Biomaterials from Nature for Advanced Devices and Therapies

By Nuno M. Neves and Rui L. Reis

In-depth information on natural biomaterials and their applications for translational medicine!

• Undiluted expertise: edited by world-leading experts with contributions from top-notch international scientists, collating experience and cutting-edge knowledge on natural biomaterials from all over the world
• A must-have on the shelf in every biomaterials lab: graduate and PhD students beginning their career in biomaterials science and experienced researchers and practitioners alike will turn to this comprehensive reference in their daily work
• Link to clinical practice: chapters on translational research make readers aware of what needs to be considered when a biomaterial leaves the lab to be routinely used

• Language: English
• Publisher: Wiley
• Release date: Oct 13, 2016
• ISBN: 9781119178071

Book preview

Contributors

Mohsen Akbari,

Center for Biomedical Engineering, Department of Medicine, Brigham and Women's Hospital, Harvard Medical School, Boston, MA, 02139, USA

Harvard-MIT Division of Health Sciences and Technology, Massachusetts Institute of Technology, Cambridge, MA, 02139, USA

Laboratory for Innovations in MicroEngineering (LiME), Department of Mechanical Engineering, University of Victoria, Victoria, BC, Canada

Manuel Alatorre-Meda,

3B's Research Group – University of Minho; Headquarters of the European Institute of Excellence on Tissue Engineering and Regenerative Medicine, AvePark, Parque de Ciência e Tecnologia, Zona Industrial da Gandra, 4805-017 Barco GMR – Portugal

ICVS/3B's – PT Government Associate Laboratory, Braga/Guimaraes, Portugal

Matilde Alonso,

G.I.R. BIOFORGE (Group for Advanced Materials and Nanobiotechnology), Universidad de Valladolid – CIBER-BBN, Spain

Luigi Ambrosio,

Institute for Composite and Biomedical Materials IMCB-CNR, Italy

Department of Chemical Science and Materials Technology DCSMT – CNR, Italy

Müge Andaç,

Department of Environmental Engineering, Hacettepe University, Ankara, Turkey

Gerard A. Ateshian,

Department of Biomedical Engineering, Columbia University, New York, NY, USA

Department of Mechanical Engineering, Columbia University, New York, NY, USA

Stephen F. Badylak,

University of Pittsburgh School of Medicine

McGowan Institute for Regenerative Medicine

Elizabeth Rosado Balmayor,

Experimental Trauma Surgery, Klinikum rechts der Isar, Technical University of Munich, Ismaninger Strasse 22, D-81675 Munich, Germany

Hernane S. Barud,

Institute of Chemistry, Sao Paulo State University – UNESP, CP 355 Araraquara-SP, 14801-970 – Brazil

Joseph J. Bellucci

Department of Biomedical Engineering, Duke University, Durham, North Carolina, USA

Jayanta Bhattacharyya,

Center for Biologically Inspired Materials and Material Systems, Duke University, Durham, North Carolina, USA

Aldo R. Boccaccini,

Institute for Biomaterials, University of Erlangen-Nuremberg, 91058 Erlangen, Germany

Assunta Borza Borzacchiello,

Institute for Composite and Biomedical Materials IMCB-CNR, Italy

José Carlos Rodríguez-Cabello,

G.I.R. BIOFORGE (Group for Advanced Materials and Nanobiotechnology), Universidad de Valladolid – CIBER-BBN, Spain

Gabriela D. Carlos,

3B's Research Group – University of Minho, Portugal

ICVS/3B's – PT Government Associate Laboratory, Portugal

Pedro Pires Carvalho,

3B's Research Group – University of Minho; Headquarters of the European Institute of Excellence on Tissue Engineering and Regenerative Medicine, AvePark, Parque de Ciência e Tecnologia, Zona Industrial da Gandra, 4805-017 Barco GMR – Portuga

ICVS/3B's PT Government Associated Lab, Braga/Guimaraes, Portugal

Guoping Chen,

Tissue Regeneration Materials Group, International Center for Materials Nanoarchitectonics, National Institute for Materials Science, 1-1 Namiki, Tsukuba, Ibaraki 305-0044, Japan

Ashutosh Chilkoti,

Center for Biologically Inspired Materials and Material Systems, Duke University, Durham, North Carolina, USA

Department of Biomedical Engineering, Duke University, Durham, North Carolina, USA

Smadar Cohen,

The Avram and Stella Goldstein-Goren Department of Biotechnology Engineering, Ben-Gurion University of the Negev, Beer-Sheva, Israel

The Center for Regenerative Medicine and Stem Cell (RMSC) Research, Ben-Gurion University of the Negev, Beer-Sheva, Israel

The Ilse Katz Institute for Nanoscale Science and Technology, Ben-Gurion University of the Negev, Beer-Sheva, Israel

Vitor M. Correlo,

3B's Research Group – University of Minho; Headquarters of the European Institute of Excellence on Tissue Engineering and Regenerative Medicine, AvePark, Parque de Ciência e Tecnologia, Zona Industrial da Gandra, 4805-017 Barco GMR – Portugal

ICVS/3B's – PT Government Associate Laboratory, Braga/Guimaraes, Portugal

Ana Costa-Pinto,

3B's Research Group – University of Minho; Headquarters of the European Institute of Excellence on Tissue Engineering and Regenerative Medicine, AvePark, Parque de Ciência e Tecnologia, Zona Industrial da Gandra, 4805-017 Barco GMR – Portugal

ICVS/3B's – PT Government Associate Laboratory, Braga/Guimaraes, Portugal

Marta L. Alves da Silva,

3B's Research Group – University of Minho; Headquarters of the European Institute of Excellence on Tissue Engineering and Regenerative Medicine, AvePark, Parque de Ciência e Tecnologia, Zona Industrial da Gandra, 4805-017 Barco GMR – Portugal

ICVS/3B's PT Government Associate Laboratory

Israel González de Torre,

G.I.R. BIOFORGE (Group for Advanced Materials and Nanobiotechnology), Universidad de Valladolid – CIBER-BBN

Adil Denizli,

Department of Chemistry, Biochemistry Division, Hacettepe University, Ankara, Turkey

Pieter J. Dijkstra,

MIRA – Institute for Biomedical Technology and Technical Medicine, Department of Developmental Bioengineering, Faculty of Science and Technology, University of Twente, P.O. Box 217, 7500 AE Enschede, The Netherlands

Alireza Dolatshahi-Pirouz,

Center for Biomedical Engineering, Department of Medicine, Brigham and Women's Hospital, Harvard Medical School, Boston, MA, 02139, USA

Harvard-MIT Division of Health Sciences and Technology, Massachusetts Institute of Technology, Cambridge, MA, 02139, USA

Laboratory for Innovations in MicroEngineering (LiME), Department of Mechanical Engineering, University of Victoria, Victoria, BC, Canada

Ana R. Duarte,

3B's Research Group – University of Minho; Headquarters of the European Institute of Excellence on Tissue Engineering and Regenerative Medicine, AvePark, Parque de Ciência e Tecnologia, Zona Industrial da Gandra, 4805-017 Barco GMR – Portugal

ICVS/3B's – PT Government Associate Laboratory, Braga/Guimaraes, Portugal

James Ferguson,

Ludwig Boltzmann Institute for Experimental and Clinical Traumatology, Donaueschingenstrasse 13, 1200 Vienna, Austria

