Blue Biotechnology E-Book

• Herausgeber: Wiley-VCH Verlag GmbH & Co. KGaA
• Kategorie: Fachliteratur
• Sprache: Englisch

Seitenzahl: 1869

Veröffentlichungsjahr: 2018

Beschreibung

With its integral treatment of ecosystem and resource management, this is the only overview of the field to address current thinking and future trends. All contributions have been written with the novice in mind, explaining the basics and highlighting recent developments and achievements.
Unmatched in scope, this two-volume reference covers both traditional and well-established areas of marine biotechnology, such as biomass production, alongside such novel ones as biofuels, biological protection of structures and bioinspired materials. In so doing, it ties together information usually only found in widely dispersed sources to assemble a grand unified view of the current state of and prospects for this multi-faceted discipline.
The combination of the breadth of topics and the focus on modern ideas make this introductory book especially suitable for teaching purposes and for guiding newcomers to the many possibilities offered by this booming field.

Leseprobe

Table of Contents

Cover

Preface

Volume 1

Part I: Bulk Marine Biomass – Industrial Applications and Potential as Primary Sources

Chapter 1: Microalgae: A Renewable Resource for Food and Fuels and More

1.1 Introduction

1.2 Sourcing Microalgae: Algal Culture Collections

1.3 Microalgal Production Systems

1.4 Uses of Microalgal Bioproducts

1.5 Chemotaxonomy: Setting the Stage for Selecting Biofuel Microalgae by Taxonomic Group

1.6 Manipulating Microalgal Lipid Composition with Culture Growth Phase and Conditions

1.7 High‐Value Lipids: Long‐Chain Polyunsaturated Fatty Acids

1.8 High‐Value Lipids: Carotenoid Pigments

1.9 High‐Value Bioproducts: Polysaccharides

1.10 Wastewater Bioremediation and Bioproducts

1.11 Other Bioapplications and the Potential for Bioengineering

1.12 Conclusions

Acknowledgments

References

Chapter 2: Commercial‐Scale Production of Microalgae for Bioproducts

2.1 Introduction

2.2 Commercial‐Scale Production Systems

2.3 Current Commercial Microalgae and Processes

2.4 Potential New Products from Microalgae

2.5 Regulations and Standards standards}"?>

2.6 Conclusion

References

Chapter 3: Ubiquitous Phlorotannins Prospects and Perspectives

3.1 Historical Background

3.2 Biosynthetic Routes and Chemistry

3.3 Subcellular Localization

3.4 Extraction and Purification of Phlorotannins

3.5 Identification Techniques

3.6 Quantification

3.7 Function of Phlorotannins in Brown Algae

3.8 Phlorotannins: Molecules of Interest in Pharmaceutical, Cosmeceutical, Agriculture Biotechnology, and Industrial Polymer Applications

3.9 Pharmacological Applications

3.10 Conclusions and Prospects

References

Chapter 4: The Potential of Microalgae for Biotechnology: A Focus on Carotenoids

4.1 Introduction

4.2 Carotenoid Synthesis

4.3 Functions of Microalgal Carotenoids

4.4 Functional Benefits of Carotenoids as Nutraceuticals

4.5 Conclusion

References

Chapter 5: Applications of Algal Biomass in Global Food and Feed Markets: From Traditional Usage to the Potential for Functional Products

5.1 Introduction

5.2 Algal Products

5.3 Applications

5.4 Conclusions

References

Chapter 6: Phytoplankton Glycerolipids: Challenging but Promising Prospects from Biomedicine to Green Chemistry and Biofuels

6.1 Introduction

6.2 Fatty Acids, Membrane Glycerolipids, and Triacylglycerol in Phytoplankton

6.3 General Principles of Glycerolipid Biosynthesis in Photosynthetic Cells

6.4 Algae‐Based Fatty Acids: Technological Challenges and Promising Applications

6.5 Conclusions

Acknowledgments

References

Chapter 7: The Bioremediation Potential of Seaweeds: Recycling Nitrogen, Phosphorus, and Other Waste Products

7.1 Introduction

7.2 Ulvales in the Bioremediation of Excess Nutrients

7.3 Kelps in the Bioremediation of Excess Nutrients

7.4 Bioremediation of Dissolved Metals with Seaweeds

Acknowledgments

References

Chapter 8: Cultivation and Conversion of Tropical Red Seaweed into Food and Feed Ingredients, Agricultural Biostimulants, Renewable Chemicals, and Biofuel

8.1 Cultivation

8.2 MUZE Processing

8.3 MUZE Products from Red Seaweed

References

Part II: Marine Molecules for Disease Treatment/Prevention and for Biological Research

