display the aggregation. Co-assembled β-sheets containing aggregates were formed by random accumulation of both peptides [30]. Also, the co-assembly of short Aβ-fragments, as well as their influence on the full-length peptide, was studied with the aim to understand the co-assembly mechanism and to reduce toxicity of Aβ plaques [31–33]. The combination of a rhodamine fluorophore-labeled Aβ(17–22) with a normal Aβ(16–22) peptide showed that peptide fibers are extended by the addition of monomeric peptides to the fiber ends [32]. Suzuki et  al. showed that the co-assembly of Aβ with designed peptides based on sequences of Aβ leads to less toxic amyloid-like fibrils [33]. Further, also mutants of the N-termini (A2V and A2T) were able to reduce or increase the Aβ formation [34]. Adding C-terminal fragments of Aβ42 to the full-length peptide led to a neuroprotective effect by lacking the cytotoxicity function of the full-length aggregates [35]. To obtain more information on the formation of natural amyloid structures with heterogeneous character, researchers are also concentrating on other amyloid-­forming peptides. The exact underlying molecular formation mechanism and the connection with diseases are still unknown. One factor is cross-seeding, as the study of the compilation of Aβ40 and hIAPP shows. The addition of hIAPP to Aβ40 accelerated amyloid Aβ40 formation from 9 to 3.5 h after mixing in a molar ratio of 1:1 [36]. The influence of the cross-seeding will be explained in the following section Prions. A closer look into the intermolecular interactions allowed the co-assembly of oppositely charged short amyloid inspired peptides, AIP-1 (Ac-EFFAAE-Am) and AIP-2 (AcKFFAAK-Am) [37]. It was shown that both peptides interacted based on the hydrophobicity of the aromatic residues and the hydrophilicity of the -Lys and -Glu residues. Amyloid-like fibers are not only an undesirable state of proteins in the human body since they are the main factor for amyloid diseases, but they are also an amazing platform for the production of functional nanomaterials [4, 5]. There are also several amyloid structures in nature with functional roles [4, 38]. Examples include biofilms of bacteria, the eggshell of insects like the silkmoths, or the catalytic scaffold of the melanin synthesis of mammals [38]. Inspired by the rules provided by nature, scientists started to create new soft materials that are harmless to humans by designing amino acid sequences which mimic the amyloid structure. Zhang et al. first demonstrated the use of β-sheets for the design of new fibrous materials in the early 1990s [39]. Their work indicated that it was possible to create a fibrous peptide structure containing βsheets by creating a pattern of peptides with hydrophobic and complementary charged amino acids, such as the pattern of (AEAEAKAK)2. Similar peptide patterns, such as (RADA)4, (RARADADA)2, and (KLDL)3, have been used to create peptide nanofibers [40–42]. Other β-structured peptides, such as β16 (GGALEAKLAALEAKLA) [43], biotinylated β16 [44], and P11-2 (QQRFQWQFEQQ) [45], can also form peptide nanofibers by self-assembly. Inspired by this work, researchers used the co-assembly of functionalized and unfunctionalized amphiphilic β-sheet moieties to optimize the properties of the resulting structures. Mixing the EAK16II peptide with the EAKIIH6, in which the C-termini is functionalized with hexahistidine, led to the exposure of

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hexahistidine at the fiber surface [46–48]. This, in particular, allows an antibody reaction, which makes the formed structures suitable for biological applications [46, 47]. Similar approaches have been shown for co-assembled hydrogels based on the peptide RADA16, which had RGDS and other biofunctional motifs, e.g., from collagen [49, 50], laminin [51], fibrin [52], or fibronectin [53]. Another system used for this approach was the Q11 peptide with and without similar modifications [54, 55]. The formation of hydrogels from Q11 which are functionalized with either RGD or the IKVAV laminin sequence is a highlight for a synergistic effect of functional motifs. Based on the RGD sequence, the resulting hydrogels showed improved cell binding, growth, and proliferation, and the laminin sequence was responsible for little to no impairment of gel viscoelasticity. The approaches described above aimed to integrate different peptides into a fibrous structure. Another interesting way to introduce the assembly of different homogeneous fiber structures is the use of enantiomeric peptides. Similar to the stereocomplex formation known from different polymers, peptides with different chirality can also be co-assembled. Swanekamp et al. reported the assembly of l- and d-(FKFE)2 peptides into amyloid fibers based on all-l or all-d peptides. The mixing of both peptides led to the formation of a new fiber type in the form of “ribbed β-sheets” consisting of alternating l- and d-(FKFE)2 peptides [56]. An alternative to using β-sheets as a basis for a fibrous nanostructure are α-helices, and one example of a well-studied and well-defined α-helical motif is the coiled-coil [3, 57]. A coiled-coil region consists of two to seven α-helices which stabilize themselves by hydrophobic and electrostatic interactions between assembled α-helices. The α-helices are formed by a sequence of seven amino acids which turn two times. An extensively studied design of a fibrous coiled-coil-based system has been explored by the Woolfson group using the “sticky ends” assembly (Fig. 4.3A–C) [57–60]. There, the α-helices contain positively and negatively charged amino acids facilitating the addition of further α-helices at the ends. Woolfson et al. used α-helices forming peptide sequences where the fifth and the seventh position amino acid was either a positively charged Lys or a negatively charged Glu [60]. This modification results in the electrostatically driven self-assembly of heterodimeric coiled-coils, whereby the “sticky ends” introduced the longitudinal extension, as shown in Fig.  4.3A–C [58]. The resulting bundles had a diameter of around 50 nm and lengths longer than 10 μm. A variation in the peptide amino acid sequence and the resulting co-assembly allows tuning the fiber morphology from linear to waved, kinked, and branched. Based on the well-understood assembly mechanism, other peptides, such as harboring biotin and FLAG-peptide tags, have been easily included in the fabricated heterogeneous fibers. Further, the introduction of fluorophores into these coiled-coil fibers allowed to investigate the growth and late stages of assembly kinetics [61, 62]. Also other groups designed coiled-coil forming peptides-based coiled-coil sequences of natural proteins. Nakaji-Hirabayashi et al. used the type II keratin K14 [63]. They co-assembled K14 with a K14 peptide modified with the laminin globular domain 3 [63]. This resulted in co-assembled fibers with a K14 core surrounded by laminin domains, which led to hydrogel formation by entanglement. The laminin domains located on the hydrogel surface provide good adhesion of stem/progenitor cells, and thus, good biocompatibility [63].

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Fig. 4.3  (A) Design principles of the self-assembling-fiber peptides: complementary charges in companion peptides direct the formation of staggered, parallel, and codirectional heterodimers, and the resulting “sticky ends” are also complementary and support longitudinal association into extended fibers. (B) Complementary charged pairs on the outer surfaces of the coiled-coil protofibrils support protofibril-protofibril interactions, fiber assembly, and thickening. (C) TEM micrographs of negative stained third-generation fibers; scale bars 50 nm (left) and 2 μm (right inset). (D) Influence of the coiled-coil forming peptide VW01 on the amyloid assembly of VW19. Panels (A–C) reprinted with permission from D. Papapostolou, A.M. Smith, E.D.T. Atkins, S.J. Oliver, M.G. Ryadnov, L.C. Serpell, D.N. Woolfson, Engineering nanoscale order into a designed protein fiber. Proc. Natl. Acad. Sci. U. S. A. 104(26) (2007) 10853–10858. Copyright 2007, National Academy of Sciences, U.S.A. and panel (D) adapted with permission from E. Brandenburg, H. von Berlepsch, J. Leiterer, F. Emmerling, B. Koksch, Formation of alpha-helical nanofibers by mixing beta-structured and alpha-helical coiled coil peptides. Biomacromolecules 13 (11) (2012) 3542–3551. Copyright 2012, American Chemical Society.

Another work showed that coiled-coil peptides have the potential to inhibit cytotoxic β-sheet formation of amyloids. Brandenburg et al. co-assembled two different peptides, VW1 and VW19, based on heptad repeat units which formed α-helical and coiled-coil structures [64]. VW1 formed coiled-coil structures, whereas VW19 was modified with Val at several positions which led to amyloid-like β-sheet fibers. Coassembly of VW1 and VW19 in an equimolar ratio or a higher content of VW1 led to the formation of co-assembled coiled-coil fibers and suppressed the amyloid formation (Fig.  4.3D). Switching the concentration to an excessive amount of VW19 initiated again the formation of amyloid structures [65].

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This section shows that co-assembled peptides have a broad field of application and their formation mechanism is well understood, which might help to develop new therapies for amyloid or plaque-related diseases such as Alzheimer’s, Parkinson’s, or Creutzfeldt-Jakob. Further details and examples of co-assembled peptide nanofibers are provided elsewhere [3–5, 57, 66, 67].

4.2.4 Co-assembled peptide-protein superstructures The creation of functional materials based on natural or synthetic proteins, instead of peptides, is often hindered by the challenges of controlling protein conformation and the resulting nanoscale assembly. Thus, the ability to generate hierarchical materials by exploiting the unique structural and functional properties remains a challenging goal [6]. One approach to solve this problem is the combination of protein and peptide assembly. Peptides can be designed to interact with specific proteins, promote their conformational changes, and control their composition, allowing adaptation of function and structure. Inostroza-Brito et al. synthesized an elastin-like, thermorescent protein based on a tropoelastin recurrent motif Val-Pro-Gly-X-Gly, wherein X, besides proline, is any amino acid [6]. The mixing of this protein with an amphiphilic peptide led to the formation of nanofibrous multilayer structures. The assembly was driven by electrostatic interactions due to the different isoelectric points (pIs) of the proteins (about 3) and the peptides (about 10) [6]. Changing the pH of the assembly solution, and thus the electrostatic interactions, allowed to switch the resulting structure between a robust and a weak tube as well as a closed membrane or a collapsed structure. They used time-of-flight-secondary ion mass spectrometry to show the anisotropic character of the resulting structures with a high proportion of ELP5 in the outer surface and PAK3 in the inner surface. The formation of composite nanofibrous structures was not mentioned. Another way to integrate proteins into a peptide-based amyloid fiber is to functionalize the protein with an additional peptide that can be easily integrated. Hudalla et al. functionalized a GFP protein with two different amyloid-forming peptides, the βtail and the Q11 peptide (Fig. 4.4) [7]. While Q11 showed a rapid β-sheet and fibril formation, βtail had a slow transition from α-helix to β-sheet and thus a much slower aggregation. Based on the slow aggregation of βtail, proteins modified with this peptide have been integrated into self-organizing Q11 nanofibers. By varying the protein to peptide ratio, it is possible to precisely control protein integration. The co-assembly was not limited to Q11; also, other β-sheet-forming peptides such as KFE8, RADA16, and HK-Q11 co-assemble with β-tail-labeled proteins [7]. Integration into coaggregated fibers has been demonstrated by immunogold labeling and fluorescence spectroscopy based on the fluorescence properties of GFP. Circular dichroism spectroscopy showed the formation of mixed β-sheet structures as shown for the heterogeneous assembly of designed peptide pairs with differently charged regions [43]. A simpler way to integrate a protein into an amyloid peptide was demonstrated by Rigdley et  al. who showed that an amyloid-containing hydrophobic template peptide from gliadin in combination with α-helical proteins (α-­ casein, α-­lactalbumin, amylase, hemoglobin, insulin, and myoglobin) can form

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Fig. 4.4  A schematic representation of engineered fusion proteins having a β-sheet fibrillizing domain integrating into Q11 nanofibers. Reprinted with permission from G.A. Hudalla, T. Sun, J.Z. Gasiorowski, H. Han, Y.F. Tian, A.S. Chong, J.H. Collier, Gradated assembly of multiple proteins into supramolecular nanomaterials. Nat. Mater. 13(8) (2014) 829–36. Copyright 2014, Nature Publishing Group.