Helena Ferreira,

3B's Research Group – University of Minho; Headquarters of the European Institute of Excellence on Tissue Engineering and Regenerative Medicine, AvePark, Parque de Ciência e Tecnologia, Zona Industrial da Gandra, 4805-017 Barco GMR – Portugal

ICVS/3B's – PT Government Associate Laboratory, Braga/Guimaraes, Portugal

Michael Floren,

Department of Mechanical Engineering, University of Colorado at Boulder, Boulder, Colorado 80309

Igor Yu Galaev,

DSM Biotechnology Center, Netherlands

Manuela E. Gomes,

3B's Research Group – University of Minho; Headquarters of the European Institute of Excellence on Tissue Engineering and Regenerative Medicine, AvePark, Parque de Ciência e Tecnologia, Zona Industrial da Gandra, 4805-017 Barco GMR – Portugal

ICVS/3B's PT Government Associated Lab, Braga/Guimaraes, Portugal

Junkal Gutierrez,

Depto. Ingenieria Quimica y del Medio Ambiente, Escuela Politécnica Donostia, Pza. Europa 1, 20018, Donostia-San Sebastian, Spain

M. David Harmon,

Institute for Regenerative Engineering, University of Connecticut Health Center, Connecticut 06030, USA

The Raymond and Beverly Sackler Center for Biomedical, Biological, Physical and Engineering Sciences, Connecticut 06030, USA

Department of Orthopaedic Surgery, University of Connecticut Health Center, Connecticut 06030, USA

Departments of Materials Science and Biomedical Engineering, University of Connecticut, Connecticut 06269, USA

Philipp Heher,

Ludwig Boltzmann Institute for Experimental and Clinical Traumatology, Donaueschingenstrasse 13, 1200 Vienna

Trauma Care Consult, Gonzagagasse 11/25, 1010 Vienna

Clark T. Hung,

Department of Biomedical Engineering, Columbia University, New York, NY, USA

Courtney L. Jenkins,

Department of Chemistry, Purdue University, West Lafayette, IN

Marcel Karperien,

MIRA – Institute for Biomedical Technology and Technical Medicine, Department of Developmental Bioengineering, Faculty of Science and Technology, University of Twente, P.O. Box 217, 7500 AE Enschede, The Netherlands

Naoki Kawazoe,

Tissue Regeneration Materials Unit, International Center for Materials Nanoarchitectonics, National Institute for Materials Science, 1-1 Namiki, Tsukuba, Ibaraki 305-0044, Japan

Markus Kerbl,

Ludwig Boltzmann Institute for Experimental and Clinical Traumatology, Donaueschingenstrasse 13, 1200 Vienna

Ali Khademhosseini,

Center for Biomedical Engineering, Department of Medicine, Brigham and Women's Hospital, Harvard Medical School, Boston, MA, 02139, USA

Harvard-MIT Division of Health Sciences and Technology, Massachusetts Institute of Technology, Cambridge, MA, 02139, USA

Laboratory for Innovations in MicroEngineering (LiME), Department of Mechanical Engineering, University of Victoria, Victoria, BC, Canada

Sangamesh G. Kumbar,

Institute for Regenerative Engineering, University of Connecticut Health Center, Connecticut 06030, USA

The Raymond and Beverly Sackler Center for Biomedical, Biological, Physical and Engineering Sciences, Connecticut 06030, USA

Department of Orthopaedic Surgery, University of Connecticut Health Center, Connecticut 06030, USA

Departments of Materials Science and Biomedical Engineering, University of Connecticut, Connecticut 06269, USA

Subhas C. Kundu,

Department of Biotechnology Indian Institute of Technology, Kharagpur-721301, India

Cato T. Laurencin,

Institute for Regenerative Engineering, University of Connecticut Health Center, Connecticut 06030, USA

The Raymond and Beverly Sackler Center for Biomedical, Biological, Physical and Engineering Sciences, Connecticut 06030, USA

Department of Orthopaedic Surgery, University of Connecticut Health Center, Connecticut 06030, USA

Departments of Materials Science and Biomedical Engineering, University of Connecticut, Connecticut 06269, USA

Lorena del Rosario Lizarraga-Valderrama,

Applied Biotechnology Research Group, Faculty of Science and Technology, University of Westminster, London W1W 6UW, UK

Ricardo Londono,

University of Pittsburgh School of Medicine

McGowan Institute for Regenerative Medicine

Wilton R. Lustri,

University Center of Araraquara- UNIARA, Araraquara-SP, Brazil

João F. Mano,

3B's Research Group – University of Minho; Headquarters of the European Institute of Excellence on Tissue Engineering and Regenerative Medicine, AvePark, Parque de Ciência e Tecnologia, Zona Industrial da Gandra, 4805-017 Barco GMR – Portugal

ICVS/3B's – PT Government Associate Laboratory, Braga/Guimaraes, Portugal

Ana L. Marques,

3B's Research Group – University of Minho; Headquarters of the European Institute of Excellence on Tissue Engineering and Regenerative Medicine, AvePark, Parque de Ciência e Tecnologia, Zona Industrial da Gandra, 4805-017 Barco GMR – Portugal

ICVS/3B's – PT Government Associate Laboratory, Portugal

Albino Martins,

3B's Research Group – University of Minho; Headquarters of the European Institute of Excellence on Tissue Engineering and Regenerative Medicine, AvePark, Parque de Ciência e Tecnologia, Zona Industrial da Gandra, 4805-017 Barco GMR – Portugal

ICVS/3B's – PT Government Associate Laboratory, Braga/Guimaraes, Portugal

Adnan Memic,

Center for Biomedical Engineering, Department of Medicine, Brigham and Women's Hospital, Harvard Medical School, Boston, MA, 02139, USA

Harvard-MIT Division of Health Sciences and Technology, Massachusetts Institute of Technology, Cambridge, MA, 02139, USA

Center of Nanotechnology, King Abdulaziz University, Jeddah, 21589, Saudi Arabia

Heather J. Meredith,

School of Materials Engineering, Purdue University, West Lafayette, IN

Claudio Migliaresi,

Department of Industrial Engineering and BIOtech Research Centre, University of Trento, Italy

European Institute of Excellence on Tissue Engineering and Regenerative Medicine, Trento, Italy

Joana Moreira-Silva,

3B's Research Group – University of Minho; Headquarters of the European Institute of Excellence on Tissue Engineering and Regenerative Medicine, AvePark, Parque de Ciência e Tecnologia, Zona Industrial da Gandra, 4805-017 Barco GMR – Portugal

ICVS/3B's – PT Government Associate Laboratory, Portugal

Antonella Motta,

Department of Industrial Engineering and BIOtech Research Centre, University of Trento, Italy

European Institute of Excellence on Tissue Engineering and Regenerative Medicine, Trento, Italy

Sunita Nayak,

Department of Biotechnology Indian Institute of Technology, Kharagpur – 721301, India

Nuno M. Neves,

3B's Research Group – University of Minho; Headquarters of the European Institute of Excellence on Tissue Engineering and Regenerative Medicine, AvePark, Parque de Ciência e Tecnologia, Zona Industrial da Gandra, 4805-017 Barco GMR – Portugal

ICVS/3B's – PT Government Associate Laboratory, Braga/Guimaraes, Portugal

Mehdi Nikkhah,

Center for Biomedical Engineering, Department of Medicine, Brigham and Women's Hospital, Harvard Medical School, Boston, MA, 02139, USA

Harvard-MIT Division of Health Sciences and Technology, Massachusetts Institute of Technology, Cambridge, MA, 02139, USA