Chapter 9: Use of Marine Compounds to Treat Ischemic Diseases

9.1 History of Natural Marine Products

9.2 Peripheral Arterial Disease and Cardiovascular Risks:  Treatments and Unmet Needs

9.3 Chemistry

9.4 Biological Properties

9.5 Conclusion

References

Chapter 10: Bioinspiration from Marine Scaffolds

10.1 History of Marine Natural Products

10.2 Chemical Space

10.3 Self‐Organizing Maps: Chemical Diversity of Marine NPs versus Plant NPs

10.4 Conclusion

References

Chapter 11: Guanidinium Toxins: Natural Biogenic Origin, Chemistry, Biosynthesis, and Biotechnological Applications

11.1 General Introduction to Guanidinium Toxins

11.2 Biogenic Source and Vector Organisms

11.3 Chemistry of Guanidinium Toxins

11.4 Synthesis

11.5 Mode of Action and Symptomology

11.6 Existing and Potential Medical and Biotechnological Research Applications

11.7 Conclusions

11.8 Future Perspectives

Acknowledgments

References

Chapter 12: Carrageenans: New Tools for New Applications

12.1 Historical Background

12.2 Chemistry

12.3 Modern Uses of Carrageenans

12.4 Blue Biotechnology for New Products and Applications

12.5 Future Developments

Acknowledgments

References

Chapter 13: Peptide Antibiotics from Marine Microorganisms

13.1 Introduction

13.2 Searching for New Peptide Antibiotics from Marine Microorganisms

13.3 Genomic Approach for New Antibiotics

13.4 Conclusions

Acknowledgments

References

Chapter 14: Recent Developments and Chemical Diversity of Cone Snails with Special Reference to Indian Cone Snails

14.1 Introduction

14.2 Cone Snails' Global Distribution and Ecology

14.3 Research on Indian Cone Snails

14.4 Biology of

Conus

14.5

Conus

Envenomation: Nonfatal and Fatal Reports

14.6 Chemical Diversity of Cone Snails

14.7 Diversity of Conopeptides in Indian Cone Snails

14.8 Therapeutic Application of

Conus

Conopeptides

14.9 Recent Developments and Future Directions

14.10 Concluding Remarks

Acknowledgments

References

Chapter 15: Marine Polysaccharides and Their Importance for Human Health

15.1 General Properties of Polysaccharides

15.2 Marine Polysaccharides from Macroalgae

15.3 Marine Polysaccharides from Marine Animals

15.4 Marine Polysaccharides (EPS) from Microalgae

15.5 Conclusions

References

Chapter 16: Marennine‐Like Pigments: Blue Diatom or Green Oyster Cult?

16.1 Introduction

16.2 Background on the Biodiversity of Blue

Haslea

Species and Marennine‐Like Pigments

16.3 Green Oysters: The Bivalve Point of View

16.4 Can Histology Elucidate the Greening Mechanism?

16.5 Raman Spectroscopy for Sensing

Haslea ostrearia

, Marennine, and Green‐Gill Oysters

16.6 Advances in Elucidating the Structure of Marennine‐Like Pigments

16.7 Colorimetric Analyses

16.8 Can Blue

Haslea

Species Be Considered as Probiotics for Use in Aquaculture?

16.9 Potential Applications for Blue Biotechnologies and Current Issues

16.10 Conclusion

Acknowledgments

References

Chapter 17: Bioprospecting and Insights into the Biosynthesis of Natural Products from Marine Microalgae

17.1 Introduction

17.2 Biosynthesis of Natural Products from Cyanobacteria

17.3 Tools for the Discovery and Characterization of Marine Bioactive Natural Products

17.4 Conclusions

Acknowledgment

References

Chapter 18: Ovothiol: A Potent Natural Antioxidant from Marine Organisms

18.1 Historical Background

18.2 Occurrence of Ovothiols

18.3 Chemistry

18.4 Biosynthesis

18.5 Biological Roles of Ovothiols

18.6 Ovothiol Derivatives

18.7 Biological Activities of Ovothiols

18.8 Conclusions

References

Chapter 19: Bioactive Marine Molecules and Derivatives with Biopharmaceutical Potential

19.1 Introduction

19.2 Challenges Facing the Discovery and Development of Marine Biopharmaceuticals

19.3 Bioactive Metabolites and Molecules

19.4 Methods Used in Biopharmaceutical Research: “From Molecule to Market”

19.5 Conclusions

References

Chapter 20: Marine Pigment Diversity: Applications and Potential

20.1 Introduction

20.2 Pigments in Aquaculture

20.3 Pigments for Cosmetics and Cosmeceutical Applications

20.4 Pigments in Functional Food and Nutraceuticals

20.5 Pigments for Pharmaceuticals and Therapies

20.6 Pigments in Other Applications

20.7 Sourcing and Beyond

20.8 Conclusion

Acknowledgment

Funding

References

Chapter 21: Potential Applications of Natural Bioactive Cyanobacterial UV‐Protective Compounds