­heterogeneous fibers [68]. The hydrophobic peptide initiated a conformational transition from α-helix to β-sheet in the protein by interacting with the hydrophobic moments within the α-helix. The resulting fibers were in the micro range and had a hierarchical structure [68]. Protofibrils were assembled to fibrils in the nanoscale and further on to fibers. The resulting fiber morphology such as width, pitch length, twist, and bands depended on the added protein, as shown in several studies [68–70]. The combination of serum albumin with a peptide of the Aβ, the fragment of the toxic region, showed that other peptides can also influence the formation of amyloid structures [71]. Fibril formation was induced by the addition of the Aβ peptide to the protein. A formation mechanism was not proposed, but a conformational change of the albumin by the peptide is possible, similar to the previously described mechanism. The literature has shown that albumin forms amyloid fibers under partial unfolding conditions [72–74]. A similar phenomenon was observed for phenylalanine fibers, which induced fibrillation of other globular proteins such as lysozyme, serum albumin, insulin, myoglobin, and cytochrome c [75]. It is assumed that the attachment of proteins to phenylalanine fibers was facilitated by electrostatic interactions. The soluble proteins were present in their native structure and had charged groups exposed to the solvent. These charged groups interacted with charged groups located on the outside of phenylalanine fibers [75]. However, it is not clear whether the proteins reacted in their native state or whether there is a conformational change from a native to an aggregation-prone state. In addition to phenylalanine, other metabolites (not peptide)

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associated with various diseases are also believed to induce protein aggregation. An interesting article on this topic has been published by Sade et al. [8]

4.3 Co-assembled protein superstructures Proteins, in comparison to peptides, cannot be simply designed to adapt their functions and interactions. Yet, they are fully biocompatible, readily available, and have defined functions determined by their primary, secondary, and tertiary structures. The co-assembly of protein allows combining the different protein functions and properties, such as biocompatibility in one material, without complicated design steps. Nevertheless, the formation of the fibrous protein-based superstructure, and in particular the co-assembly of different proteins, is more demanding. In this section, we will focus on three formation methods, i.e., electrospinning, extrusion, and self-assembly mechanism.

4.3.1 Electrospinning of co-assembled protein superstructures The electrospinning process was initially used to produce polymer fibers with a diameter in the nm to μm range. The literature describes two different types of electrospinning, wet and melt spinning. The principles are based on electrical forces which are responsible for fiber formation. The high voltage used for this process, about 30 kV, first leads to the melting of the polymer in the spinneret during melt spinning, and then to a charged polymer melt/solution. When the charge inside the spinneret and the drop on the open side is high enough to overcome the surface tension, a cone is formed between the spinneret and the collector. This creates a constant liquid jet between the spinneret and collector in which the solvent evaporates (wet-spinning) and the whipping occurs before the stream reaches the collector. By adjusting the voltage, the distance between spinneret and collector, and the concentration in the solution, it is possible to control the fiber diameter. It was shown that it is possible to use more than 100 different polymers for electrospinning and further that it is possible to use natural matrix forming proteins, such as collagen [76], elastin, and fibrinogen, or the most common example for natural protein fibers, spider silk. That proteins form fibers in a natural state is not a prerequisite for electrospinning, and thus, globular molecules such as albumin, hemoglobin, or myoglobin were used [77, 78]. Since the introduction of electrospinning for the production of scaffolds in tissue engineering on the basis of biopolymers and/or proteins, researchers tried to mimic the changing composition of the ECM [77, 79–81]. Protein-based electrospun heterogeneous superstructures are typically produced from mixtures of a synthetic and often biodegradable polymer and a protein [77, 82–88]. Here, we concentrate on the production of fibrous superstructures based only on the co-assembly of protein blends. Among the first researchers were Boland et al. and later Buttafoco et al., who described the electrospinning of nanofibers from a collagen-elastin mixture [79, 81]. They observed homogeneous fibers with diameters in the range of 200 to 800 nm, depending

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on the weight ratio of collagen to elastin in the spinning solution [79, 81]. A direct differentiation between different proteins in the fibers was not possible with AFM [79], nor with SEM [81]. Both came to the conclusion that the composition of the fibers is equal to the distribution of proteins in the mixed solution, but no evidence for this was presented. To stabilize the electrospun fibers, a cross-linking step with glutaraldehyde or carbodiimide (EDC)/N-hydroxysuccinimide (NHS) was required; otherwise, the fibers dissolved immediately in water [81]. The cross-linking step indicates that no covalent bonds were formed during the spinning process. Furthermore, Buttafoco et al. suggested that the arrangement of proteins was based on hydrophobic interactions produced by the used NaCl [81]. However, it is likely that the used high voltage and thus the high temperature, as well as the used solvents 1,1,1,3,3,3-hexafluoro-2-­propanol (HFIP) or 10 mM hydrochloric acid, influenced the secondary structure of the proteins, which can lead to a partial unfolding. These changes may be responsible for the arrangement of proteins in the fibers, as HFIP induces the formation of α-­helical structures or the exposure of hidden docking regions. Several studies focused on the electrospinning of the well-investigated silk fibroin. Co-assembled fibrous structures via electrospinning were reported for elastin [89] and collagen [90]. The fibrous structures from the blends of silk fibrous and elastin were stabilized by cross-linking with genipin. Genipin introduced a conformational transition from random coils to β-sheets in the secondary structure of silk fibroin. Further, an increase in β-sheets was also observed for elastin after adding the cross-linker. It is supposed that the conformational changes are needed to allow the formation of secondary amine linkages between the amino groups of silk fibroin or elastin and the ester groups of genipin [89]. In contrast, for the formation of collagen-blended silk fibroin fibers, a cross-linking agent was not needed. These co-assembled superstructures were produced from a solution containing both proteins [90]. However, a formation mechanism was neither described nor assumed. Based on the formation conditions, an unfolding and change in the secondary structure can be expected, similar to the previous description for heterogeneous collagen fibers. For both studies, the co-assembly was not directly confirmed, but implicitly assumed. After addition of the second protein, identification of different as well as single protein molecules inside the fibers was not possible. For silk fibroin-elastin co-assembly structures, the formation of scaffolds was described for 100% silk fibroin which changes to a fibrillary structure for 50% silk fibroin and 50% elastin [89]. The diameters of silk fibroin collagen fibers decreased depending on the silk fibroin:collagen ratio from 2.15 ± 0.13 µm for 100:0 to 1.03 ± 0.04 µm for 85:15 [90]. Both examples show that the composition influences the resulting structure or fiber. The co-assembly of three different proteins was described by Ravichandran et al. [91], who aimed to produce an ECM mimicking fibrous scaffold based on hemoglobin, gelatin, and fibrinogen. Hemoglobin and gelatine were dissolved in TFE and mixed with fibrinogen dissolved in HFIP. The resulting fibrous structures were cross-linked with phytic acid. In this case, the cross-linking was based on bonding phytic acid anions to cations of the proteins [91]. The cross-linking influenced the fiber diameter, i.e., 290 ± 130 nm for 1% phytic acid and 337 ± 113 nm for 5% phytic acid. The effect of the cross-linking to the biocompatibility and cell proliferation was not observed. Yet, the authors assumed that the high oxygen binding capability of hemoglobin in

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the fibers was responsible for the regeneration of ischemic myocardium, and thus, the scaffold is suitable for myocardial tissue engineering [91]. In conclusion, electrospinning is a suitable tool to co-assemble different proteins into fibrous structures, which properties can be controlled by varying the composition. However, commonly, a cross-linker is needed to stabilize and bind the different molecules in the resulting fibrous structures. Also, most of the mentioned examples resulted in fibers with a diameter in the micrometer range and only few are in the nanometer range, which we then call nanofibrous structures.

4.3.2 Extrusion of co-assembled protein superstructures Another technique to create fibrous structures of (co)-assembled proteins is the extrusion process which is related to electrospinning. Instead of an electrical field which aligns the proteins to a fiber, a protein containing solution flows through a needle onto a substrate or into a coagulation bath. The resulting shear stress, appearing when the protein solution passes the needle, leads to the uniaxial alignment of the protein molecules inside the fibrous structure. Underwood et al. reported one of the first studies about the co-assembly of fibrinogen and fibronectin into “cables” by extrusion [92]. A fibrinogen and fibronectin containing buffer solution which further contains urea was pressed through a spinneret into a coagulation bath with a pH of 2. The added urea led to a partial unfolding of fibronectin [93] as well as fibrinogen [94], which allowed the aggregation of the proteins inside the coagulation bath. The low pH of the coagulation bath further induced the interaction of both proteins. Several studies showed that a low pH of 2 led also to the self-assembly of proteins, including fibrinogen, to fibrous structures [72, 95–98]. It is expected that the conformational changes in the proteins are responsible for the self-assembly mechanism. Thus, it is challenging to determine which factors of this extrusion technique are responsible for the co-assembly. In particular, Raoufi et al. reported the formation of fibronectin fibers by extrusion without a coagulation bath or urea [99]. In contrast to Underwood et al., they used a nanoporous anodic aluminum oxide membrane (pore size: 20 and 200 nm) for fiber formation. They showed that shear stress also induces a partial unfolding of the fibronectin, leading to intermolecular interactions resulting in a lateral assembly. The formation of co-assembled fibers consisting of collagen and fibronectin, collagen and elastin, as well myosin and actin was reported (Fig. 4.5A–C) [100]. Varying the protein concentration and the membrane pore size, it was possible to control the diameter of the resulting fibrous structures in the range between 15 and 151 nm. Formed single fibers assemble further to larger fiber bundles in micrometer scale directly after extrusion. Although they proved the existence of collagen and fibronectin in such a fiber bundle by immunofluorescence, it was not possible to determine the presence and distribution of both proteins in one fiber [100]. Yet, the existence of a heterogeneous fiber is quite likely: The proved heterogenous character of the fiber bundles showed that either homogenous fibers of two different proteins can interact with each other and form heterogeneous bundles or the bundle was formed by heterogeneous fibers. A rapid formation of fiber bundles after extrusion should be introduced by strong interactions between different single fibers. This means that either different homogenous

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Fig. 4.5  SEM images of extruded protein composite nanofibers and polysaccharide nanofibers: (A) fiber bundle from a collagen-fibronectin blend, (B) fiber assembly from a collagen-elastin blend, (C) nanofibrous assembly of an actin-myosin blend. (D) Incorporation of fluorescently labeled fibronectin into the silk fibroin fiber, at increasing doping concentrations, greater amounts of fibronectin are incorporated into the alloy fibers. Scale bar is 15 μm. Panels (A–C) reprinted with permission from M. Raoufi, N. Aslankoohi, C. Mollenhauer, H. Boehm, J.P. Spatz, D. Bruggemann, Template-assisted extrusion of biopolymer nanofibers under physiological conditions. Integr. Biol. (Camb.) 8(10) (2016) 1059–1066. Copyright 2016, The Royal Society of Chemistry and panel (D) from M.M. Jacobsen, D. Li, N. Gyune Rim, D. Backman, M.L. Smith, J.Y. Wong, Silk-fibronectin protein alloy fibres support cell adhesion and viability as a high strength, matrix fibre analogue. Sci. Rep. 7 (2017) 45653. Copyright 2017, Nature Publishing Group.

fibers or heterogeneous fibers were interacting with each other. We assume that both proteins interact with each other; otherwise, heterogeneous fiber bundles would not be formed. Further, since both proteins were randomly distributed in the feed solution and then pressed through pores, it is very unlikely that only one type of protein enters the same pore. However, to confirm this assumption, an analysis of the unknown inner fiber structure is needed. The co-assembly of silk fibroin and fibronectin to heterogeneous fibers was shown by Jacobsen et al. These heterogeneous fibers were prepared by wet-spinning combined with a coagulation bath (70% methanol) [101]. Jacobsen et  al. assumed that a combination of different interactions led to the co-assembly [101]. Besides ionic and hydrophobic interactions, fibronectin should form intermolecular β-sheets with silk fibroin chains which results in a physical cross-linking. Further, an entropic entanglement of the fibronectin molecule is assumed to bind within the silk fibroin lattice. The heterogeneous character of the fibers was proved with fluorescence imaging where the fibronectin was labeled with Alexa Fluor 633 prior to the fiber formation.