School of Biological and Health Systems Engineering (SBHSE), Arizona State University, Tempe, AZ 85287, USA

Adam B. Nover,

Department of Biomedical Engineering, Columbia University, New York, NY, USA

Joaquim Miguel Oliveira,

3B's Research Group – University of Minho; Headquarters of the European Institute of Excellence on Tissue Engineering and Regenerative Medicine, AvePark, Parque de Ciência e Tecnologia, Zona Industrial da Gandra, 4805-017 Barco GMR – Portugal

ICVS/3B's – PT Government Associate Laboratory, Braga/Guimaraes, Portugal

Bijal Panchal,

Applied Biotechnology Research Group, Faculty of Science and Technology, University of Westminster, London W1W 6UW, UK

Arghya Paul,

Center for Biomedical Engineering, Department of Medicine, Brigham and Women's Hospital, Harvard Medical School, Boston, MA, 02139, USA

Harvard-MIT Division of Health Sciences and Technology, Massachusetts Institute of Technology, Cambridge, MA, 02139, USA

Laboratory for Innovations in MicroEngineering (LiME), Department of Mechanical Engineering, University of Victoria, Victoria, BC, Canada

Department of Chemical and Petroleum Engineering, Bioengineering Program, University of Kansas, KS, USA.

Maristela F.S. Peres,

Institute of Chemistry, Sao Paulo State University – UNESP, CP 355 Araraquara-SP, 14801-970, Brazil

A. Hari Reddi,

Ellison Center for Tissue Regeneration, Department of Orthopaedic Surgery, University of California Davis, School of Medicine, Sacramento, California 95817, USA

Heinz Redl,

Ludwig Boltzmann Institute for Experimental and Clinical Traumatology, Donaueschingenstrasse 13, 1200 Vienna, Austria

Trauma Care Consult, Gonzagagasse 11/25, 1010 Vienna, Austria

Rui L. Reis,

3B's Research Group – University of Minho; Headquarters of the European Institute of Excellence on Tissue Engineering and Regenerative Medicine, AvePark, Parque de Ciência e Tecnologia, Zona Industrial da Gandra, 4805-017 Barco GMR – Portugal

ICVS/3B's – PT Government Associate Laboratory, Portugal

Sidney J.L. Ribeiro,

Institute of Chemistry, Sao Paulo State University – UNESP, CP 355 Araraquara-SP, 14801-970, Brazil

Brendan L. Roach,

Department of Biomedical Engineering, Columbia University, New York, NY, USA

Marcia Rodrigues,

3B's Research Group – University of Minho; Headquarters of the European Institute of Excellence on Tissue Engineering and Regenerative Medicine, AvePark, Parque de Ciência e Tecnologia, Zona Industrial da Gandra, 4805-017 Barco GMR – Portugal

ICVS/3B's PT Government Associated Lab, Braga/Guimaraes, Portugal

Ipsita Roy,

Applied Biotechnology Research Group, Faculty of Science and Technology, University of Westminster, London W1W 6UW, UK

Luisa Russo,

Institute for Composite and Biomedical Materials IMCB-CNR, Italy

Emil Ruvinov,

The Avram and Stella Goldstein-Goren Department of Biotechnology Engineering, Ben-Gurion University of the Negev, Beer-Sheva, Israel

Ryosuke Sakata,

Ellison Center for Tissue Regeneration, Department of Orthopaedic Surgery, University of California Davis, School of Medicine, Sacramento, California 95817, USA

Mercedes Santos,

G.I.R. BIOFORGE (Group for Advanced Materials and Nanobiotechnology), Universidad de Valladolid – CIBER-BBN, Spain

Tircia C. Santos,

3B's Research Group – University of Minho; Headquarters of the European Institute of Excellence on Tissue Engineering and Regenerative Medicine, AvePark, Parque de Ciência e Tecnologia, Zona Industrial da Gandra, 4805-017 Barco GMR – Portugal

ICVS/3B's – PT Government Associate Laboratory, Braga/Guimaraes, Portugal

Sybele Saska,

Institute of Chemistry, Sao Paulo State University – UNESP, CP 355 Araraquara-SP, 14801-970, Brazil

Tiago H. Silva

3B's Research Group – University of Minho; Headquarters of the European Institute of Excellence on Tissue Engineering and Regenerative Medicine, AvePark, Parque de Ciência e Tecnologia, Zona Industrial da Gandra, 4805-017 Barco GMR – Portugal

ICVS/3B's – PT Government Associate Laboratory, Portugal

Joana Silva-Correia,

3B's Research Group – University of Minho; Headquarters of the European Institute of Excellence on Tissue Engineering and Regenerative Medicine, AvePark, Parque de Ciência e Tecnologia, Zona Industrial da Gandra, 4805-017 Barco GMR – Portugal

ICVS/3B's – PT Government Associate Laboratory, Braga/Guimaraes, Portugal

Pranav K. Singh,

College of Dairy Science & Technology, GADVASU, Ludhiana, India

Harjinder Singh,

Riddet Institute, Massey University, Palmerston North, New Zealand

Ana Maria Testera,

G.I.R. BIOFORGE (Group for Advanced Materials and Nanobiotechnology), Universidad de Valladolid – CIBER-BBN, Spain

Agniezska Tercjak,

Depto. Ingenieria Quimica y del Medio Ambiente, Escuela Politécnica Donostia, Pza. Europa 1, 20018, Donostia-San Sebastian, Spain

Christy Thomas,

Applied Biotechnology Research Group, Faculty of Science and Technology, University of Westminster, London W1W 6UW, UK

Mark Van Dyke,

Associate Professor, Virginia Tech – Wake Forest School of Biomedical Engineering and Sciences, Virginia Polytechnic Institute and State University, 323 Kelly Hall (0298), Blacksburg, VA 24061, USA

Martijn van Griensven,

Experimental Trauma Surgery, Klinikum rechts der Isar, Technical University of Munich, Ismaninger Strasse 22, D-81675 Munich, Germany

Nicola Volpi,

Department of Life Sciences, University of Modena and Reggio Emilia, Italy

Rong Wang,

MIRA – Institute for Biomedical Technology and Technical Medicine, Department of Developmental Bioengineering, Faculty of Science and Technology, University of Twente, P.O. Box 217, 7500 AE Enschede, The Netherlands

Jonathan J. Wilker,

Department of Chemistry, Purdue University, West Lafayette, IN, USA

School of Materials Engineering, Purdue University, West Lafayette, IN, USA

Preface

The biomaterials field has enabled the development of numerous medical devices and solutions for different clinical conditions, and enormously improved the quality of life of millions of patients. Researchers in this field helped to pioneer landmark developments in devices such as hip and knee implants, catheters, fistulas and hollow fibers for hemodialysis, heart valves or stents. New medical devices always require the development and refinement of biomaterials to obtain enhanced performance. This need provides an ideal motivation to explore natural biomaterials as a source, and inspires progress in the discovery of highly biocompatible and high performance biomaterials for new medical devices. The possibilities offered by nanoscience and nanotechnology open new avenues for the progress in biomaterials science to tackle very demanding health-related problems from a radical new angle. It is expected that the advent of regenerative medicine and tissue engineering, together with the exciting developments in drug delivery systems, will enable development of long-lasting and highly effective therapeutic solutions to previously unmet clinical needs, and also improve the safety and efficacy of currently available medical devices.