21.1 Introduction

21.2 UV Screening Compounds

21.3 Biosynthesis of Cyanobacterial Photoprotective Compounds

21.4 Functions and Applications of UV Protective Compounds

21.5 Conclusion

Acknowledgments

References

Chapter 22: Bio‐Inspired Molecules Extracted from Marine Macroalgae: A New Generation of Active Ingredients for Cosmetics and Human Health

22.1 What are Marine Macroalgae/Seaweeds?

22.2 Life in the Marine Environment and Its Constraints

22.3 Selected Chemical Strategies Developed by Macroalgae

22.4 Extraction of Ingredients (Osmolytes, Polyphenols, and Alginates)

22.5 Cosmetological Applications of Ingredients

22.6 Medical Applications of Ingredients: Wound Dressing and Skin Regeneration

22.7 Conclusion

References

Chapter 23: Emerging Therapeutic Potential of Marine Dinoflagellate Natural Products

23.1 Introduction

23.2 Neosaxitoxin and Gonyautoxin: A New Class of Analgesics

23.3 Brevenal: A Potential New Therapeutic for Cystic Fibrosis

23.4 Cyclic Imine Toxins: Potential Neurodegenerative Disease Drug Leads

23.5 Neuropharmacology and Biotechnology Applications of Cyclic Imine Toxins

23.6 Conclusions

References

Chapter 24: How Fluorescent and Bioluminescent Proteins Have Changed Modern Science

24.1 Introduction

24.2 Bioluminescence

24.3 Organisms that Fluorescence

24.4 Conclusion

References

Volume 2

Part III: Biostructures, Biomaterials, and Biomolecules for other Applications

Chapter 25: Antimicrobial and Antibiofilm Molecules Produced by Marine Bacteria

25.1 Introduction

25.2 Antimicrobial Compounds from Marine Bacteria

25.3 Antibiofilm Molecules

25.4 AlpP and LodA: More Than Just Antimicrobial Proteins

25.5 Conclusion

Acknowledgments

References

Chapter 26: Chitin of Poriferan Origin as a Unique Biological Material

26.1 Historical Background

26.2 Sponges (Porifera) as a Source of Chitin

26.3 Principles of Sponge Chitin Isolation and Identification

26.4 Structural and Physicochemical Properties of Sponge Chitin

26.5 Poriferan Chitin, Tissue Engineering, and Stem Cell Research

26.6 Poriferan Chitin and Extreme Biomimetics

26.7 Conclusions

Acknowledgments

References

Chapter 27: Marine Biominerals with a Biotechnological Future

27.1 Introduction

27.2 Calcium Carbonate‐Based Biominerals

27.3 Silica‐Based Marine Biominerals

27.4 Heavy‐Metal Bioaccumulations

27.5 Marine Biominerals and Composites in Novel Technologies

27.6 Biointegrative Solutions from Nano to Macro to Giga

Acknowledgments

References

Postface

Index

End User License Agreement

List of Tables

Chapter 1

Table 1.1 A selection major algal culture collections.

Table 1.2 Summary of features based on fatty acid composition of the chemotaxonomic microalgal groups, using screening data compiled from >40 species held in the Australian National Algae Culture Collection.

Table 1.3 Pigment composition of vegetative cells and resting‐stage aplanospores of

Haematococcus

.

Chapter 2

Table 2.1 Commercial microalgae culture systems currently used, the algal species cultured, and approximate maximum culture volume.

Table 2.2 Summary of existing and potential high‐value products from microalgae, alternative sources, and application of these products.

Chapter 3

Table 3.1 Classification of the elucidated chemical structures of phlorotannins.

Table 3.2 Overview of the different staining of phlorotannin compounds.

Table 3.3 Summary of phlorotannin structures and their possible pharmacological applications.

Chapter 4

Table 4.1 Stressor‐induced carotenoid production leading to increased pigment contents in microalgae.

Table 4.2 Saturating light intensities for photosynthesis (

P

max

) and maximum cultivation light intensities for several microalgal species.

Table 4.3 Metal‐induced reactive oxygen species (ROS) formation and resulting enhancement of carotenoid content in microalgae.

Table 4.4 Molybdenum uptake potential by various green microalgae.

Table 4.5 Carotenoid pigments and macroalgal species with commercial potential for their production.

Table 4.6 Pigment contents in a number of microalgal species.

Chapter 5

Table 5.1 Average composition of some microalgae (% dry weight), after [5].

Table 5.2 Average composition of some macroalgae (% dry weight).

Table 5.3 Average composition of human food sources (% dry weight).

Table 5.4 Examples of macroalgal species with potential human health benefits.

Table 5.5 List of applications (not related to bioenergy) for selected microalgae.

Table 5.6 Macroeconomic data linked to animal feeds worldwide.

Table 5.7 Main raw material production for human food in the world, after [21].

Table 5.8 Worldwide production of microalgae and seaweed in 2012, extracted from the Fisheries and Aquaculture database of the FAO (http://www.fao.org/fishery/statistics/en).

Table 5.9 List of Food forSpecified Health Uses (FOSHU) ingredients in Japan in 2005, after [150].