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The average volumetric fluorescence increased with increasing fibronectin content (Fig. 4.5D). Further, the properties of silk fibroin fibrous structures have been tuned by incorporating the fibronectin. Interestingly, the mechanical integrity of the fibers was not influenced by the fibronectin due to the supposed integration mechanism of physical cross-linking. Furthermore, fibronectin improved the biocompatibility of the fibrous structures as shown by improved cell attachment and viability of fibroblasts, endothelial, and vascular smooth muscle cells [101]. Although the fiber diameters were in the range of micrometers, instead of nanometers, it is a good example of the synergistic effect of co-assembled proteins.

4.3.3 Self-assembly of co-assembled protein superstructures The fiber formation in the previously described techniques was induced by aligning the molecules with an external force, e.g., electronic field or shear stress. Often, this resulted in fibers with diameters in the range of several micrometers. Thus, the nanoscale advantage of a large surface-to-volume ratio, which is advantageous for ­surface-induced processes, is not given for the fibers and they are not really “nano”. Here, the self-assembly process is suitable to produce nanofibrous structures. There are several reports in the literature dealing with the coaggregation of different ­disease-relevant proteins into amyloid structures, in particular with the coaggregation of prions (infectious proteins). This coaggregation is similar to the co-assembly process and leads to the formation of amyloid fibers. An intensively investigated prion is the α-synuclein which relates to the early onset of Parkinson’s. α-Synuclein is known to form amyloid structures under defined conditions [102–104]. Typical for the formation of amyloid structures is the lag phase where the proteins unfold and form nuclei for the fiber formation. It is known that adding seeds reduces or eliminates this lag phase. Wood et  al. observed that the addition of homo-mutant (A53T) α-synuclein seeds to a wild type of α-synuclein accelerated the amyloid fiber formation [102]. This shows that the pathogenic A53T mutant acts as a heterogeneous seed and introduces the amyloid formation of the soluble wild-type α-synuclein which is the first step of co-assembly to heterogeneous structures. This result was unexpected due to the structural differences in the mutant and wild-type fibers [102]. Besides a mutant version of the α-synuclein, a truncated version of this protein induces the amyloid fiber formation [104–106]. Liu et al. simulated an incomplete degradation of the α-synuclein and tested the influence of truncated α-synuclein on the full-length α-synuclein [104]. Thioflavin T-binding studies showed that the truncated proteins accelerate the full-length α-synuclein protofibril formation. The existence of a co-assembled fiber was demonstrated by the use of HIS-tagged full-length α-synuclein. HIS-tagged full-length and untagged truncated proteins were detected by gel filtration chromatography. However, a formation mechanism and the associated intermolecular interactions were not proposed. Horvath et al. showed that prions of different diseases can also crosstalk and reported coaggregation between α-synuclein and the islet amyloid polypeptide (IAPP) associated with type-2 diabetes [107]. Both proteins showed differences in their aggregation kinetics. IAPP assembled within several minutes to amyloids, while α-synuclein needed hours. Horvath et al. assumed the existence of ­heterogeneous

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fibers due to their different influences as seeds by accelerating and inhibiting protein aggregation [107]. The authors also claimed that additional experiments, such as immunogold staining, are necessary for a true proof of coaggregation [108]. A detailed overview of further studies regarding the coaggregation of prions was given by Bondarev et al. [109]. They subdivided the coaggregation into four classes, titration, sequestration, axial, and lateral coaggregation. Titration occurs when soluble proteins interact with already formed fibers, or in this case, amyloids via specific binding sites. If the interactions are not based on specific binding sites, it is called sequestration. Further, axial coaggregation describes the molecular stacking of different proteins along the fiber axis of one (amyloid) fiber. In such a coaggregated fiber, the proteins can theoretically be arranged statistically, alternately, with a gradient or as blocks [109, 110]. When different types of homogeneous amyloid fibers, consisting of only one protein, coaggregate through interactions between their side surfaces, this is referred to as lateral coaggregation. Based on the experimental data of different studies, a distinction between class three and four is often not possible. Several studies only reported a coaggregation or a cross-seeding of proteins without determining the molecule arrangement. Especially, the verification of axial coaggregation is one of the most challenging tasks [109]. However, for some prions, the axial coaggregation was shown for Sup35 with Rnq1 [111–115], csgA with csgB [116, 117], and Rip1 with Rip3 [118]. After cross-seeding experiments of Sup35 fibers with Rnq1 seeds, the Rnp1 was detected by immunogold staining inside the formed fibers [111]. Other studies indirectly proved axial coaggregation by cell uptake experiments and resulting overexpression of marker molecules [112, 113, 115]. The co-assembly for this combination was driven by multiple binding sites in both proteins. It is assumed that blocking of one crucial binding site might be sufficient to alter the resulting interactions [114]. For csgA and csgB, the axial co-assembly was also indirectly proven [119]. Both proteins are part of the curli fiber formation which is involved in cell adhesion and aggregation as well as biofilm formation [120]. Adding csgB to soluble csgA induces amyloid formation. CsgB, which is amyloidogenic, starts to rapidly form oligomers. Based on five repeating units of csgB which are similar to five repeating units of csgA, csgB is a perfect nucleator for csgA and starts the aggregation [116]. Hammer et al. mentioned that nature separates in this case the nucleation and elongation properties into two separate molecules to guide the place and time of interaction [116]. The axial-aggregation of Rip1 and Rip3 is correlated with cytokine TNF-induced necrosis. Their axial arrangement was proven by HIS-tagged truncated Rip1 and Rip3 versions. Further, this method was used to determine the shortest sequences which are needed for interaction [118]. Inspired by the works on coaggregation regarding prions and Aβ oligomers, Dubey et  al. aimed to expand the current knowledge by revealing the underlying principles leading to coaggregation of different globular proteins [121]. As a suitable model system, they choose four different proteins, bovine serum albumin, lysozyme, insulin, and cytochrome C, which are well-investigated and form individual amyloids. Heating up these individual molecules up to 70°C, near to their melting temperatures, introduced the amyloid formation with their typical formation kinetics including a lag phase. Mixing now these protein monomers at 70°C led to a dramatically decreased lag phase and a rapid coaggregation, independent from the mixture

Fig. 4.6  Coaggregation studies of mixed monomers of different globular proteins in PBS at ∼ 70°C. (A) Coaggregation of albumin (BSA) and insulin; (B) Coaggregation of insulin and lysozyme; (C) Coaggregation of BSA and lysozyme; (D) Coaggregation of BSA, lysozyme, and insulin. (E) AFM topography image of a single albumin (HSA)-hemoglobin (HGB) fiber. The white line represents the TERS measurement area and the two points where the spectra in (F) were taken. (F) Two representative TERS spectra with and without characteristic HGB bands. (G) Schematic depiction of hPNF labeling with AuNPs. Representative STEM image of hPNFs immunolabeled with antibodies conjugated with AuNPs. Particles assigned to HSA are marked orange, whereas nanoparticle assigned to HGB are marked purple. Panels (A–D) reprinted with permission from K. Dubey, B.G. Anand, M.K. Temgire, K. Kar, Evidence of rapid coaggregation of globular proteins during amyloid formation. Biochemistry 53(51) (2014) 8001–8004 (https://pubs.acs.org/doi/10.1021/bi501333q). Copyright 2014, American Chemical Society and panels (E–G) from C. Helbing, T. Deckert-Gaudig, I. Firkowska-Boden, G. Wei, V. Deckert, K.D. Jandt, Protein handshake on the nanoscale: how albumin and hemoglobin self-assemble into nanohybrid fibers. ACS Nano 12(2) (2018) 1211–1219. Copyright 2018, American Chemical Society.

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(Fig. 4.6A–D). Reducing the temperature prevents coaggregation as well as amyloid formation. Interestingly, bioinformatical analysis identified no significant similarities in their sequences [121]. This contradicts early reports, which claimed that protein sequences with more than 70% identity favor aggregation and, further, less than 30–40% identity are not facilitating aggregation [122]. This reveals the important role of partial unfolding in the co-assembly of proteins. Dubay et al. assumed that the coaggregation and also cross-seeding depend more on the intermediate partial unfolded species and less on their sequence identities [121]. Based on that, they draw two interesting conclusions. First, there is a net gain of additional intermolecular interactions during aggregation, and second, that there are some conserved regions inside the protein’s sequences which hinder their individual self-assembly [121]. A similar conclusion was drawn by Oki et al., who investigated the coaggregation of β-lactoglobulin and lysozyme [123]. Both proteins coaggregated at a temperature of 70°C and above. Besides the already mentioned partial unfolding and hydrophobic interaction, these proteins were also coaggregated by electrostatic interaction due to the different pIs of 5.1 for β-lactoglobulin and 10.7 for lysozyme. Oki et al. identified that hydrophobic interactions, electrostatic affinity, and unfolding of proteins are involved in coaggregation and nucleation by adding different additives to eliminate one of these interactions [123]. Moreover, they concluded that nucleation is triggered by a slight partial unfolding [123]. However, direct proof of a coaggregated fiber was not provided. A new approach to prove the existence of co-assembled fibers was reported by Helbing et  al. who investigated the ethanol-induced self-assembly of human serum albumin and hemoglobin fibers [124]. They used tip-enhanced Raman spectroscopy (TERS) to identify the hemoglobin molecule inside the co-assembled fibers, based on their heme-group. TERS allows performing Raman spectroscopy with a nm resolution. (Fig. 4.6E and F) These findings were complementary proved by immunogold labeling. (Fig. 4.6G) Interestingly, both methods revealed the very high albumin to hemoglobin ratio. This high ratio has been explained by the fast self-assembly kinetics of albumin compared to hemoglobin. Interestingly, a cross-seeding as described by Dubay et  al. was not observed. Helbing et  al. assumed that the ethanol-induced unfolding exposed corresponding regions, amino acid sequences in both proteins with similar properties, which are necessary to self-co-assemble into fibers [122, 124]. The interaction between these sequences was called protein “handshake” [124]. Co-assembly of proteins is not only associated with diseases, but is also found in nature, e.g., in the heterogeneous structure of ECM. One of the first experiments, which “almost” describes a co-self-assembly process of two different proteins, was reported by Underwood et al. in 1999 [125]; almost, because they drew protein cables containing fibrinogen and fibronectin from a solution with a pH between 4.0 and 4.5. The lowering of the pH led to fine precipitation, which aggregated through gentle movement and formed fine protein fibers. These precipitating fibers were slowly drawn upwards from the solution which led to the arrangement of small fibers in the drawn direction to bigger bundles, the protein “cables.” It is assumed that isoelectric forces led to precipitation, and thus, to the interaction of proteins and fine fibers. The diameter of the fiber bundle

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formed is in the range of 200 μm. However, such a bundle consists of several small fibers with a diameter of about 1 μm, which are still not in the nano range. To conclude this section, based on the complex structures of proteins, identifying the mechanism of co-assembly is far more challenging than for small designed peptides. However, co-assembled proteins are not always related to diseases, for example, the ECM. It is still a major goal of several research projects to mimic this unique scaffold, as it may lay the foundation for artificially created organs.