The evolution of the biomaterials field is a multidisciplinary endeavor that pursues many different avenues in the quest for high performance materials which are able to positively interact with living organisms. There is passionate discussion as to whether there are particular benefits in following the synthetic biomaterials route instead of using natural and biologically derived biomaterials. It is our perspective that both strategies are worth developing, but we have a particular commitment in exploring the second of those routes. We hope with this book to provide strong evidence that natural and biologically-derived materials provide a wealth of exciting opportunities as sources of biomaterials.

Natural biomaterials such as catgut for sutures and collagen were among the first natural biomaterial systems used in patients. Today, there are a growing number of natural resources that are being explored for biomedical applications such as silk fibroin, DNA-based solutions or protein-based biomaterials. The more we learn about the interaction between cells and biomaterials, the more we realize that we still do not have the knowledge required to rationally design the chemistry, the surfaces and the bulk properties of the biomaterials that would have a perfect integration/interaction with living systems.

We aim to bring to the scientific community a reference book for beginners in the field, and also an in-depth analysis of the achievements and challenges ahead. We hope that this will be useful for experts in many areas of biomedical sciences. Each section of the book is intended to cover not only important developments in the various natural biomaterials properties and medical applications, but also the advances required for the successful development of new and high-end natural-based biomaterials.

Nuno M. Neves and Rui L. Reis

Part I

1

Collagen-Based Porous Scaffolds for Tissue Engineering

Guoping Chen and Naoki Kawazoe

Tissue Regeneration Materials Group, International Center for Materials Nanoarchitectonics, National Institute for Materials Science, 1-1 Namiki, Tsukuba, Ibaraki 305-0044, Japan

1.1 Introduction

Collagen is one of the main components of extracellular matrices that provide mechanical support and biological signals to cells for cellular activities [1]. Collagen has attracted wide attention for biomedical applications because of its versatile property [2–5]. It has been used to construct scaffolds in different forms either with or without hybridization with other biodegradable synthetic or naturally derived materials for tissue engineering. Collagen-based porous scaffolds have been developed through many methods and widely used for tissue engineering of a variety of tissues and organs such as skin [3], bone [5], cartilage [6], ligament [7], blood vessel [8] and nerve [9]. Their pore structures have been well designed and controlled to meet the requirements for cell distribution and cell interaction to promote functional tissue regeneration [10–15]. Hybridization of collagen with mechanically strong synthetic polymers has also developed to improve its mechanical property. Some of the latest developments of collagen-based scaffolds with controlled pore structures and composite structures are summarized and highlighted in this chapter.

1.2 Collagen Sponges

Collagen is a water-soluble polymer and is very easy to prepare its porous sponge by using freeze-drying method [16]. Collagen aqueous solution or collagen gel can be frozen at a low temperature, subsequently freeze-dried at a low pressure and finally cross-linked to prepare collagen porous sponges. During freezing process, freezing temperature may affect the formation and growth of ice crystals in the aqueous solution. Therefore, controlling of freezing temperature has been used to control the porous structure of collagen sponges. Fast freezing at a lower temperature induces cracking, uniform small channels and formation of a fibrous structure. Slow freezing at a higher temperature results in nonuniformity and large pores with more collapsed pores than continuous channels. A unidirectional freezing-drying method has been developed to prepare unidirectionally structured collagen sponge [17]. Collagen sponge resembling the extracellular matrix structure of a particular tissue has been prepared by specific freezing regimes [18]. Although some methods have been deve-loped to prepare collagen sponges with partially controlled pore structures, it has been pursued by many researchers to make the sponge pore open and increase the interconnectivity. Recently a method by using embossing ice particulates as a temperature to precisely control the pore structure of collagen sponges has been developed [10].

The preparation scheme using embossing ice particulates is shown in Figure 1.1. At first, water droplets are prepared by spraying pure water on the surface of a hydrophobic film and water droplets are formed on the surface. The size of the water droplets can be controlled by spraying condition such as spraying speed and spraying time. Or the water droplet can be printed on the hydrophobic surface by a dispenser and the size of the ice droplets can be controlled by the volume of injected volume of water. Subsequently, the water droplets are frozen at a low temperature to form ice particulates embossing the membrane surface. The size and density of embossing ice particulates are controllable. Finally, collagen aqueous solution is eluted onto the embossing ice particulates, frozen, freeze-dried and cross-linked to prepare collagen sponges with a controlled pore structure. Usually the temperature of ice particulates and collagen aqueous solution should be balanced before eluting collagen aqueous solution onto the ice particulates. The prepared collagen porous sponges have large open pores on the surface and interconnected bulk pores underlying the large surface pores. Such structure is very similar to a funnel and therefore the collagen sponges prepared by this method are referred as funnel-like collagen sponges.

Diagram shows preparation of water droplets leads to ice particulate template to collagen solution to keep at designated temperature for 1 hour to funnel-like collagen scaffold.

Figure 1.1 Preparation scheme of the funnel-like collagen sponge using embossing ice particulates. Adapted and reproduced from Ref. 10 (DOI: 10.1177/0883911510370002).

The photo of funnel-like collagen sponge prepared with 398 μm-diameter ice particulates and 1.0% collagen aqueous solution shows clear large pores are evenly distributed on the surface of the collagen sponge (Fig. 1.2a). Scanning electron microscopy images show that large pores are formed on the top surface and interconnected bulk pores are formed beneath the surface pores (Fig. 1.2b and c). The mean diameter of the large surface pores is almost the same as that of the embossing ice particulates which are used as templates because the large surface pores should be the replicas of the embossing ice particulates.

Image described by surrounding text and caption.

Figure 1.2 Photograph (a) and SEM photomicrographs (b, c) of top surface (b) and cross-section (c) of funnel-like collagen sponge prepared with 398 μm-diameter ice particulate template at –3°C. Adapted and reproduced from Ref. 10 (DOI: 10.1177/0883911510370002).

The underlying bulk pores are interconnected with the large surface pores and extend into the bulk body of the sponge from the surface pores. The underlying bulk pores are the replicas of the ice crystals that are formed during freeze-drying. Therefore, the pore structure of the funnel-like collagen sponges is mainly dependent on the size and density of embossing ice particulates and the freezing temperature. The size and density of surface large pores are determined by the size and density of embossing ice particulates. The size and interconnectivity of underlying bulk pores are dependent on the freezing temperature. Funnel-like collagen sponges prepared with the same size of ice particulates (398 μm) but four different freezing temperatures have the same surface large pores (Fig. 1.3a–d). However, the underlying bulk pores have different size. The size of the bulk pores decreases with a decrease of freezing temperature. Figure 1.3e shows the speculated schematic diagram of the formation of the ice crystals during the freezing process and the effect of temperature on the formation of the ice crystals. When the temperature of the aqueous collagen solution is lowered to its freezing point in the absence of embossing ice particulates, random flake-like ice crystals are formed. Some ice crystals may start to form on the surface while some may start from the inner bulk solution. The connectivity of these ice crystals is low. Therefore, the collagen sponges prepared without embossing ice templates have a random porous structure and low pore interconnectivity. In contrast, when the ice particulate templates are used, the ice particulates can serve as nuclei to initiate ice crystallization at the freezing interface of the liquid phase collagen solution. The newly formed ice particulates gradually grow into connected dendritic network. The embossing ice particulates and the newly formed dendritic ice crystal network should result in the formation of the unique funnel-like porous structure of the collagen sponges. Formation of the new ice crystals during freezing process is affected by the temperature. Low temperature results in quick formation of dense, small ice crystals and therefore formation of small bulk pores. On the other hand, high temperature results in slow formation of sparse, big ice crystals and formation of large bulk pores.