Table 5.10 Functionality claims approved for functional food in China in 2005, after [152].

Chapter 6

Table 6.1 Linear, even‐numbered, and odd‐numbered fatty acids.

Table 6.2 The three different diacyl precursors of glycerolipids in microalgae.

Table 6.3 Molecular diversity of membrane lipids' polar heads.

Chapter 8

Table 8.1 Some known producers of seaweed concentrates (SWC) and their base countries as of 2016, with larger producers designated by an asterisk (*).

Table 8.2 Postulated efficacies of seaweed concentrates (SWC) components applied to plants and animals in agriculture showing indications of the state of knowledge at the time of writing.

Table 8.3 A matrix of impacts that application of seaweed concentrates (SWC) can have in agricultural plant and animal applications.

Chapter 9

Table 9.1 Examples of marine‐derived carbohydrate‐based therapeutic molecules (GlycoMar www.glycomar.com).

Table 9.2 Examples of carbohydrate‐based drugs in use or in development.

Chapter 12

Table 12.1 Early events in the use of carrageen (from

Chondrus crispus

) and its mucilages, the carrageenans.

Table 12.2 Carrageenophyte global production data 1994–1995.

Table 12.3 Summary of global commercial carrageenophyte production and values in 2012.

Table 12.4 Comparison of the characteristics of

1

H and

13

C NMR in the study of carrageenans.

Table 12.5

1

H NMR chemical shifts of the three main commercial carrageenans in D

2

O.

Table 12.6

13

C NMR chemical shifts of the three main commercial carrageenans in D

2

O.

Table 12.7

1

H and

13

C NMR chemical shifts of methyl and pyruvate substituents of carrageenans in D

2

O at 65 °C in reference to DSS, after [78, 138].

Table 12.8 New products and potential opportunities for carrageenan uses.

Table 12.9 Carrageenan sulfurylases described in the literature.

Table 12.10 Characterized carrageenan hydrolases described in the literature.

Chapter 14

Table 14.1 Geographical distribution of cone snails in Indian waters.

Table 14.2 Cone shells' envenomation nonfatal and fatal case report 1850–2009.

Table 14.3 Classification of Conotoxin superfamilies.

Table 14.4 Novel conotoxins isolated from Indian

Conus

species.

Chapter 15

Table 15.1 Summary of classified polysaccharides, with examples for each criterion.

Table 15.2 Main marine algae polysaccharides and their biological properties.

Table 15.3 Some registered trademarks based on marine algae polysaccharides.

Table 15.4 General characteristics of hydrogel‐based wound dressings, with examples of commercial products.

Table 15.5 General characteristics of commercial alginate hydrogel dressings for wound dressing.

Table 15.6 Chitosan biological properties and some related applications.

Table 15.7 Chitosan derivatives.

Chapter 17

Table 17.1 Examples of natural products from marine microalgae.

Chapter 18

Table 18.1 Occurrence of thiohistidine‐related metabolites.

Table 18.2 Spectral and optical properties of ovothiol A disulfide.

Chapter 19

Table 19.1 Marine‐derived compounds approved by the US

Food and Drug Administration

(

FDA

), which are in phases I–III of drug development as of April 2016 [6, 61].

Table 19.2 Marine peptides that have antimicrobial biopharmaceutical potential, their source, and target of inhibition.

Chapter 20

Table 20.1 Categories of organic pigments and examples of chemical structures.

Table 20.2 Ro5 parameters for a subset of 56 pigments.

Chapter 21

Table 21.1 Molecular structure and absorption maxima of MAAs commonly present in cyanobacteria.

Chapter 22

Table 22.1 Examples of marine active ingredients derived from macroalgae and present in commercial brands on the European and world cosmetic markets.

Table 22.2 Examples of commercial alginate‐based dressings for wounds.

Chapter 23

Table 23.1 Clinical trials with GTX 2/3 or NeoSTX.

Chapter 24

Table 24.1 Commonly used marine luciferases.

Chapter 25

Table 25.1 Antibiofilm marine compounds.

Chapter 26

Table 26.1 Overview of sponge chitin‐based composite materials obtained via extreme biomimetics route.

Chapter 27

Table 27.1 Examples of current or potential uses for diatom frustules in industrial applications.

List of Illustrations

Chapter 1

Figure 1.1 (a) Microalgal cultures maintained in constant environment room. (Australian National Algae Culture Collection (ANACC).). (b) Raceway pond for

Dunaliella bardawil

production. Nature Beta Technologies Ltd., Eilat, Israel. (Courtesy of Dr Ami Ben‐Amotz.). (c) Microalgal cultures grown in disposable plastic bags supported vertically in metal frames. Shellfish Culture Ltd., Tasmania. (Courtesy of Shellfish Culture Ltd.). (d) 60 L vertical annular column photobioreactors. Algal Culture Laboratory, CSIRO, Hobart, Australia. (Courtesy of Dion Frampton.). (e) 100 L stainless steel bioreactor.