4.4 Conclusion The co-assembly of peptides and proteins was initially reported in relation to diseases such as Alzheimer’s, Parkinson’s, type II diabetes, or Creutzfeld Jacob. However, the observation that such structures are not necessarily harmful to the human body opens up an interesting field of research that offers the opportunity to develop new functional materials with excellent and controllable properties. The well-studied field of self-assembled peptides allows easy adaptation of properties by using well-established ways to design the peptide structure. We have shown in this chapter that the different assembly mechanisms such as π-π stacking, amphiphilic peptides, β-sheet folding, and coiled-coil-forming peptides are well-understood. The combination of known assembly mechanisms and well-described design rules makes them a suitable material for heterogeneous self-assembly. It allows extending the possible functions of these materials. The control of the peptide design makes it possible to further optimize the interactions between peptides and proteins. This enables the use of natural protein properties in more synthetic peptide structures. At the end, we presented different techniques for the production of co-assembled protein fibers. Especially electrospinning and extrusion are suitable for this purpose. We presented examples showing that the different protein properties in a fiber can be synergistically mixed. However, there are still some challenges to be overcome, such as the production of stable composite fibers without cross-linking or the production of fibrous structures on the nanoscale. These can be overcome, for example, by using the self-­ assembly mechanism. Several studies, in particular on the coaggregation of prions, have shown the formation of co-assembled nanofibers at the nanoscale. In addition, only a few studies reported on the co-assembly of non-disease-related proteins into nanofibers. Thus, they are suitable as building blocks for mimicking natural hierarchical structures. However, the greatest challenges in this area are the verification of the heterogeneous composition of nanofibers and the identification of the underlying co-assembly mechanisms. In several studies, the composition of the heterogeneous fibers was only assumed and not proven, which is due to a lack of possible characterization methods with the resulting resolution on the nanoscale. Further studies focused on the investigation of cross-seeding to answer the question of how and why proteins can induce nucleation and further control nanofiber formation kinetics. The responsible mechanisms are not well-understood, which means that work still needs to be done. This is a prerequisite for the development of new therapies for amyloid-related diseases.

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[106] W. Li, N. West, E. Colla, O. Pletnikova, J.C. Troncoso, L. Marsh, T.M. Dawson, P. Jäkälä, T.  Hartmann, D.L.  Price, M.K.  Lee, Aggregation promoting C-terminal truncation of α-synuclein is a normal cellular process and is enhanced by the familial Parkinson's disease-linked mutations, Proc. Natl. Acad. Sci. U. S. A. 102 (6) (2005) 2162–2167. [107] I. Horvath, P. Wittung-Stafshede, Cross-talk between amyloidogenic proteins in type-2 diabetes and Parkinson's disease, Proc. Natl. Acad. Sci. U. S. A. 113 (44) (2016) 12473–12477. [108] I. Horvath, S. Rocha, P. Wittung-Stafshede, In vitro analysis of alpha-synuclein amyloid formation and cross-reactivity, Meth. Mol. Biol. 1779 (2018) 73–83. [109] S.A. Bondarev, K.S. Antonets, A.V. Kajava, A.A. Nizhnikov, G.A. Zhouravleva, Protein co-aggregation related to amyloids: methods of investigation, diversity, and classification, Int. J. Mol. Sci. 19 (8) (2018) 2292. [110] A.J. Dear, T.C.T. Michaels, T.P.J. Knowles, Dynamics of heteromolecular filament formation, J. Chem. Phys. 145 (17) (2016) 175101. [111] Y.A. Vitrenko, E.O. Gracheva, J.E. Richmond, S.W. Liebman, Visualization of aggregation of the Rnq1 prion domain and cross-seeding interactions with Sup35NM, J. Biol. Chem. 282 (3) (2007) 1779–1787. [112] J. Sharma, S.W. Liebman, Exploring the basis of [PIN +] variant differences in [PSI +] induction, J. Mol. Biol. 425 (17) (2013) 3046–3059. [113] I.L. Derkatch, S.M. Uptain, T.F. Outeiro, R. Krishnan, S.L. Lindquist, S.W. Liebman, Effects of Q/N-rich, polyQ, and non-polyQ amyloids on the de novo formation of the [PSI+] prion in yeast and aggregation of Sup35 in vitro, Proc. Natl. Acad. Sci. U. S. A. 101 (35) (2004) 12934–12939. [114] K.M.  Keefer, K.C.  Stein, H.L.  True, Heterologous prion-forming proteins interact to cross-seed aggregation in Saccharomyces cerevisiae, Sci. Rep. 7 (1) (2017) 5853. [115] F.  Arslan, J.Y.  Hong, V.  Kanneganti, S.-K.  Park, S.W.  Liebman, Heterologous aggregates promote De Novo prion appearance via more than one mechanism, PLoS Genet. 11 (1) (2015) e1004814. [116] N.D. Hammer, J.C. Schmidt, M.R. Chapman, The curli nucleator protein, CsgB, contains an amyloidogenic domain that directs CsgA polymerization, Proc. Natl. Acad. Sci. U. S. A. 104 (30) (2007) 12494–12499. [117] Q. Shu, S.L. Crick, J.S. Pinkner, B. Ford, S.J. Hultgren, C. Frieden, E. The, coli CsgB nucleator of curli assembles to beta-sheet oligomers that alter the CsgA fibrillization mechanism, Proc. Natl. Acad. Sci. U. S. A. 109 (17) (2012) 6502–6507. [118] J. Li, T. McQuade, A.B. Siemer, J. Napetschnig, K. Moriwaki, Y.S. Hsiao, E. Damko, D.  Moquin, T.  Walz, A.  McDermott, F.K.  Chan, H.  Wu, The RIP1/RIP3 necrosome forms a functional amyloid signaling complex required for programmed necrosis, Cell 150 (2) (2012) 339–350. [119] L.P. Blanco, M.L. Evans, D.R. Smith, M.P. Badtke, M.R. Chapman, Diversity, biogenesis and function of microbial amyloids, Trends Microbiol. 20 (2) (2012) 66–73. [120] M.M. Barnhart, M.R. Chapman, Curli biogenesis and function, Annu. Rev. Microbiol. 60 (2006) 131–147. [121] K. Dubey, B.G. Anand, M.K. Temgire, K. Kar, Evidence of rapid coaggregation of globular proteins during amyloid formation, Biochemistry 53 (51) (2014) 8001–8004. [122] C.F.  Wright, S.A.  Teichmann, J.  Clarke, C.M.  Dobson, The importance of sequence diversity in the aggregation and evolution of proteins, Nature 438 (2005) 878. [123] S. Oki, K. Iwashita, M. Kimura, H. Kano, K. Shiraki, Mechanism of co-aggregation in a protein mixture with small additives, Int. J. Biol. Macromol. 107 (2018) 1428–1437.

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Characterization techniques of protein and peptide nanofibers: Self-assembly kinetics

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Gang Weia,b a College of Chemistry and Chemical Engineering, Qingdao University, Qingdao, PR China, b Faculty of Production Engineering, University of Bremen, Bremen, Germany

5.1 Introduction The self-assembly and formation of both natural and artificial protein and peptide nanofibers are related to biological functions and diseases of human body [1–4]. With the development of protein bioengineering, peptide synthesis, and nanotechnology, a lot of functional protein and peptide molecules with ability to self-assemble into nanofibers/nanofibrils have been designed and synthesized for the applications in tissue engineering, biomedical engineering, nanodevices, sensors, energy storage materials, and environmental materials [5–7]. It is well-known that unfolded proteins and peptides are easy to form one-­ dimensional (1D) nanofibers under a broad range of conditions. However, most of the native folded proteins should undergo a pretreating process like unfolding and hydrolysis to form molecular aggregation to nanofibers. For instance, in order to promote the conformation transition and self-assembly of protein and peptide molecules, some physical experimental conditions such as pH, ionic strength, temperature, and solvents can be adjusted [8–11]. The addition of metal ions, polymers, and enzymes into the protein and peptide systems has exhibited great effects on the self-assembly kinetics of nanofibers [12–14]. In addition, recent studies on the self-assembly of functional peptides to nanofibers have indicated that the motif design of peptide is one of the helpful ways to synthesize peptide nanofiber-based biomaterials for specific applications [15, 16]. To help the optimal design and synthesis of functional nanofibers, it is necessary to understand the molecule aggregation and self-assembly kinetics [17]. Usually, the kinetic studies of nanofiber self-assembly provide high sensitivity and high resolution in detecting the changes of molecular aggregation. For instance, spectroscopy techniques such as circular dichroism (CD) [18], ThT fluorescence [19], Fouriertransform infrared spectroscopy (FTIR) [20], and others have been well-established to monitor the aggregation of molecules over a series of factors (time, temperature, pH, concentration), and the microscopy assays such as transmission electron microscopy (TEM) [21], atomic force microscopy (AFM) [22], and other super-resolution microscopy (SRM) [23] have been used to observe the length scale of nanofibers with high resolution. In addition, the developments of theoretical computer simulations and Artificial Protein and Peptide Nanofibers. https://doi.org/10.1016/B978-0-08-102850-6.00005-X © 2020 Elsevier Ltd. All rights reserved.

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optimized analytical methods and models have provided the possibility to study the self-­assembly of nanofibers at both atomic and molecular levels [24, 25]. The aim of this chapter is to present an overview on various characterization techniques for the self-assembly kinetics of protein and peptide nanofibers. The kinetic triggering for molecular self-assembly to nanofibers by adjusting some experimental parameters will be introduced, and then the self-assembly kinetic characterizations of nanofibers with the spectroscopy, microscopy, crystallography, and other analytical techniques will be demonstrated and discussed in detail. It is expected that this work will be very helpful for readers to understand the kinetic characterizations of protein and peptide self-assembly towards various nanostructures from 1D to 3D in one way, and in another way, to design and develop novel nanofiber-based nanomaterials for advanced applications.

5.2 Kinetic triggering for molecular self-assembly Protein and peptide molecules can be mediated to self-assemble to form 1D amyloid nanofibers/nanofibrils. It is not easy for the native folded proteins to aggregate and form nanofibers, which should be first activated by unfolding and hydrolysis before the self-assembly process [5]. However, the unfolded proteins and peptides exhibited excellent ability to self-assemble into nanofibers through a fast nucleation process via applying various experimental conditions, as shown in Fig. 5.1. The adjusting of these experimental factors is helpful for triggering the molecular aggregation and self-­ assembly kinetics of protein and peptide nanofibers. One of the typical strategies is to trigger the self-assembly kinetics of protein and peptide nanofibers by adjusting the solution properties such as the pH [26–28], ionic strength [29], and temperature [30], or using organic solvents [10, 31, 32]. For instance, Kobayashi et al. investigated the effects of various pH on the aggregation of amyloid-β (Aβ)1–42 protein [26]. It has been found that when the solution pH was smaller than 5 or larger than 9.5, the protein molecules only changed their conformation and no nanofibers formed. However, when the pH was adjusted to 6–8, the protein

Fig. 5.1  Triggering of the self-assembly kinetics of protein and peptide nanofibers by adjusting various experimental parameters.