Four SEM images on top taken at scale of 100mm. Diagram below show formation of large crystal by slow crystallization to small ice crystals by quick crystallization from -3 to -10°C.

Figure 1.3 SEM photomicrographs of top surfaces of funnel-like collagen sponges prepared by using the embossing ice particulate template with ice particulate diameter of 398 μm at different temperature of –1 (a), –3 (b), –5 (c) and –10°C (d). Schematic diagram of the effect of temperature on the formation of new ice crystals on the ice particulate template and formation of new ice crystals on the surface without ice particulates (e). Adapted and reproduced from Ref. 10 (DOI: 10.1177/0883911510370002).

The funnel-like porous structure facilitates cell seeding and homogeneous cell distribution in the collagen sponges. Compared to control collagen sponge prepared at –3°C without ice template, the funnel-like collagen sponge prepared with 398 μm-diameter embossing ice particulates at –3°C shows good cell penetration and spatially more homogeneous distribution [10]. The funnel-like collagen sponges have been used for three-dimensional culture of fibroblasts and chondrocytes for dermal and cartilage tissue engineering, respectively [19].

The embossing ice particulates method can also be used for preparation of porous scaffolds of other materials such as gelatin, chitosan and hyaluronic acid as long as the freezing temperature of the solution of the other materials is not higher than the melting temperature of the ice particulates. Funnel-like porous scaffolds of chitosan, hyaluronic acid and hyaluronic acid/collagen composite are prepared by the method and have the same effect on cell seeding and distribution as that of funnel-like collagen sponges [20–22].

1.3 Collagen Sponges with Micropatterned Pore Structures

Micropattern structures of porous scaffolds are important to guide the regeneration of tissues and organs with complex structures. The micropatterned structures can be micropatterned pores or micropatterned bioactive molecules. They can arrange cells into a predesigned location and guide the regeneration of complex networks such as capillary and neuronal networks in accordance with the micropatterns [23–24]. Micropatterned structures in three-dimensional (3D) porous scaffolds are desirable because they can mimic the same biological and physiochemical cues as those present in the in vivo microenvironments that surround cells. The embossing ice particulates method can be used to prepare such porous scaffolds with micropatterned structures [25].

Template of micropatterned ice particulates or ice lines is used to prepare the micropattern-structure collagen sponges. The ice particulate and ice line templates are prepared by ejecting water droplets through a dispensing machine on a film at a low temperature. The micropatterns of the ice particulates and ice lines can be designed using a computer program. The other procedures are the same as the embossing ice particulates method as above described. Figure 1.4 shows four types of micropatterned ice templates that are prepared by designing the micropattern and size of the ice particulates (Figure 1.4a, c, e and g). Collagen sponges with different micropatterned pore structures are prepared by using the micropatterned ice particulates as templates (Figure 1.4b, d, f and h). The micropatterned pore structures are the negative replica of the ice templates. The other pores surrounding and underlying the micropatterned pores and lines are negative replica of ice crystals generated during the freezing process.

Image described by surrounding text and caption.

Figure 1.4 Photomicrographs of four types of micropatterned ice particulates templates (a, c, e and g) and SEM Photomicrographs collagen porous scaffolds prepared with the micropatterned templates (b, d, f and h) and collagen sponge with three-dimensionally micropatterned pores that is prepared with micropatterned templates shown in a (i: top surface, j: cross section). Adapted and reproduced with permission from Ref. 25 (DOI: 10.1002/ adma.201200237).

The micropatterned pore layer can be stacked to construct collagen sponges with 3D micropatterned pores (Figure 1.4i and j). In this case, the frozen collagen solution on the first layer of micropatterned ice particulates should be used to prepare the second layer of micropatterned ice particulates instead of the film. By repeating the procedure and later following it with the freeze-drying, cross-linking and washing processes, collagen sponges with micropatterned 3D structures can be prepared. The 3D micropatterned collagen sponge has its top surface similar to that of the collagen sponge with one layer of micropatterned structure as shown in Figure 1.4b. The cross-section has stacked pore structure (Figure 1.4j).

The micropatterning method can also be used to micropattern bioactive molecules in the 3D porous collagen scaffolds [11]. For incorporation of bioactive molecules, a collagen aqueous solution containing bioactive molecules other than pure water should be used. Bioactive molecules are mixed with the collagen aqueous solution. The mixture solution is ejected onto the low-temperature film through a nozzle using a dispensing machine. Different micropatterns composed of the collagen/bioactive molecule solutions can be prepared by designing a program. The ice micropatterns of bioactive molecules are used to prepare collagen porous scaffolds with micropatterned bioactive molecules. The other preparation procedures are the same as above described. The bioactive molecules can also be 3D micropatterned in collagen sponges by repeating the above-mentioned micropatterning procedure. The stacking method is the same as that of collagen sponges with 3D micropatterned pores. Not only single bioactive molecules but also multibioactive molecules can be co-micropatterned in collagen sponges. The multibioactive molecules can be mixed and co-micropatterned together or the multibioactive molecules can be micropatterned separately to construct a co-micropattern structure.

1.4 Collagen Sponges with Controlled Bulk Structures

Although funnel-like collagen sponges prepared with embossing ice particulates have unique pore structures for easy cell seeding and cell penetration to underlying pores, obtaining a homogeneous cell distribution in thick and large scaffolds remain challengeable. To control the bulk pore structures of large and thick scaffolds, some preparation methods have been developed [26–27]. Among these methods, the porogen-leaching method offers many advantages for the easy manipulation and control of pore size and porosity. Although the porogen materials can leave replica pores after leaching, they cannot initiate the formation of surrounding pores. As a result, isolated pores are formed in the scaffold, a situation which is not desirable for tissue engineering scaffolds. To improve pore interconnectivity, the porogen materials are bonded before mixing them with polymer matrix [28–30]. However, the bonded porogen materials require organic solvents for leaching of the porogen materials and the residual solvents are toxic to cells. Penetration of the polymer solution into the bonded porogen material becomes difficult if the polymer solution has a high viscosity.

Embossing ice particulates method shows advantages to control the surface pore structure and to increase the interconnectivity between the surface pores and surrounding pores [10, 22]. By taking this advantage, the ice particulates method should solve the interconnection problem of large scaffolds because the ice particulates can initiate formation of interconnecting ice crystals from their interface in aqueous solution. In this case, free ice particulates should be used as a porogen material. The pre-prepared free ice particulates not only work as porogens to control the pore size and porosity but also work as nuclei to initiate the formation of new ice crystals in the surrounding aqueous solution, therefore increasing the interconnectivity of the collagen sponges [12,31].

Ice particulates are prepared by spraying Milli Q water into liquid nitrogen using a sprayer. The ice particulates can be sieved by sieves with different mesh pores to obtain ice particulates having a specific diameter. The sieving process should be conducted at a low-temperature to avoid melting of ice particulates. The pre-prepared ice particulates are spherical. The free ice particulates are mixed with collagen aqueous solution. The collagen aqueous solution is prepared by dissolving collagen in a solution of ethanol and acetic acid. The mixing process is conducted at a low temperature (e.g. –4°C) at which temperature the ice particulates does not melt and the collagen aqueous solution does not freeze. The two components should be well mixed to obtain an even distribution of ice particulates in the collagen aqueous solution. The mixture of ice particulates and collagen aqueous solution is further frozen at –80°C and freeze-dried and cross-linking to obtain collagen sponges with controlled bulk pore structures.