Figure 1.2 Average percentage compositions of the long‐chain polyunsaturated fatty acids (PUFAs) docosahexaenoic acid (DHA; 22:6n‐3), eicosapentaenoic acid (EPA; 20:5n‐3), and arachidonic acid (AA; 20:4n‐6) of different microalgal classes.

Figure 1.3 Pigment profiles of different microalgal classes from strain representatives held in the Australian National Algae Culture Collection. Within one class, there can be variation between genera, species, and even strains. Pigment key: But‐fuco, 19′‐butanoyloxyfucoxanthin; Fuco, fucoxanthin; Neo, 9′‐

cis

‐neoxanthin; 4‐keto‐hex‐fuco, 4‐keto‐19′‐hexanoyloxyfucoxanthin; Pras, prasinoxanthin; Viola, violaxanthin; Hex‐fuco,19′‐hexanoyloxyfucoxanthin; Astax, astaxanthin; Diadino, diadinoxanthin; Antherax, antheraxanthin; Allo, alloxanthin; Myxo, myxoxanthophyll; Diato, diatoxanthin; Zea, zeaxanthin; Lut, lutein; Canthan, canthaxanthin; DV chl

b

, divinyl chlorophyll

b

; Chl

b

, monovinyl chlorophyll

b

; DV chl

a

, divinyl chlorophyll

a

; β,ɛ‐car, β,ɛ‐carotene; β,β‐car, β,β‐carotene.

Chapter 2

Figure 2.1 Center pivot pond used for

Chlorella

production in Taiwan.

Figure 2.2 Raceway ponds at the

Arthrospira

(

Spirulina

) facility operated by Earthrise Nutritionals LLC in Calipatria, California, USA.

Figure 2.3 Schematic diagram of the stages of production of

Arthrospira

.

Figure 2.4

Dunaliella salina

β‐carotene production plant operated by BASF at Hutt Lagoon, Western Australia.

Figure 2.5 Schematic diagram of

Dunaliella salina

β‐carotene production by BASF in Australia.

Figure 2.6 Schematic diagram of the process of

Haematococcus pluvialis

astaxanthin production. Note that the culture of the green and red cells may be either in open ponds, closed photobioreactors, or internally lit reactors.

Figure 2.7 Outdoor tubular photobioreactors at the Algatechnologies

Haematococcus pluvialis

plant in Kibbutz Ketura, Israel.

Chapter 3

Figure 3.1 Biosynthesis of a phloroglucinol unit by a type III polyketide synthase from malonyl‐CoA as the sole substrate.

Figure 3.2 Theoretical extraction and purification chart for phlorotannins.

Figure 3.3 Proton NMR spectra and 2D HMBC NMR spectra for two polymers isolated in semi‐purified extracts of

F. spiralis

: (a) fucol type and (b) fucophlorethol type.

Figure 3.4 ESI mass spectrum of a defatted ethyl acetate fraction of

Ascophyllum nodosum

after dialysis on a 2 kDa cutting size membrane showing various oligomeric forms of phlorotannins between DP9 and DP15.

Figure 3.5 Roles of phlorotannins at different stages of the life of algae. (1) Inhibition of spermatozoid motility and prevention of polyspermy; (2) construction of the cell wall by cross‐linking of phlorotannins and carbohydrates; (3) cell division and polarization; (4) adhesion to the substratum synthesis of bioadhesive material; (5) antibacterial and antifouling activities; (6) chelation of heavy metals; (7) protection against UV radiations; (8) chemical signaling by waterborne cues, prevention of attacks; and (9) chemical defenses against grazers and wound‐healing processes.

Chapter 4

Figure 4.1 Schematic microalgal carotenoid synthesis pathway based on Refs [8–14]. BCH, β‐carotene hydroxylase; BCK, β‐carotene ketolase;

CTI

,

carotenoid isomerase

;

GGR

,

geranylgeranyl reductase

;

GGPPS

,

geranylgeranyl pyrophosphate synthase

;

IDI

,

isopentenyl pyrophosphate isomerase

;

LCY

,

lycopene cyclase

; PDS, phytoene desaturase; PSY, phytoene synthase; ZDS, zeta carotene desaturase.

Figure 4.2 Schematic of pigment function. Anth, antheraxanthin; Astx, astaxanthin; β‐car, β‐carotene; Chl, chlorophyll; Lut, lutein; Neo, neoxanthin; ROS, reactive oxygen species; Viola, violaxanthin; Zea, zeaxanthin.

Figure 4.3 Schematic overview of reactive oxygen species (ROS) formation and microalgal ROS detoxification mechanisms.

Figure 4.4 Irradiance regulation of epoxidation and de‐epoxidation of xanthophyll cycle pigments.

VDE

,

violaxanthin de‐epoxidase

;

ZE

,

zeaxanthin epoxidase

.