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aggregation and formation of nanofibers were observed. In another study, Dehsorkhi et al. studied the effect of pH on the self-assembly of a collagen-derived peptide amphiphile, C16-KTTKS, and demonstrated that, with the increasing of pH, the peptide self-assembled into different structures from micelles (pH 2) to flat tape (pH 3), twisted nanofibers (pH 4), and then flat tape (pH 7) [27]. Therefore, the triggering of pH on the self-assembly kinetics of proteins and peptides exhibits high potential for understanding the self-assembly mechanism of nanofibers. The ionic strength of solution also shows great effects on the molecular aggregation and the formation of protein and peptide nanofibers. In a typical study, Abelein and coworkers characterized the explicit effect of ionic strength on the microscopic aggregation rates of Aβ1–40 and found that physiological ionic strength could accelerate the self-assembly kinetics of proteins by promoting surface-catalyzed secondary nucleation reactions [29]. It is clear that the salt decreased the free-energy barrier for the folding of Aβ1–40 to the mature stable state, promoting the creation of mature nanofibrils. Usually, high temperature (for instance 90°C) is favorable for the molecular conformation transition and the formation of amyloid nanofibers. Previously, Loveday et al. studied the effect of temperature (from 75°C to 120°C) on the fibril formation kinetics, fibril structure, and rheological properties of β-lactoglobulin nanofibrils [30]. Besides the solution properties, the type of solutions can also affect the self-assembly kinetics of proteins and peptides to nanofibers. It has been found that the organic solvents and natural surfactants revealed high effects on the formation of protein and peptide nanofibers [10, 31]. The self-assembly kinetics of protein and peptide nanofibers can be triggered by adding various biological (such as enzyme) [13, 33], organic [34], and inorganic (metal ions and nanoparticles) [12, 35, 36] mediators to induce/inhibit the formation of nanofibers. For example, Xu’s group has developed enzyme-based synthesis strategies for creating a series of small d-amino acids and d-peptides nanofibers for biomedical applications [13, 33]. It is clear that enzymes could be applied as a useful trigger for understanding the self-assembly kinetics and tuning the cytotoxicity of nanofibers against different cancer cells. Recently, Li and coworkers demonstrated the preparation of peptide nanofibers by using the polyoxometalate-driven self-assembly of short peptide molecules [34]. Due to the electrostatic interactions between cationic peptide and polyanions, the ionic self-assembly resulted in the formation of multivalent peptide nanofibers with improved antimicrobial performances. Besides biological and organic promoters, inorganic metal ions and nanoparticles have been proved to induce the nanofiber formation. For instance, Zn2 + and Cu2 + could be useful for creating amyloid β protein and peptide nanofibers [12, 35], and silver and gold nanoparticles with a diameter of 20 nm accelerated the aggregation kinetics of two peptides, NNFGAIL and GNNQQNY, greatly [36]. In addition, the solid-liquid [37–39] and air-water [40] interfaces have potential for affecting the self-assembly kinetics of nanofibers. Reichert et al. investigated the adsorption of fibrinogen on the highly oriented pyrolytic graphite (HOPG)-water interface and found that the surface structure of graphite guided the formation and topotactical orientation of fibrinogen nanofibers [37]. Recently, Liao and coworkers demonstrated the self-assembly and formation of peptide (NapFFKYp) nanofibers and nanosheets on the substrates of mica, HOPG, and polystyrene film [39].

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They  found that all the substrates could promote the formation of rod-like peptide nanofibers with substrate-dependent thickness. Besides the solid-liquid interface, the air-­water interface can affect the formation of protein nanofibrils. For instance, Campioni et al. investigated the aggregation of α-synuclein at the air-water interface by using multi-techniques [40]. It has been found that α-synuclein aggregated readily into nanofibrils that adsorbed on the interface. However, the aggregation and formation of nanofibrils were reduced greatly when the air-water interface was replaced by a solid-liquid interface. Therefore, it can be concluded that both solid-liquid and air-water interfaces are one of the key triggers for studying the self-assembly kinetics of protein and peptide nanofibers. It should be noted that some biological interfaces such as the cell membranes and self-assembled bio-membranes could also be used for guiding the formation of amyloid protein nanofibers [41]. Besides, the external stimulus such as ultrasonication [28, 42], mechanical stress [43], and pressure [44] can affect the self-assembly kinetics of protein and peptide nanofibers. For instance, Chatani et al. investigated the ultrasonication-based production of monodispersed amyloid β2-microglobulin nanofibrils with a well-defined molecular size, which were very useful for characterizing the structure and dynamics of amyloid nanofibrils [42]. In this case, the use of ultrasonication breaks down the preformed fibrils to shorter, uniform nanofibrils with minimum free energy, which will be helpful to achieve the direct measurements of nuclear magnetic resonance (NMR) spectra of amyloid nanofibrils as the shorter nanofibrils exhibited more sharp NMR peaks than the longer fibrils. However, sometimes the ultrasonication is also responsible for the disassembly of peptide nanofibers [28]. In another study, Macchi and coworkers studied the effect of different mechanical stresses on the self-assembly of flucagon to nanofibrils [43]. It has been observed that the low-stress- and high-stressprepared protein nanofibrils exhibited different kinetic profiles, secondary structures, and morphologies. Mishra and Winter further suggested that cold denaturation and mechanical pressure are effective to understand disaggregation and refolding mechanisms of amyloid fibrils [44].

5.3 Characterizations of self-assembly kinetics of nanofibers/nanofibrils The self-assembly kinetics of protein and peptide nanofibers can be characterized with many techniques, including spectroscopy, microscopy, crystallography, and advanced analytical techniques.

5.3.1 Spectroscopy analysis CD spectroscopy is a powerful technique for understanding various types of secondary structures of biomacromolecules [45] and has been widely used for the characterization of the self-assembly kinetics of protein and peptide nanofibers/nanofibrils previously [46–49]. CD spectroscopy can reveal the conformation states of protein

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and peptide molecules under specific experimental conditions due to its advantage of real-time monitoring of molecular conformations in aqueous solution. For example, Paramonov et al. studied the roles of hydrogen bonding and amphiphilic packing in the self-assembly of peptide amphiphiles (C16-GGGGGGGERGDS) with spectroscopy and microscopy techniques [46]. The CD measurements indicated that the alanine (A)-mutated PAs showed different conformation states, as shown in Fig. 5.2A. The A mutants of PA at the first to fourth position of G (closed to the ­hydrophobic core)

Fig. 5.2  Spectroscopic study on the self-assembly kinetics of nanofibers: (A) CD, (B) fluorescence, (C) FTIR, and (D) tip-enhanced Raman spectroscopy (TERS). Images (A–D) are reproduced by permission from (A) S.E. Paramonov, H.W. Jun, J.D. Hartgerink, Self-assembly of peptide-amphiphile nanofibers: the roles of hydrogen bonding and amphiphilic packing, J. Am. Chem. Soc. 128 (2006) 7291–7298, Copyright 2006, (B) J. Kardos, A. Micsonai, H. Pal-Gabor, E. Petrik, L. Graf, J. Kovacs, Y.H. Lee, H. Naiki, Y. Goto, Reversible heat-induced dissociation of beta(2)-microglobulin amyloid fibrils, Biochemistry 50 (2011) 3211–3220, Copyright 2011, (C) Q. Lu, H.S. Zhu, C.C. Zhang, F. Zhang, B. Zhang, D.L. Kaplan, Silk self-assembly mechanisms and control from thermodynamics to kinetics, Biomacromolecules 13 (2012) 826–832, Copyright 2012, and (D) C. Helbing, T. Deckert-Gaudig, I. Firkowska-Boden, G. Wei, V. Deckert, K.D. Jandt, Protein handshake on the nanoscale: how albumin and hemoglobin self-assemble into nanohybrid fibers, ACS Nano 12 (2018) 1211–1219, Copyright 2018, American Chemical Society.

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caused the f­ormation of β-sheet structure, while the mutants at the fifth to seventh position (distal from the core) of G resulted in polyproline type II conformation. In another case, Bartolini et al. investigated the kinetics of Aβ1–42 peptide s­ elf-aggregation by CD spectroscopy [47]. Under optimized conditions, the reproducible CD kinetic study indicated that the tested Aβ1–42 peptide reveals a three-step conformation transition from unordered α-helix to increasing β-sheet and prevailing β-sheet secondary structure. Their kinetic analysis of peptide aggregation makes it clear for the ­self-assembly mechanism of peptides, and at the same time, bring insight into the inhibiting mechanism of peptide. Recently, Ye et al. investigated the effects of peptide hydrolysis for fibrillation kinetics and fibril morphology with far-UV CD spectroscopy and other techniques [49]. The obtained CD data indicated that the minimum of the CD spectra of created fibrils shifted from 208 nm in the starting peptide solution to about 215–220 nm after self-assembly for 3 days, proving the formation of β-sheet structure with increasing content. Fluorescence spectroscopy is a useful technique to characterize the time-dependent aggregation and self-assembly of protein and peptide molecules. To show the fluorescent signals of protein or peptide molecules in the process of self-assembly, fluorescent probes such as thioflavin-T (ThT) and Nile Red (NR) could be added to bind to the self-assembled/aggregated structures to result in enhanced emission signals [49–52]. It is well-known that both ThT and NR can bind with the amyloid form of protein and peptide aggregates, but not with their monomers, and therefore, are valuable for amyloid detection in vitro and in vivo. For instance, Kardos and coworkers investigated the reversible heat-induced dissociation of β2-microglobulin, α-synuclein, and K3 amyloid fibrils with CD and ThT fluorescence spectroscopy [50], as shown in Fig. 5.2B. It was found that the polymerized amyloid fibrils could be dissociated to monomers very quickly under incubation at 99°C through the ThT fluorescence measurements. In addition, the dissociation of fibrils is a reversible and dynamic process to reach the equilibrium between fibrils and monomers quickly. In another study, Jacob et al. investigated the self-assembly of Fmoc-protected Aβ40–42 peptides to βrich nanofibrils with spectroscopy and microscopy techniques [52], in which the ThT binding proved the formation of amyloid peptide (Fmoc-Aβ40–42-Val) structure and the NR binding delayed the gel formation without interfering the fibril formation. Therefore, the fluorescence spectroscopy provides a simple and direct evidence for the formation of amyloid nanofibers. Two important vibrational spectroscopy techniques, FTIR [53, 54] and Raman [55, 56], can provide dynamical and structural analysis of self-assembled protein and peptide nanofibers. It is well-known that FTIR spectroscopy provides light absorption due to the changes of molecular dipole moments and Raman spectroscopy reveals the inelastic scattering information of molecules caused by bond polarization, and therefore, these two techniques could show complementary information for analyzing the molecular structure. Lu and coworkers characterized silk nanofibers formed by mimicking the natural spinning process with FTIR and morphological techniques [53]. It is possible to distinguish the organized but not β-sheet structure (peak of silk I) and the stable β-sheet structure (silk II) with FTIR, as shown in Fig. 5.2C. The obtained FTIR data indicated that hydrophilic interactions, concentration, charge, and temperature