Collagen sponges prepared with free ice particulates have interconnected large pores and small pores (Fig. 1.5) [12]. The large pores are spherical and are the same size as the free ice particulates. The small pores have a random morphology and different sizes. The small pores surround the large spherical pores. The large pores are negative replicas of the pre-prepared free ice particulates, while the small ice particulates are from the ice crystals formed during freezing. The density of the large spherical pores is dependent on the percentage of free ice particulates.

Image described by surrounding text and caption.

Figure 1.5 SEM photomicrographs of cross sections of collagen sponges prepared with 2% collagen aqueous solution and free ice particulates at a ratio of ice particulates/collagen solution of 50% at low (a) and high (b) magnification. Adapted and reproduced from Ref. 12 (DOI: 10.1177/0883911513494620).

Usually 2% (w/v) collagen aqueous solution is used to prepare collagen sponges when free ice particulates are used as a porogen material. Low collagen concentration (e.g. 1%) may result in collapse of some large pores due to less dense collagen matrix surrounding the large pores. High collagen concentration has very high viscosity and difficulty to be completely mixed with free ice particulates, and therefore resulting in partial collapse of the pore structure.

The concentration of collagen aqueous solution and the ratio of ice particulates can affect the mechanical property of collagen sponges. Young's modulus increases as collagen concentration increases because high collagen concentration can form dense collagen matrix surrounding the large pores. For the influence of ratio of ice particulates, there is an optimal range. When 25, 50 and 75% ice particulates are used, the Young's modulus of the collagen scaffolds increases in an order of 75% <25% <50%. The collagen sponges prepared with 50% ice particulates show the highest Young's modulus. It is much higher than that of the collagen sponge prepared without usage of ice particulates. The difference in the mechanical properties can be explained by the different pore structures. The spherical pores formed by ice particulates are thought to reinforce the collagen scaffolds. The high mechanical strength of the collagen scaffold prepared with 50% ice particulates should be due to the most appropriate packing of the large spherical pores and appropriate filling of the collagen matrix between the large spherical pores.

The collagen sponges prepared with ice particulates show a homogenous cell distribution throughout the sponges when they are used for 3D culturing of bovine articular chondrocytes. In vivo implantation in nude mouse shows a uniform spatial distribution of cells, a uniform ECM distribution and homogeneous tissue formation in the collagen sponges. The collagen sponges prepared with ice particulates are more favorable to the production of cartilaginous ECM and chondrocyte maturation than the collagen sponges prepared without ice particulates [31].

Collagen sponges with a gradient change of pore size from 150 μm to 500 μm are prepared by this method [32]. Ice particulates with diameters of 150–250, 250–355, 355–425 and 425–500 μm are used. Spherical large pores with diameters in a good agreement with the sizes of the ice particulates are formed in the gradient collagen sponges. The effect of pore size on cartilage tissue formation can be directly compared by culturing bovine articular chondrocytes in the gradient collagen scaffolds. The micropores in the scaffolds prepared with ice particulates in the range of 150–250 μm show the most beneficial effect on the gene expression and the production of cartilaginous matrix proteins as well as on cartilage regeneration. The good interconnectivity among the spherical large pores in the gradient collagen sponges facilitates smooth delivery of cells throughout the sponges. Cells can be smoothly delivered and homogenously distributed not only in the larger pores, but also in the small pore region. Easy filling of the seeded cells and proliferated cells in the pores in a diameter of 150–250 μm can increase the cell–cell interactions and provide a real 3D microenvironment to promote chondrogenic differentiation and cartilage tissue formation.

1.5 Hybrid Scaffolds

Although collagen sponges prepared with the free ice particulates have improved mechanical property compared to collagen sponges prepared without ice particulates, their mechanical property is still inferior to porous scaffolds of biodegradable synthetic polymers. Hybrid porous scaffolds of biodegradable synthetic polymers and collagen are prepared by forming collagen microsponges or sponge in the space or opening of porous mechanical skeleton of biodegradable synthetic polymers [13–15, 33–45].

Biodegradable synthetic polymer such as poly(glycolic acid) (PGA), poly(lactic acid) (PLA) and their copolymers of poly(lactic-co-glycolic acid) (PLGA) can be easily formed into porous scaffolds with designated shapes and strong mechanical property, which are the drawbacks of collagen. To combine the advantages avoiding the problems of biodegradable synthetic polymers and collagen, they have been hybridized to construct their hybrid porous scaffolds. A few types of such hybrid porous scaffolds are prepared by introducing collagen microsponges in the pores or interstices of PLGA sponges or meshes (Figure 1.6) [34,40]. The biodegradable synthetic polymer sponge or mesh serving as a mechanical skeleton allows for easy formation into the desired shapes and provides the hybrid scaffolds with appropriate mechanical strength, while the collagen microsponges formed in pores or interstices provide the hybrid scaffolds with a microporous structure, hydrophilicity and good cell interaction.

Image described by surrounding text and caption.

Figure 1.6 SEM photomicrographs of PLGA-collagen hybrid sponge (a) and PLGA-collagen hybrid mesh (b). Adapted and reproduced with permission from Ref. 34 (DOI: 10.1002/(SICI)1521-4095(200003)12:6<455::AID-ADMA455>3.0.CO;2-C) and Ref 40 (DOI: 10.1002/jbm.a.10164).

Collagen sponges can also be formed in the middle space of cup-type porous scaffolds of PLLA and PLGA to prepare hybrid porous scaffolds having high porosity and cell linkage protection capacity. In this case, the mechanical skeleton is not a simple sponge or mesh. A cup-shaped porous skeleton of biodegradable synthetic polymers is used. Collagen sponge is formed in the central space of the cup-shaped porous skeleton and collagen microsponges are formed in the pores or interstices of the porous wall of the cup skeleton [14, 45]. The central collagen sponge and interstitial collagen microsponges are connected. Combination of the highly porous central collagen sponge and the surrounding PLLA-collagen sponge cup results in the high porosity of the hybrid sponge. The cup-shaped porous skeleton serves as a mechanical support and reinforces the hybrid sponge to keep its shape during cell culture and transplantation. The cup-type hybrid porous scaffolds have another effect that the surrounding outside PLLA-collagen sponge cup can protect against cell leakage from the scaffold during cell seeding. The high cell seeding efficiency helps the scaffolds to hold more cells and decreases cell loss. The high porosity provides more space for cell accommodation and tissue formation. The hybrid scaffolds can be used for tissue engineering of skin [38, 44], cartilage [39, 40], bone [41–43] and ligament [37].

1.6 Conclusions

Many methods have been developed to control the pore structures and mechanical properties of polymeric porous scaffolds to meet the requirements for functional tissue engineering. Method using ice particulates as a template or porogen material can prepare collagen sponges with open surface pores, highly interconnected bulk pores and micropatterned structures. Hybridization of collagen sponges with biodegradable synthetic polymers can increase the mechanical property of collagen sponges. All these porous scaffolds have precisely controlled pore and hybrid structures with unique property. They are very useful for tissue engineering of a variety of tissues and organs.