Figure 4.5 Metal stress‐induced cellular generation of reactive oxygen species (ROS) and hypothesized sites of carotenoid action. APX, ascorbate peroxidase; CAT, catalase; GPX, glutathione peroxidase;

GSSG

,

two molecules of glutathione linked by a disulfide bond

;

MDAsc

,

monodehydroascorbate

; SOD, superoxide dismutase. (

Figure 4.6 Chemical structures of astaxanthin isomers, after [63].

Chapter 6

Figure 6.1 Building blocks and general structure of glycerolipids found in microalgae. (a) Glycerol. This tri‐alcohol is the universal backbone of glycerolipids. The stereospecific numbering nomenclature (

sn‐x

) is shown. (b) Glycerol 3‐phosphate. This molecule harbors a phosphate group at the

sn

‐3 position. (c) Acyl‐CoA and acyl‐ACP thioesters. The thioester bond allows the activation of fatty acids, such that the hydrolysis of this bond releases the chemical energy required for an acyl transfer. (d) Lyso‐phosphatidic acid (Lyso‐PA). (e) Phosphatidic acid (PA). (f) Diacylglycerol (DAG). (g) Lipids with polar heads. Phosphoglycerolipids (PC, PE, PS, PI, PG, and DPG) and galactoglycerolipids (MGDG, DGDG, and SQDG) constitute the membrane glycerolipids (Table 6.3). (h) Triacylglycerol (TAG). By contrast with polar lipids, TAG accumulates in the form of lipid droplets (LDs) or

oil bodies

(

OB

s) in microalgae.

Figure 6.2 Challenges faced by the development of an algae‐based industry. Microalgae need to be screened and selected from the phytoplankton biodiversity. Their growth needs to be optimized and controlled in various systems such as open ponds, photobioreactors, and so on. Microalgae can be improved by genetic engineering or directed evolution. A stock of variants, preserving a sufficient genetic diversity, has to be maintained, just like cultivar collections in agriculture. The design of large‐scale cultivation systems has to implement supplies of water (either from natural source or ideally waste water from urban or industrialized area), CO

2

(including flue gas from industrial plants), other carbon sources, nutrients such as phosphorus and nitrogen, light, energy, and so on. The process to extract biomolecules is one of the most important challenges for the future in order to exploit all possible by‐products, become economically viable in the long term, and respect sustainability criteria.

Chapter 7

Figure 7.1 Simplified schematic of the role of seaweed bioremediation in the recycling of water (blue line) and solid resources (red line) through diverse industries and bio‐products.

Chapter 8

Figure 8.1 Schematic depiction of interrelationships among crop and environmental factors that impact farm productivity. The “Goldilocks zones” are regions where crop factors and environmental factors are “just right” for profitable farm development and operation.

Figure 8.2 Comparison of (A) single‐stream processing as is prevalent for tropical red seaweed processing as of 2016 and (B) multi‐stream, zero‐effluent processing as it is being applied at present and developed for the future.

Chapter 9

Figure 9.1 Colorful seaweed in rock pool on the coast of Brittany in France. From right bottom to the upper left: brown algae

Laminaria digitata

, green algae

Ulva lactuca

, red algae

Palmaria palmata

, green algae

Enteromorpha intestinalis

, brown algae

L. digitata

, and

Fucus vesiculosus

. (Courtesy of S. La Barre.)

Figure 9.2 Peripheral arterial disease in the lower limb is characterized by chronic obstruction of the arteries supplying the leg, gradually leading to CLI. High blood pressure, smoking, and high cholesterol can damage the endothelium and lead to the development of a fatty streak under the endothelium. An unstable plaque develops with a fatty core that can sometimes rupture into the bloodstream. Clotting of the blood begins at the site of the plaque rupture. The artery becomes partially blocked and blood flow is diminished. Accordingly, there is insufficient blood flow to redirect toward the collaterals. Furthermore, acute arterial occlusion leads to distal tissue hypoxia and causes the activation of an inflammatory response against ischemic tissue injury owing to endothelium dysfunction and several factors. The insufficient collaterals and angiogenic signaling limit tissue regeneration and can lead to necrosis and loss of tissue function if revascularization is not reestablished.

Figure 9.3 The angiogenesis and vasculogenesis processes. Recruitment of the endothelial cells from preexisting vessels plays a critical role in the regulation of angiogenesis. (1) Mobilized bone marrow endothelial progenitor cells (EPCs) with high proliferative capacity migrate and (2) home to the site of angiogenesis where (3) they deliver signals that activate pericytes, which then in turn play an active role by secreting mediators that stimulate migration and replication of local endothelial cells [46, 47]. A small amount differentiates and incorporates in the new vessel wall. EPC, endothelial progenitor cells; VEGF, vascular endothelial growth factor;

erythropoietin

(

EPO

), MMPs,

nitric oxide

(

NO

).