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control the formation of silk nanofibers. Therefore, the self-assembly of silk to nanofibers is not only a thermodynamic process (conformation transition to β-sheets), but also a kinetic process that involves in solution properties (such as hydrophilic interactions, concentration, charge, and others). In another case, Guo and Wang provided spectroscopic (CD and FTIR) evidence on the self-assembly and aggregation of Aβ fragments (Aβ1–42, Aβ1–25, and Aβ33–42) to different structures [54]. By using Raman spectroscopy, Flynn et al. characterized the structural features of α-synuclein amyloid fibrils that formed at various pH, ionic strength, and molecular sequences [55]. Raman spectra of samples provided clear differences in the amide-I, amide-III, and the fingerprint regions of amyloid fibrils, indicating that the self-­ assembly of protein to fibrils is kinetic-controlled. Recently, Helbing et al. investigated the molecular co-assembly of albumin and hemoglobin to hybrid nanofibers in ethanol [56]. The synthesized hybrid protein nanofibers were characterized by tip-­enhanced Raman spectroscopy (TERS) with precisely selected points. When there is hemoglobin in the hybrid nanofibers, the TERS spectrum (spectrum 1) revealed iron vibrations at 1355 (Fe2 +) and 1378 (Fe3 +) cm− 1, while the TERS spectrum (spectrum 2) of nanofibers without hemoglobin did not show the iron-related peaks, as shown in Fig. 5.2D. Therefore, TERS is highly useful to collect spatially resolved Raman spectra over several nm to determine the presence of hemoglobin in the created hybrid nanofibers. NMR spectroscopy can also be utilized for characterizing the structure and ­self-assembly kinetics of nanofibers. The structural features of molecules or molecular aggregates, including the secondary structure, intra- or intermolecular distance, and ­dihedral angles, can be determined by NMR. For instance, Wälti and coworkers ­determined the 3D structure of a disease-relevant polymorph of Aβ1–42 amyloid fibrils at atomic resolution with the solid-state NMR technique [57]. By combining with the data of mass-per-length measurements from electron microscopy, they found that the 3D structure is composed of two molecules per fibril layer by forming a d­ ouble-horseshoe-like cross-β-sheet entity. Their finding is highly helpful to know the self-assembly mechanism of Aβ1–42 protein at the atomic level and further to develop potent anti-amyloid diseases’ (AD) drugs and AD diagnostic biomarkers. In another case, van der Wel investigated the structure of GNNQQNY amyloid fibrils by magic angle spinning (MAS) NMR [58] and found that the conformers were three predominantly β-sheet in structure, but with a highly localized distortion in one of them. This study provided insights into the distinction between fibril growth and crystal formation. To improve the sensitivity of MAS NMR experiments, dynamic nuclear polarization (DNP) NMR has been developed for studying the intermolecular structure of amyloid fibrils [59, 60]. It is clear that using DNP technique could short the data acquisition time from days to hours and improve the signal-to-noise ratios of the obtained NMR spectra significantly. The DNP NMR made it possible to define not only the arrangement of peptide β-strands to β-sheets, but also the β-sheet interfaces within each protofilament [60], leading to the understanding of the self-assembly mechanism of nanofibrils at the atomic and molecular levels. Besides the above techniques, a traditional spectrophotometric UV-vis technique has been applied by Cirulis et al. for studying the kinetics and morphology of peptide self-assembly [61]. By simple analysis of the obtained spectrophotometric data, they

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were able to describe the self-assembly kinetics of elastin-like peptide in  vitro and in vivo. In addition, by binding probe dye like Congo Red (CR) to the self-assembled amyloids, it is possible to understand the molecular aggregation mechanism through UV-vis measurement of CR absorption [52].

5.3.2 Microscopy analysis The self-assembly kinetics of protein and peptide molecules can be understood by characterizing the nanofiber length with several microscopy techniques including TEM, AFM, Stochastic optical reconstruction microscopy (STORM), and confocal fluorescence microscopy (CM). TEM and AFM have been the most popular techniques for the characterizations of both the morphology and properties of self-assembled protein and peptide nanofibers [21, 24, 62–64]. By statistical analysis of the obtained nanofiber length and height scales by various kinetic factor controls, it is possible to obtain the potential aggregation and self-assembly mechanism of molecules. For instance, previously, Dong and coworkers utilized ice cryo-TEM to observe the formation of self-assembled multidomain peptide (MDP) nanofibers and demonstrated the effects of molecular frustration on the conformation and final self-assembled nanostructures [21]. Marshall et al. observed the self-assembly of Sup35 yeast prion fragment (GNNQQNY) formed with various assembly periods (0–192 h) with TEM and found the structural transition from nanofibers to crystals [62]. It is clear that TEM technique provides high resolution to see the nanostructure of nanofiber, but the original nanofibers should be stained or ice-frozen for TEM sample preparation. AFM provides quick and convenient characterization to the structural properties of self-assembled protein and peptide nanofibers, which can be directly deposited onto flat solid substrates (Si wafer and mica) for imaging in air or liquid. The unique nanofiber structures with helix and ribbon properties can be seen clearly due to the high lateral resolution of AFM (about 0.1 nm). For instance, Marini et al. investigated the self-assembly of peptide (FKFEFKFE) in water over time and observed the formation of left-handed helical nanofibers by AFM [63]. In another case, Adamcik and coworkers demonstrated their statistical analysis of AFM images of heat-denatured β-lactoglobulin fibrils and found that the formed fibrils have a multistranded helical shape with twisted ribbon-like structures [24]. This study provided a general model for amyloid fibril assembly, which is highly helpful for understanding the growth kinetics and aggregation mechanism of protein and peptide nanofibers. Besides TEM and AFM, novel super-resolution microscopy techniques such as STORM [23, 65] and CM [66, 67] have been used to study the self-assembly and growth kinetics of amyloid peptide nanofibrils. Previously, Pinotsi et  al. observed directly the heterogeneous growth kinetics of α-synuclein nanofibrils via two-color STORM with a resolution of better than 20 nm [23]. They were able to distinguish the structures of single nanofibrils formed at different stages by using different fluorescent labeling protein seeds, revealing high capabilities for studying the growth kinetics of nanofibers at the molecular level. In a recent study, Beun et al. elucidated the growth dynamics, exchange kinetics, and assembly mechanism of silk-like protein

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block nanofibrils by using AFM and two-color STORM techniques [65]. It was found that the red-labeled protein can self-assemble into nanofibrils and then the addition of green protein monomers promotes the green extensions to grow on the red nanofibrils, as shown in the scheme and corresponding STORM images of Fig. 5.3A and B. Meanwhile, the length and height analyses on formed nanofibrils with AFM snapshots indicate the growth kinetics of nanofibrils by adjusting the protein concentration and assembly period (Fig. 5.3C–E). The in situ study of protein self-assembly by combining AFM and STORM mediates the understanding of protein aggregation mechanism and opens perspectives for creating functional biomaterials through hierarchical self-assembly. Esbjörner and coworkers performed direct observations on the aggregation and self-assembly kinetics of Aβ1–40 and Aβ1–42 proteins with fluorescence CM technique [66]. The obtained noninvasive fluorescence lifetime images of nanofibers indicate that the fluorescently labeled proteins exhibit different growth kinetics. In another

Fig. 5.3  Two-color super-resolution microscopy for amyloid fibril growth kinetics: (A) Scheme of two-step nanofibril growth, (B) typical 5 STORM images of two-step grown nanofibrils, (C) fiber length over time, (D) growth speed, and (E) fiber size over concentration. Reproduced with permission from L.H. Beun, L. Albertazzi, D. van der Zwaag, R. de Vries, M.A.C. Stuart, Unidirectional living growth of self assembled protein nanofibrils revealed by super-resolution microscopy, ACS Nano 10 (2016) 4973–4980. Copyright 2016, American Chemical Society.

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study, Pinotsi et al. studied the growth of protein nanofibers of human tau K18, Aβ1–42, and α-synuclein with AFM and CM techniques, which provide a label-free, quantitative assay for understanding the protein aggregation and seeded growth kinetics [67]. It is important that the fluorescent CM technique can be utilized for studying the protein aggregation in vivo, which can promote the in situ study of AD in biomedical fields.

5.3.3 X-ray crystallography analysis To understand the atomic-level structure and possible formation mechanism of protein and peptide nanofibers, X-ray crystallography characterization is also very helpful. Previously, X-ray diffraction (XRD) of microcrystals [68, 69], X-ray fiber diffraction (XRFD) [70, 71], and femtosecond X-ray coherent diffraction (XRCD) [72] have been utilized for studying the atomic structures of a lot of amyloid nanofibers. For example, Eisenberg and coworkers utilized XRD to study the atomic structures of the cross-β spine of more than ten amyloid-like nanofibrils [68, 69]. In their experiments, the amyloid nanofibrils were firstly fabricated to form microcrystalline clusters and the single microcrystals were mounted for XRD tests. For the statistical analysis of the obtained XRD data, several segments identified from the microcrystals were measured. Their finding indicated the common structural features of the tested amyloid nanofibrils at the molecular level and provided an atomic-level hypothesis for the basis of prion strains. Madine et  al. analyzed the structure of amyloid hIAPP20–29 fibrils by using ­solid-state NMR and XRFD techniques [70]. To achieve the fiber diffraction, a droplet of amyloid fibril solution was placed between two wax-filled capillary tubes on a stretch frame, and then the fibrils were dried to form a partially aligned fiber sample for testing. Therefore, it is possible to gain the high-resolution X-ray crystallographic structures of nanofibrils, which in another way are helpful to create the structural models and self-assembly mechanisms of fibrils. To increase the signal-to-noise ratio in XRFD test, Seuring and coworkers, recently, reported an advanced femtosecond XRCD characterization of aligned bombesin amyloid fibrils on low background graphene substrate [72], as shown in Fig. 5.4. They first covered the silicon chips with a graphene monolayer and then a layer of fibrils, and then mounted the sample in vacuum at the Coherent X-ray Imaging (CXI) beamline of the Linac Coherent Light Source (LCLS) beam, and then performed test with the X-ray free-electron laser (XFEL), as indicated in Fig. 5.4A. The use of silicon frames and monolayer graphene reduced greatly the signals of background, as proved by the obtained XRCD patterns in Fig. 5.4B. The tobacco mosaic virus (TMV) fibers were used as reference to compare with amyloid bombesin fibrils. It was found that both TMV fiber and bombesin fibrils exhibited clear diffraction traces compared to the background signals (Fig. 5.4C). This technique provided high resolution of 2.7 and 2.4 Å in single patterns of TMV fibers and amyloid bombesin fibrils, respectively, revealing high potentials for investigating the unsolved structures of nanofibers/fibrils with weak scattering property.

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old

er

CSPAD Detector

(A) 0.2

(B)

Background

0

10

TMV

0

5

Bombesin

0

101

Photons

100 10–1 10–2

Bombesin TMV Background

10–3

(C)

0

0.1

0.2

0.3

0.4

0.5

R (Å–1)

Fig. 5.4  Femtosecond XRCD characterization of amyloid fibrils aligned on graphene: (A) experimental design, (B) diffraction images of different samples (background, TMV fibers, and bombesin amyloid fibrils), and (C) traces from the diagonal lines in the diffraction images of (B) over the reciprocal resolution R (Å− 1). Reproduced with permission from C. Seuring, K. Ayyer, E. Filippaki, M. Barthelmess, J.N. Longchamp, P. Ringler, T. Pardini, D.H. Wojtas, M.A. Coleman, K. Dorner, S. Fuglerud, G. Hammarin, B. Habenstein, A.E. Langkilde, A. Loquet, A. Meents, R. Riek, H. Stahlberg, S. Boutet, M.S. Hunter, J. Koglin, M.N. Liang, H.M. Ginn, R.P. Millane, M. Frank, A. Barty, H.N. Chapman, Femtosecond x-ray coherent diffraction of aligned amyloid fibrils on low background graphene, Nat. Commun. 9 (2018) 1836. Copyright 2018, Nature Publishing Group.