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2

Marine Collagen Isolation and Processing Envisaging Biomedical Applications

Joana Moreira-Silva, Gabriela S. Diogo, Ana L. P. Marques, Tiago H. Silva, and Rui L. Reis

3B's Research Group – University of Minho; Headquarters of the European Institute of Excellence on Tissue Engineering and Regenerative Medicine, AvePark, Parque de Ciência e Tecnologia, Zona Industrial da Gandra, 4805-017 Barco GMR – Portugal ICVS/3B's – PT Government Associate Laboratory, Portugal

2.1 Introduction

Known for thousands of years due to its gluing properties, collagen is the most abundant protein in mammals and a main structural component of extracellular matrix [1, 2]. In mammals, collagen constitutes approximately 25% of the total proteins, being found as structural component of tissues as skin, bone and cartilage [1].

Collagen is structurally formed by a triple helix of three extended protein chains that wrap around one another [3, 4]. Collagen and gelatin are different forms of the same macromolecule, with gelatin being the denatured form of collagen. Collagen represent a unique class of proteins due to a specific amino acid content: glycine in each amino acid triplet, high contents of proline and the only proteins bearing hydroxyproline found in animals [5].

There are, at least, 28 types of collagen known so far, diverse on its composition and tissue distribution on human body [4, 6].

From terrestrial sources collagen is being extracted from different organisms: bovine skin [7], equine tendon [8], porcine skin [9], and other sources. Being the main raw-materials for production of collagen the bovine and porcine skin, but alternative sources are being pursued, namely of marine origin, motivated by the unpopularity of the former due to religious constrains and to active discussion on the associated diseases, such as bovine spongiform encephalopathy (BSE) [8], and the risk they pose to humans. In contrast, marine origin collagen and gelatin have a relatively low risk of possessing unknown pathogens [10]. For this reason, collagen from marine origin is receiving more interest and being considered as an important alternative source [11, 12], and many studies have been conducted to extract collagen from marine sources and have used to screen their potential industrial applications [13–17].

Marine origin collagen has different characteristics from the one obtained from terrestrial sources. It has been found to have different amino acid composition, glycosaminhoglycans content and denaturation temperature [18]. These different characteristics, summed with the low risk of carrying diseases transmissible to humans and not possessing religious constrains, turns the marine collagen superior to its terrestrial equivalents. Collagen from marine organisms has gaining ever growing attention, due to its properties [19–22].

Also different types of collagens can be isolated from different marine organisms. In marine environment, collagen can be found in several organisms, namely in fish (including their skins), marine sponges, jellyfish, among others [23–25]. Type I collagen can be isolated from fish skins, bones and scales [10]. When considering marine sponges, the available methodologies differ and different types of collagens can be isolated [26]. Jellyfish are frequently considered as gelatinous animals, mostly composed of water and a developed collagen-rich mesogloea, from which Type II collagen can be extracted.

Being a ubiquitous structural protein in mammals, collagen is consequently an important ingredient for health-related applications, finding uses in cosmetics, dental composites, skin regeneration templates, biodegradable matrices and shields in ophthalmology, cardiovascular surgery, plastic surgery, orthopedics, urology and neurology. In this perspective, the industrial relevance of collagen is undoubtedly high. Fish by-products, such as skins, bones and scales, are abundant in fish processing plans, are currently directed to low-added value ends (animal feed). Besides the added-value attributed to collagen per se, there is an additional advantage of valorization of a by-product (economic and environmental benefits) [15, 21, 27]. Jellyfish is an abundant organism and have, as described above, a gelatinous nature. The extraction can follow an acetic acid treatment, with the methodology being then tuned according to the species, the amount of biomass and the seasonality [28]. Moreover, marine sponges might possess collagens with unique properties, being clear already the extraction procedure, which needs to be based in alkaline treatments [16].

In the present chapter you will find a comprehensive description of the different methods used for collagen extraction from different by-products, originated by fish processing plans, and from different marine resources. We will give you also insights on the diverse possible applications of marine origin collagen in the biomedical field, particularly for Tissue Engineering and Regenerative Medicine.

2.2 Extraction of Collagen From Marine Sources

The demand for collagen from marine origin has been growing in the last years. The higher number of publications about marine collagen reveals a growing interest from the academic community to come up with new profitable sources.

Marine collagens can be obtained from different sources and the focus is the extraction from the abundant ones with methods that could be easily scale up, so that processes can be profitable for the industry. Therefore, with the appropriate technology, marine collagen can be used as a potential alternative to the use of mammals for this purpose. From the marine environment there are studies to extract collagens from:

Fishes and, more specifically, from:

Skins (ocellate puffer fish [29], seaweed pipefish [30], brownstripe red snapper [31], brownbanded bamboo shark [32], carp [33], largefin longbarbel catfish [23], Japanese sea-bass, chub mackerel, bullhead shark [34], small-spotted catshark [35], skate [36], brown backed toadfish [37], bigeye snapper [38], blacktip shark [39], Atlantic salmon [22], chum salmon [40], Nile perch [41], surf smelt [42], Baltic cod [43]);

Bones (carp [33], skipjack, Japanese sea-bass, ayu, yellow sea bream, horse mackerel [34], Baltic cod [44]);

Cartilages (brownbanded bamboo shark, blacktip shark [24])

Scales (carp [33], spotted golden goatfish [45], tilapia [20], [46], rohu and catla (Catla catla) [47]).

Marine sponges (Chondrosia reniformis [26], Ircinia fusca [48], [49]);

Jellyfishes (Stomolophus meleagris [50], [51], Rhopilema esculentum, Rhopilema asamushi, Catostylus tagi, Rhizostoma pulmo [51], [52], Aurelia aurita, Cotylorhiza tuberculata, Pelagia noctiluca, [52].

Molluscs such as squids (Illex argentinus [53], Dosidicus gigas [54]), cuttlefish (Sepia lycidas [55]) and paper nautilus (Argonauta argo [56]).

Among collagen marine sources, fish are the most widely used, since they are wide available, have no risk of transmitting diseases, have no religious constraints and its extraction processes reveal good yield results. As referred before, considering the total weight of a fish, 75% is discarded, resultant from processing industries, in the form of skins, bones (about 15% of the fish weight), fins, heads, guts and scales (about 2% of the fish weight). More specifically, in cod fillet production, 60% of the fish weight are by-products [57].

According to the occurrence of each type of collagen in nature, it is possible to acquire, for example, type I collagen from fish skins, type II collagen from fish cartilage and jellyfish or other type of collagen from marine sponges [23–25].

Depending on the collagen source, different techniques have been proposed to obtain collagens macromolecules. It involves the preparation of samples, isolation of collagen and its posterior recovery. For fish skins, scales and bones, jellyfish and also molluscs, the extraction process has a common thread and identical steps, with the collagen being isolated with acidic solutions. For marine sponges, however, the same methods cannot be applied, because collagen from this source has different features, and thus, different solvents and reagents will be needed to acquire it.

2.2.1 Extraction of Collagen from Fish, Jellyfish and Molluscs

2.2.1.1 Preparation of samples

2.2.1.1.1 Mechanical preparation

In the case of fish skins, bones and molluscs, before the starting of an extraction process, samples need to be cleaned from any muscle remaining attached to them. Cleaning samples will influence the extraction yield, as this is related to the initial mass of skins (wet or dry) and, along with the cut into small pieces, or even mince them; it will facilitate the action of the chemical solutions used in the extraction process.

2.2.1.1.2 Removal of noncollagenous proteins, pigments and lipids

Marine raw materials have a huge number of unwanted compounds in the final product, such as noncollagenous proteins, pigments or lipids. To remove them, raw materials need to undergo several chemical pre-treatments to obtain a higher yield of collagen in the final stage of the extraction process.