Figure 9.4 Activation of local angiogenesis is a promising approach to patients with critical limb ischemia with a high risk of amputation. Stem/progenitors cells isolated from bone marrow or peripheral blood can be pre‐activated with SDF‐1, VEGF, fucoidan, hypoxia, and so on or modified to improve survival, homing, functional engraftment, and functional activity (e.g., differentiation) before being infused or injected in patients. Infusion of these

in vitro

expanded bone marrow cells enhances neovascularization in animal models of hindlimb ischemia.

Figure 9.5 Fucoidan (Fuc) can prevent thrombosis by catalyzing thrombin inhibition and promoting fibrinolysis. It reduces circulating level of thrombin by catalyzing its inhibition:

tissue factor

(

TF

) bound to the endothelium initiates the coagulation cascade. It binds to activate circulating

factor VII

(

FVIIa

) and catalyzes the activation of factor X. Then factor Xa (FXa) binds to

factor V

(

FV

) present on the phospholipid surface of platelets or endothelial cells to catalyze the conversion of prothrombin to activate

thrombin

(

IIa

). Thrombin cleaves fibrinogen to form fibrin that leads to the formation of fibrin clot and then a thrombus. The propagation of the cross‐linked fibrin clot is limited by plasmin, which cleaves fibrin. Endothelial cells secrete

tissue plasminogen activator

(

tPA

) to activate plasmin and control the expansion of the clot on the injured endothelial cell surface. Fuc is a direct inhibitor of thrombin and catalyzes the inhibition of serine proteases Xa and thrombin by antithrombin (AT) and the inhibition of thrombin by heparin cofactor II (HCII). It promotes fibrinolysis by potentiating plasminogen activators such as tPA and reducing plasminogen activator inhibitor‐1 (PAI‐1) release by endothelial cells.

Figure 9.6 Fucoidan induces a proangiogenic phenotype in human EPC. (1)  Fucoidan enhances EPC proliferation in a concentration‐dependent manner starting at 1 µg mL

−1

. (2) Fucoidan pretreatment promotes EPC motility (no chemoattractant) and enhances EPC chemotaxis toward VEGF. 3. Fucoidan pretreatment enhances basic fibroblast growth factor 2 (FGF2)‐induced vascular tube formation by EPC: (3A) EPCs do not form tubular structures in control medium, (3B) 18 h after seeding, FGF2‐pretreated cells are elongated and interconnected, and (3C) the tubular network is significantly more extensive in the presence of FGF‐2 and fucoidan. (4) Fucoidan pretreatment enhances EPC adhesion on activated endothelium under static and dynamic conditions and their extravasation toward VEGFs (40 ng mL

−1

). (5) Fucoidan induces mobilization of immature CD34

+

CD31

+

CD45

−

murine progenitors. Wild mice were intraperitoneally injected with PBS (negative control, CTRL), 5 mg kg

−1

of low molecular weight fucoidan (LMWF), or 0.5 µg kg

−1

VEGF (positive control, VEGF) and bled 30 min after the injection. Blood was assayed for CD34

+

CD31

+

CD45

−

cells, and plasma samples were analyzed for SDF‐1 concentration by enzyme‐linked immunosorbent assay test. (5A) Progenitor mobilization after intraperitoneal injection of PBS (negative CTRL), LMWF, or VEGF. (5B) SDF‐1 level in mice plasma after intraperitoneal injection of fucoidan compared with controls. Results are expressed as the means ±SEM *

p

 < 0.05; **

p

 < 0.01; ***

p

 < 0.001.

Figure 9.7 Fucoidan protects ischemic tissue against necrosis. Fucoidan pretreatment of cultured EPC (

endothelial colony‐forming cells

,

ECFC

) increases neovascularization in hindlimb ischemia of mice with PAD 14 days after artery ligation and intravenous injection of pretreated ECFC. Macroscopic aspects of ECFC‐injected mice (a) and fucoidan‐ECFC‐injected mice (b). Cumulative incidence of clinical necrosis (c), foot perfusion (d), and quantitative analysis of angiographic score (e) of normal saline (CTRL, ▪), untreated ECFC (ECFC, □), and fucoidan‐stimulated ECFC (ECFC, green □) transplanted mice. Hematoxylin and eosin staining of the same distal gastrocnemius muscle sections on day 14 and quantification of histologically preserved area, ischemic infiltrated area, and necrotic area type surface (f). The surface of each area type is reported as a percentage of the entire histological section surface. (g) To evaluate the effect of fucoidan on critical ischemia, two groups of animals underwent the surgical procedure and received two intramuscular injections 1 and 2 days after surgery of normal saline and fucoidan solution (15 mg kg

−1

Fuc). Quantification of histologically preserved area, ischemic infiltrated area, and necrotic area type surface 14 days after surgery and intramuscular bolus administration of fucoidan. Values are expressed as means ±SEM (

n

 = 15) *

p

 < 0.05, **

p

 < 0.01, and ***

p

 < 0.001 versus normal saline‐injected mice (CTRL). NI, non‐ischemic hindlimb; isch, ischemic hindlimb.