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5.3.4 Other analytical assays Besides the above characterization techniques, a few other analytical assays such as quartz crystal microbalance (QCM) and isothermal titration calorimetry (ITC) have also been utilized for studying the self-assembly and growth kinetics of amyloid nanofibers [73–78]. QCM is a new type of surface-binding-based bioanalytical assay, which offers a sensitive and real-time way to determine the surface mass changes and the relationship between the frequency of oscillation and the material mass by using the piezoelectric property of quartz crystals. Previously, Kotarek et al. investigated the real-time growth kinetics of Aβ1–40 protein by using QCM analysis [73]. To achieve the aim, biotin-­ labeled protein monomers were bound onto the avidin-modified gold-quartz crystal surface (Fig.  5.5A), and then non-labeled Aβ1–40 aggregates were added to interact with labeled monomer. Therefore, the growth kinetics and elongation rate of protein nanofibrils can be measured by QCM by using monomers (Fig. 5.5B) and aggregation Aβ1-40 aggregation intermediate

Au electrode surface

avidin

∆m (pmol cm–2)

10 µM

40

7 µM 5 µM

20

3 µM

0

0 µM

0

(B)

rate (pmol cm–2 min–1)

(A)

biotinylated Aβ1-40 monomer

5 Time (min)

4

2

0

10

0

(C)

5 [M] (µM)

10

Fig. 5.5  QCM studying on the growth kinetics of Aβ1–40 protein: (A) surface modification and binding of monomers and aggregates, (B) growth kinetic over monomer concentration, and (C) growth kinetic over aggregate concentration. Reproduced with permission from J.A. Kotarek, K.C. Johnson, M.A. Moss, Quartz crystal microbalance analysis of growth kinetics for aggregation intermediates of the amyloid-beta protein, Anal. Biochem. 378 (2008) 15–24. Copyright 2008, Elsevier Ltd.

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111

­intermediate (Fig. 5.5C) with various concentrations. It was found that the concentration of monomers, the density of aggregates, ionic strength, and acidic pH could greatly affect the growth kinetics of mature Aβ1–40 fibrils. Buell et al. elucidated and quantified the correlation between the growth rate of nanofibrils and the population of nonnative states by using QCM and other spectroscopic methods [74]. In another work, Walters and coworkers measured the elongation kinetics of polyglutamine peptide nanofibrils by using QCM with dissipation monitoring [75]. In addition, the QCM assay showed qualitative information on the mechanical properties of the elongating amyloid fibrils. ITC has been an effective assay to study the heat effect of ligand binding to proteins or protein aggregation. For instance, Kardos et al. for the first time utilized ITC to directly measure the thermodynamic parameters of β2-microglobulin nanofibrils [76]. They measured the enthalpy and heat capacity changes of the reaction and found that the monomeric, acid-denatured molecules promoted the formation of an ordered, cross-β-sheet structure in the rigid amyloid nanofibrils. Later, Lin and coworkers investigated the effects of temperature, ionic strength, and pH on the aggregation mechanisms of Aβ nanofibrils by combining ITC and spectroscopic methods [77]. In another study, Ikenoue et al. monitored directly the formation of β2-microglobulin nanofibrils by ITC assay [78]. All the above cases indicate that ITC assay could be a promising approach for understanding the protein folding, unfolding, aggregation, and self-assembly kinetics.

5.4 Summary In the above sections, the characterization techniques for studying the self-assembly kinetics of protein and peptide nanofibers are introduced and discussed in detail. To make it more clear, a comparison (Table 5.1) is presented here to show the advantages and disadvantages of every techniques.

5.5 Conclusion and outlooks In summary, the study of the self-assembly kinetics of protein and peptide nanofibers plays important roles in the design and synthesis of novel nanofibrous materials for various applications. To trigger the aggregation and self-assembly of both protein and peptide molecules, a lot of potential strategies, such as adjusting the solution properties, adding organic solvents or surfactants, introducing biopolymers/enzymes/ metal ions/metal nanoparticles, using the solid-liquid or air-water interfaces, and applying external force or pressure, could be utilized. In addition, the characterization techniques including spectroscopy, microscopy, crystallography, and other analytical assays provided enough supports for understanding the aggregation and self-­ assembly mechanism of protein and peptide molecules. This chapter will be helpful for readers to select suitable techniques for characterizing the structure and properties

Table 5.1  Comparison of characterization techniques of the self-assembly kinetics of nanofibers. Techniques Spectroscopy

CD

Fluore-scence FTIR

Raman

NMR

UV-vis

Microscopy

TEM

AFM

STORM CM

Advantages

Disadvantages

Application

Ref.

Real-time, quick measurement, easy operation, small sample amount Real-time, in vitro and in vivo tests, sensitive High resolution, wide wavenumber detection

High requirements for sample, high cost

Measuring secondary structure of biomolecules, protein folding

[45–49]

Fluorescent labeling

Time-dependent growth kinetics of nanofibers Dynamical and structural analysis of nanofibers

[49–52]

Dynamical and structural analysis of nanofibers

[55, 56]

Structure and selfassembly kinetics of nanofibers Quantitative analysis of growth kinetics of nanofibers Length and structural scale for self-assembly kinetics of nanofibers Length scale for selfassembly kinetics of nanofibers Growth kinetics of nanofibers Protein aggregation and growth kinetics

[57–60]

Real-time, easy sample preparation, quick test, high sensitivity Real-time, atomic-level structural information, 3D structure information Low cost, quick test, high sensitivity and selectivity, real-time, quantitative High resolution, surface and structural analysis High resolution, surface analysis, easy sample preparation, air/liquid tests Real-time, high resolution (better than 20 nm) Real-time, in vivo, quantitative

Complicate sample preparation, peak duplicate, non-real-time Affected by some factors like system parameters and fluorescence Low sensitivity compared to FTIR, Raman, UV-vis No structural and conformation information for nanofibers Nanofibers should be stained or frozen Small imaging scale, low speed Needs fluorescent labeling Lower resolution, needs labeling

[53, 54]

[52, 61]

[21, 62]

[24, 63, 64]

[23, 65] [66, 67]

X-ray crystallography

XRD

On atomic level, quantitative and qualitative, easy sample preparation High-resolution structure of nanofibers

Only accurate for large crystalline, relatively low sensitivity Complicate nanofiber sample preparation

XRCD

High resolution, high signalto-noise ratio, structure imaging

Complicate sample preparation

QCM

Real-time, sensitive, quantitative

Surface modification, no conformation information

ITC

Thermodynamic measurement, quantitative

Large sample quantity, low throughput

XRFD

Other assays

Measuring atomic structure of nanofiber microcrystals Measuring molecular structure of long nanofibers Detecting atomic structure of noncrystalline single nanofibers Growth kinetics of protein and peptide nanofibers Binding affinity, protein folding, unfolding, aggregation

[68, 69]

[70, 71]

[72]

[73–75]

[76–78]

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of artificial protein and peptide nanofibers. Meanwhile, it is valuable for investigating the self-assembly kinetics and formation mechanism of nanofibers at the atomic and molecular level.

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[67] D. Pinotsi, A.K. Buell, C.M. Dobson, G.S.K. Schierle, C.F. Kaminski, A label-free, quantitative assay of amyloid fibril growth based on intrinsic fluorescence, ChemBioChem 14 (2013) 846–850. [68] R. Nelson, M.R. Sawaya, M. Balbirnie, A.O. Madsen, C. Riekel, R. Grothe, D. Eisenberg, Structure of the cross-beta spine of amyloid-like fibrils, Nature 435 (2005) 773–778. [69] M.R.  Sawaya, S.  Sambashivan, R.  Nelson, M.I.  Ivanova, S.A.  Sievers, M.I.  Apostol, M.J. Thompson, M. Balbirnie, J.J.W. Wiltzius, H.T. McFarlane, A.O. Madsen, C. Riekel, D. Eisenberg, Atomic structures of amyloid cross-beta spines reveal varied steric zippers, Nature 447 (2007) 453–457. [70] J. Madine, E. Jack, P.G. Stockley, S.E. Radford, L.C. Serpell, D.A. Middleton, Structural insights into the polymorphism of amyloid-like fibrils formed by region 20-29 of amylin revealed by solid-state nmr and x-ray fiber diffraction, J. Am. Chem. Soc. 130 (2008) 14990–15001. [71] A.E. Langkilde, K.L. Morris, L.C. Serpell, D.I. Svergun, B. Vestergaard, The architecture of amyloid-like peptide fibrils revealed by x-ray scattering, diffraction and electron microscopy, Acta Cryst D71 (2015) 882–895. [72] C. Seuring, K. Ayyer, E. Filippaki, M. Barthelmess, J.N. Longchamp, P. Ringler, T. Pardini, D.H.  Wojtas, M.A.  Coleman, K.  Dorner, S.  Fuglerud, G.  Hammarin, B.  Habenstein, A.E. Langkilde, A. Loquet, A. Meents, R. Riek, H. Stahlberg, S. Boutet, M.S. Hunter, J.  Koglin, M.N.  Liang, H.M.  Ginn, R.P.  Millane, M.  Frank, A.  Barty, H.N.  Chapman, Femtosecond x-ray coherent diffraction of aligned amyloid fibrils on low background graphene, Nat. Commun. 9 (2018) 1836. [73] J.A. Kotarek, K.C. Johnson, M.A. Moss, Quartz crystal microbalance analysis of growth kinetics for aggregation intermediates of the amyloid-beta protein, Anal. Biochem. 378 (2008) 15–24. [74] A.K.  Buell, A.  Dhulesia, M.F.  Mossuto, N.  Cremades, J.R.  Kumita, M.  Dumoulin, M.E.  Welland, T.P.J.  Knowles, X.  Salvatella, C.M.  Dobson, Population of nonnative states of lysozyme variants drives amyloid fibril formation, J. Am. Chem. Soc. 133 (2011) 7737–7743. [75] R.H. Walters, K.H. Jacobson, J.A. Pedersen, R.M. Murphy, Elongation kinetics of polyglutamine peptide fibrils: a quartz crystal microbalance with dissipation study, J. Mol. Biol. 421 (2012) 329–347. [76] J. Kardos, K. Yamamoto, K. Hasegawa, H. Naiki, Y. Goto, Direct measurement of the thermodynamic parameters of amyloid formation by isothermal titration calorimetry, J. Biol. Chem. 279 (2004) 55308–55314. [77] M.S.  Lin, L.Y.  Chen, H.T.  Tsai, S.S.S.  Wang, Y.  Chang, A.  Higuchi, W.Y.  Chen, Investigation of the mechanism of beta-amyloid fibril formation by kinetic and thermodynamic analyses, Langmuir 24 (2008) 5802–5808. [78] T.  Ikenoue, Y.H.  Lee, J.  Kardos, H.  Yagi, T.  Ikegami, H.  Naiki, Y.  Goto, Heat of ­supersaturation-limited amyloid burst directly monitored by isothermal titration calorimetry, Proc. Natl. Acad. Sci. U. S. A. 111 (2014) 6654–6659.