To remove noncollagenous proteins, a sodium hydroxide (NaOH) solution is widely used, with a concentration of 0.1 M NaOH, for a variable period of time. This is a very important step, which will influence the purity of the acquired collagen. This procedure is applicable to squid skin [54], jellyfish [50], cuttlefish [55] and fishes (skin [23], cartilages [24], scales [45] and bones [57]). Another solution that is also used to remove noncollagenous proteins is 1.0 M NaCl (0.05 M Tris-HCl, pH 7.5) for 24 h, but this is not broadly used [58].

Pigments of skins are removed with H2O2 in a NaOH solution, and it is applied mainly when working with squids, due to their rosy appearance [53].

Some fish skins have a higher content in lipids than others (e.g. salmon skins from farmed fish), therefore, it is necessary to defat them. An additional step of cleaning is needed: the use of 10% butyl-alcohol in a ratio of 1:10 (w/v) or 10% ethanol is usually enough to remove the excess of lipids from the samples [37, 42].

*** **Demineralization of fish bones, cartilages and scales

This step is only applicable in mineralized samples. Fish bones are mineralized materials, mainly constituted by calcium phosphate and carbonate. Because some properties of collagens from warm-blooded animals are different (e.g. solubility, denaturation temperature) compared to the cold water species, demineralization methods cannot be the same. To extract collagen from this type of raw materials, it is necessary to first remove its mineral content. For this, two different methods are used: HCl solution with different concentrations being applied (0.1–1.0 M) and ethylenediaminetetraacetic acid (EDTA) also used with different concentrations (0.1–0.5 M) [44]. Its effectiveness is dependent on the concentration and time of reaction. Demineralized bone, that is called ossein, can now proceed to the next extraction phase. The demineralization process with EDTA is also applied to cartilages [24] and scales [45].

These procedure parameters (reaction time, concentration) have been adapted to the different marine sources being used, because they are distinct between each other. After samples pass through the removal of noncollagenous proteins, removal of pigments and/or fats, and/or demineralization, they can now undergo the process of collagen extraction.

2.2.1.2 Collagen extraction

For the extraction of marine collagen with acidic solutions, as it is the case of fish, jellyfish and molluscs, polypeptides in solution have a positive charge and, in acidic conditions, it becomes dominant. Thus, the solubilization increases due to the enhanced repulsion among tropocollagen. The collagen extraction with acidic solutions is suitable for squid [54], jellyfish [50], cuttlefish [55] and fishes (skin [22], cartilages [24], scales [45] and bones [38]).

Several acidic solutions can be used to solubilize collagen: organic acids (acetic, lactic, citric or chloracetic) and inorganic acids (hydrochloric). However, the solvents that presented better yield results are the acetic and lactic acids, being the acetic acid the most used [59]. After solubilized in an acidic solution, collagen is then called Acid Soluble Collagen (ASC).

The yield will be dependent on the extract solution chosen, concentration, time of reaction and also on the source of the raw material (species and age of the animal). Collagen solubility in 0.5 M acetic acid ranges from 2% to 90% and is also dependent on time of exposure. The studies show that it can be 24 h, 48 h or 72 h. Even with more time, no significant improvement is achieved.

If a lower collagen extraction yield is achieved, a re-extraction with acidic solution can be performed or an enzymatic treatment by proteolytic enzymes nonspecific for collagen is a very good aid: trypsin, pancreatin, ficin, bromelain, papain or pepsin. The last one being the most used enzyme for this purpose. These enzymes affect only the telopeptides or the nonhelical ends of the collagen, and thus, they remove intermolecular crosslinks, even the ones that are most stable in acidic solutions. Also after these treatments, the solubility of collagen depends on each fish species: collagen can be totally dissolved or its solubility can only be increased by 5%. This is attributed to different location and type of intermolecular crosslinks, especially the ones that include saccharide [59]. Advantages of using pepsin go beyond the increased extraction efficiency: it has influence on the removal of noncollagenous proteins that still remain in the samples, by hydrolyzing and turning them easily removed, and it can reduce antigenicity caused by telopeptides. After the effect of pepsin, the obtained collagen is then called pepsin soluble collagen (PSC).

2.2.1.3 Collagen recovery

After extraction, collagen will be dissolved in the acidic solution and it is necessary to recover it in a dry final form. At least, two different methods are currently used.

*** **Precipitation with NaCl, dialysis and freeze-drying

This is the most used procedure to precipitate collagen. The supernatant, where collagen is dissolved, is salted out by the addition of NaCl and then it is precipitated with a NaCl solution until reaches a final concentration, that can vary between 2.3 M to 2.6 M NaCl in 0.05 M Tris-HCl (pH 7.5). The resultant precipitate is collected by centrifugation, at 4°C to avoid protein denaturation, dissolved in acetic acid 0.5 M. This collagen in solutions is dialyzed against acetic acid solution or water and finally freeze dried to obtain the dry collagen, which can be stored at room temperature until further usage [24, 33, 37, 39, 45, 55].

*** **Precipitation with K-carrageenan

Collagen fibrils can also be precipitated with K-carrageenan, a polysaccharide isolated from the cell walls of the red seaweed Chondrus crispus. When collagen is solubilized in acidic solutions, it can be precipitated by adding a K-carrageenan solution of 0.5% in 0.5 M acetic acid. The process efficiency depends on the concentration of collagen and K-carrageenan [57].

All these phases, including the samples preparation, extraction and recovery, are performed at low temperature, around 4°C, to reduce the risk of obtaining denatured collagen.

2.2.2 Extraction of Collagen from Other Sources: Marine Sponges

Extraction of collagens from marine sponges do not follow the same steps as for fish, jellyfish and molluscs, since collagen from this source is not soluble in acidic solutions.

As an example, collagen is isolated from the marine sponge Chondrosia reniformis with a basic solution at pH 9.5. In brief, ethanol conserved sponges are washed in tap water, size is reduced and, with a blender, little pieces of sponge are homogenized in a solution of a 100 mM Tris–HCl buffer (pH 9.5; 10 mM EDTA; 8 M urea; 100 mM 2-mercaptoethanol) in a proportion 1:5. A dark colored dispersion is obtained and pH is stabilized with NaOH to 9. After 24 h of extraction, solution is centrifuged and collagen, which is solubilized in the supernatant, is precipitated by addition of acetic acid, lowering pH to 4, and collected by centrifugation. The precipitate is re-suspended in distilled water for washing and removal of 2-mercaptoethanol, centrifuged and later lyophilized [26].

2.3 Collagen Characterization

From mammals to marine resources and by-products, there are diverse sources from where it is possible to extract collagen. Although the same type of collagen can be extracted from those different environments, its features can be different, its composition can vary and it may affect its properties. For instance, mammals' collagen have a higher denaturation temperature than the one observed from marine organisms, due to the lower composition of imino acids, which directly affect stability [35].

Collagens perform the role of structural and mechanical functions in our body, especially in connective tissues, that makes it unique. In regenerative medicine field, the goal is to apply biomaterials with or without cells, in the way that they could best mimic living tissues and enable the recovery of their initial properties and functions. Therefore, it is of uttermost importance that implantable materials, such as collagen, are well known and characterized, regarding both chemical and physical properties.

There are several techniques that are being used to best characterize and know the composition of the extracted collagen. Fourier Transform InfraRed spectroscopy, differential