Protein synthesis and characterization

6

Shobhit Kadakeria, Michael R. Arulb, Rosalie Bordettb, Nithyadevi Duraisamyb, Hemantkumar Naikc, and Swetha Rudraiahd a Burlington High School, Burlington, MA, United States, bDepartment of Orthopaedic Surgery, University of Connecticut Health Center, Farmington, CT, United States, c PPD, FSP Department, 320 Research Way, Middleton, WI, United States, dDepartment of Pharmaceutical Sciences, School of Pharmacy, University of Saint Joseph, Hartford, CT, United States

6.1 Introduction Proteins are the most versatile macromolecules in all living systems and serve crucial functions in essentially all biological processes. They are one of the four life’s basic building blocks with carbohydrates (sugars), lipids (fats), and nucleic acids (DNA and RNA) comprising the other three. Proteins make up about 42% of the dry weight of human body. They form part of the body’s structure and perform many essential functions, such as allowing us to move, distributing oxygen around the body, fighting infections, transporting substances into and out of cells, controlling chemical reactions, and carrying messages from one part of the body to another, etc. They also perform a vast array of functions within organisms, including catalyzing metabolic reactions, DNA replication, responding to stimuli, providing structure to cells and organisms, transporting molecules from one location to another, etc. Proteins are large biomolecules, or macromolecules, consisting of one or more long chains of amino acid residues. The individual amino acid residues bond together to form a linear chain of amino acids and the bond between adjacent amino acids is known as peptide bond. Hence, the linear amino acid chain is called a peptide. A short chain containing less than 20–30 amino acid residues is commonly called peptides, or oligopeptide. A linear long chain of amino acid residues containing more than 20–30 amino acid residues is called a polypeptide. A protein contains at least one long polypeptide; Fig. 6.1 shows the peptide bonding. The peptide bond has two resonance forms that contribute some double-bond character and inhibit rotation around its axis, so that the alpha carbons are roughly coplanar. The other two dihedral angles in the peptide bond determine the local shape assumed by the protein backbone. The end with a free amino group is known as the N-terminus or amino terminus, whereas the end of the protein with a free carboxyl group is known as the C-terminus or carboxy terminus.

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Peptide bond

Fig. 6.1  Peptide bonding and general primary protein structure.

6.2 Types of proteins There are many different types of proteins in human body. They all serve important roles in our growth, development, and everyday functioning. These proteins have been divided into different types based on their functions. Some examples are: ●

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Structural proteins: Structural proteins provide support in our bodies, for example, the proteins in our connective tissues, such as collagen and elastin. Catalytic proteins: Enzymes are proteins that facilitate biochemical reactions, for example, pepsin is a digestive enzyme in your stomach that helps to break down proteins in food. Defense proteins: Antibodies are proteins produced by the immune system to help remove foreign substances and fight infections. Nucleotide proteins: DNA-associated proteins regulate chromosome structure during cell division and/or play a role in regulating gene expression, for example, histones and cohesin proteins Contractile proteins: Contractile proteins are involved in muscle contraction and movement, for example, actin and myosin Hormonal proteins: Hormone proteins coordinate bodily functions, for example, insulin controls our blood sugar concentration by regulating the uptake of glucose into cells. Transport/carrier proteins: Transport proteins move molecules around our bodies, for example, hemoglobin transports oxygen through the blood.

6.3 Protein structure Proteins serve crucial functions in essentially all biological processes. One of the key properties that enable proteins to participate in such a wide range of functions is their structure. Proteins are linear polymers built of monomer units called amino acids. The chemical bonding between portions of the polypeptide chain helps in holding the protein together, providing its shape. There are two general classes of protein molecules: globular proteins and fibrous proteins. Globular proteins are generally compact, soluble, and spherical in shape. Fibrous proteins are typically elongated and insoluble. Globular and fibrous proteins may exhibit one or more of four types of protein structure.

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There are four levels of protein structure distinguished from one another by the degree of complexity in the polypeptide chain. They are: primary, secondary, tertiary, and quaternary structures. A single protein molecule may contain one or more of the protein structure types.

6.3.1 Primary structure Primary Structure describes the unique order in which amino acids are linked together to form a protein. An example of this structure can be seen in the Fig. 6.2 depiction. Proteins are constructed from a set of 20 or more amino acids. Generally, all amino acids have the alpha carbon bonded to a hydrogen atom, carboxyl group, an amino group, and a variable side chain/group. The side chain/group varies among amino acids and determines the differences between these protein monomers. The amino acid sequence of a protein is determined by the information found in the cellular genetic code. The order of amino acids in a polypeptide chain is unique and specific to a particular protein. Even altering just one amino acid in a protein’s sequence can affect the protein’s overall structure and its function.

6.3.2 Secondary structure Secondary structure refers to local folded structures that form within a polypeptide due to interactions between atoms of the backbone. The most common types of secondary structures are the α helix and the β-pleated sheet. Both structures are held in shape by hydrogen bonds, which form between the carbonyl O of one amino acid and the amino H of another. In an α-helix, the carbonyl (CO) of one amino acid is hydrogen-bonded to the amino H (NH) of an amino acid that is fifth from it in the chain. This pattern of bonding pulls the polypeptide chain into a helical structure that resembles a curled ribbon, with each turn of the helix containing 3.6 amino acids. The side chain/groups of the amino acids stick outward from the α-helix, where they are free to interact. In a β sheet, two or more segments of a polypeptide/protein chain line up next to each other, forming a sheet-like structure held together by hydrogen bonds. The hydrogen bonds form between carbonyl and amino groups of backbone, while the side chain groups extend above and below the plane of the sheet. The strands of a β-pleated sheet may be parallel, pointing in the same direction (meaning that their N- and C-termini match up), or antiparallel, pointing in opposite directions (meaning that the N-terminus of one strand is positioned next to the C-terminus of the other).

Fig. 6.2  Primary structure of protein (R1, R2, and R3 are side chain functional groups).

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Certain amino acids are more or less likely to be found in α-helices or β-pleated sheets. For instance, the amino acid proline is sometimes called a “helix breaker” because its unusual side chain group (which bonds to the amino group to form a ring) creates a bend in the chain and is not compatible with helix formation [1]. Proline is typically found in bends, unstructured regions between secondary structures. Similarly, amino acids such as tryptophan, tyrosine, and phenylalanine, which have large ring structures in their R groups, are often found in β-pleated sheets, perhaps because the β-pleated sheet structure provides plenty of space for the side chains [2].

6.3.3 Tertiary structure The tertiary structure of a polypeptide or protein is the three-dimensional arrangement of the atoms within a single polypeptide chain. The tertiary structure is primarily due to interactions between the side chain/groups of the amino acids that make up the protein. There are several types of bonds and forces that hold a protein in its tertiary structure. ●

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Hydrophobic interactions greatly contribute to the folding and shaping of a protein. The side chain/group of the amino acid is either hydrophobic or hydrophilic. The amino acids with hydrophilic side chains/groups will seek contact with their aqueous environment, while amino acids with hydrophobic side chains/groups will seek to avoid water and position themselves towards the center of the protein. Hydrogen bonding in the polypeptide chain and between amino acid side chain/groups helps to stabilize protein structure by holding the protein in the shape established by the hydrophobic interactions. Due to protein folding, ionic bonding can occur between the positively and negatively charged side chains/groups that come in close contact with one another. Folding can also result in covalent bonding between the sulfur-containing side chains/groups of cysteine amino acids. This type of bonding forms what is called a disulfide bond. Interactions called van der Waals forces also assist in the stabilization of protein structure. These interactions pertain to the attractive and repulsive forces that occur between molecules that become polarized. These forces contribute to the bonding that occurs between molecules.

These interactions/bondings act like molecular “safety pins,” keeping parts of the polypeptide firmly attached to one another leading to their three-dimensional structure.

6.3.4 Quaternary structure Quaternary structure refers to the structure of a protein macromolecule formed by interactions between multiple polypeptide chains. Each polypeptide chain is referred to as a subunit. Proteins with quaternary structure may consist of more than one of the same type of protein subunit. They may also be composed of different subunits. The arrangement of the monomers in the three-dimensional protein is the quaternary structure. Hydrophobic interaction is the main stabilizing force for subunits in quaternary structure. When a single monomer folds into a three-dimensional shape to expose its polar side chains to an aqueous environment and to shield its nonpolar side chains,

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there are still some hydrophobic sections on the exposed surface. Two or more monomers will assemble so that their exposed hydrophobic sections are in contact. Hemoglobin is an example of a protein with quaternary structure. Hemoglobin, found in the blood, is an iron-containing protein that binds oxygen molecules. Hemoglobin’s quaternary structure is the package of its monomeric subunits. Hemoglobin is composed of four monomers. There are two α-chains, each with 141 amino acids, and two β-chains, each with 146 amino acids. Because there are two different subunits, hemoglobin exhibits hetero-quaternary structure. If all of the monomers in a protein are identical, it is called homo-quaternary structure.

6.4 Applications of protein in medicine The origin of diseases often lies in a complex network of biological interactions that need to be understood not only at a clinical level, but also at phenotypic and molecular levels. Modern drug discovery is molecular target driven with the aim of identifying new therapeutic agents that can selectively target disease-specific molecular mechanisms or pathways. The critical cellular functions, including cell growth, DNA replication, transcriptional activation, translation and transmembrane signal transduction, are all regulated by multiprotein complexes. The function, activity, and specificity of these complexes are usually controlled by the Protein-Protein Interactions (PPIs) that occur between the various complexes. The dynamic processes of living organisms depend on the coordinated formation of such PPIs; for example, they occur during DNA replication, gene regulation, transcription and splicing of mRNA, protein synthesis and secretion, formation of intracellular structures and oligomeric enzymes, and in the many pathways associated with cell signaling [3]. PPIs can also be responsible for the development of pathological processes, for example, Alzheimer’s and prion diseases [4, 5], and several types of human cancer. In addition, PPIs between virus-encoded components or between viral proteins and cellular factors occur during the replication and assembly of human viruses in host cells [6]. Thus, inhibition of these interactions is a promising novel approach for rational drug design against a wide number of cellular and microbial targets. The immune system acts as defense against various infectious agents that cause different forms of diseases. Two major components are the antibody-mediated and cell-­mediated immune responses. The antibody-mediated immune system which comprises B-lymphocytes recognizes the type of foreign invading antigens and produces specific antibodies against them. Antibodies are protein components of an adaptive immune system whose main function is to bind antigens, or foreign substances in the body, and target them for destruction. Antibodies stand out for their unique ability to recognize with relatively high specificity and affinity a virtually unlimited number of target biomolecules, known as antigens. This capability makes antibodies truly indispensable tools in biological research today, which can be used for modulation of PPI and silencing.

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Monoclonal antibodies (mAb or moAb) are antibodies that are made by identical immune cells that are all clones of a unique parent cell. Monoclonal antibodies can have monovalent affinity, in that they bind to the same epitope (the part of an antigen that is recognized by the antibody). Monoclonal antibody therapy is a form of immunotherapy that uses monoclonal antibodies (mAb) to bind specifically to certain cells or proteins. The objective is that this treatment will stimulate the patient’s immune system to attack those cells. In the past few years, attempts to inhibit PPIs using antibodies have met with varying degrees of success, In 1975, monoclonal antibody (mAb) (antibodies that are made by identical immune cells that are all clones of a unique parent cell) technique was created by Georges Köhler, César Milstein, and Niels Kaj Jerne by using mouse × mouse hybridoma [7]; they shared the Nobel Prize in Physiology or Medicine in 1984 for the discovery. In 1992, FDA approved first therapeutic mAb Muromonab-CD3 (trade name Orthoclone OKT3) to reduce acute rejection in patients with organ transplants. Since then, FDA has approved many therapeutic mAbs. The humanized antibodies represent the majority of PPI Inhibitors in the clinic. The antibodies are very attractive as therapeutic agents due to their high target specificity. However, they have poor cell and blood-brain barrier permeability properties, are not orally bioavailable, and have high production cost. Their applications remain mainly limited to extracellular targets.

6.5 Bioactive/functional peptides Peptides with structures based on amino acid sequences found at protein-protein interfaces make excellent leads for the development of PPI modulators. Peptides, which are distinguished from proteins based on their smaller size (