Figure 9.8 Proposed mechanisms involved in beneficial effects of fucoidan in peripheral ischemia. (a) Fucoidan can bind to, potentiate, and inactivate cationic proteins such as adherent proteins expressed in extracellular matrix (ECM) and bone marrow, enzymes, growth factors, cytokines, and so on. (b) Fucoidan can prevent thrombosis by catalyzing thrombin inhibition, promoting fibrinolysis, and stimulating the release of

tissue factor pathway inhibitor

(

TFPI

). (c) Fucoidan can promote EPC recruitment by displacing sequestered SDF‐1 from HSPG. It can also compete with EPC for binding

heparan sulfate proteoglycan

s (

HSPG

s) or ECM proteins. (d) Fucoidan can bind to EPC and induce a proangiogenic phenotype to EPC by enhancing their mobility. (e) Fucoidan enhances EPC chemotaxis toward VEGF, EPC attachment to laminin, and growth factor‐induced vascular tube formation, improving revascularization of ischemic tissue.

Chapter 10

Figure 10.1 Chemical structures of morphine, quinine, and salicylic acid.

Figure 10.2 Chemical structures of spongothymidine and spongouridine, vidarabine (ara‐A), and cytarabine (ara‐C).

Figure 10.3 Key events in natural product isolation and discovery. The exploration of natural products started in the early eighteenth century with analyzing medicinal plants to identify active ingredients. In 1929, the discovery of penicillin changed the course of medicinal chemistry and started the era of antibiotics, while marine life exploration started in the 1960s, and attention increased slowly. However, in the last decade marine compounds played a pivotal role in natural product drug discovery.

Figure 10.4 Amino acid sequence of ziconotide.

Figure 10.5 Chemical structure of ecteinascidin‐743 and mechanism of action.

Figure 10.6 Chemical structure of halichondrin B and eribulin.

Figure 10.7 Chemical structure of dolastatin 10, MMAE, and brentuximab vedotin.

Figure 10.8 Stepwise scaffold tree generation shown for the example of varioxepine A. Starting from the quite complex molecule varioxepine A, first all side chains are pruned and the molecular network is obtained. Iterative removal of less characteristic peripheral rings is performed, for example, first of all the red marked aromatic ring and the linker are pruned (level 6). The remaining rings are removed one by one until a root structure remains (level 1), which should characterize the scaffold in a chemically intuitive way. Consequently, the number of levels depends on the complexity of the starting molecules [45, 46]. (This scaffold tree was calculated with Scaffold Hunter 2.6.0.)

Figure 10.9 Circular representation of the scaffold tree of marine compounds, reported in 2013 [48]. (a) To analyze huge sets of compounds according to their scaffolds, obtained through repetitive pruning, scaffolds trees can be generated to visualize the hierarchical classification of scaffolds and identify common core structures. (b) Scaffolds are represented by the leaf nodes of such a tree on each level. (c) Branching displays shared scaffolds, leading to different molecules on a higher level of the scaffold tree. The number of levels depends on the complexity of the source molecules. However, these scaffolds can be further clustered in heat maps to explore relations between properties and molecules. (This scaffold tree was calculated and visualized with Scaffold Hunter 2.6.0.)

Figure 10.10 Scaffold network of varioxepine A, a recently isolated alkaloid from marine algal‐derived endophytic fungus

Paecilomyces variotii

with a new oxa‐cage [47]. Red‐colored structures refer to the scaffolds, obtained through the scaffold tree approach. The network consists of a total number of 37 structures (5 with 5 rings, 9 with 4 rings, 9 with 3 rings, 7 with 2 rings, 7 with 1 ring).

Figure 10.11 Scaffold network of the tetracyclic‐fused alkaloid neosartin C [52], a recently isolated metabolite of the marine fungus

Neosartorya pseudofischeri

(starfish

Acanthaster planci

). Red‐colored structures refer to the scaffolds, obtained through the scaffold tree approach. The network consists of a total number of 27 structures (3 with 5 rings, 5 with 4 rings, 7 with 3 rings, 6 with 2 rings, 5 with 1 ring).

Figure 10.12 Scaffold network of granatumin Y, a limonoid recently isolated from the seeds of the Indian mangrove

Xylocarpus granatum

with a C

1

–C

29

oxygen bridge. Red‐colored structures refer to the scaffolds, obtained through the scaffold tree approach. The network consists of a total number of 27 structures (4 with 5 rings, 5 with 4 rings, 6 with 3 rings, 7 with 2 rings, 5 with 1 ring).

Figure 10.13 Comparison of structural diversity of the marine NPs with the DNP dataset and analysis of diversity within marine NPs employing SOMs. (a) SOM generated with the DNP dataset (sdf version 211.9); (b) distribution of marine NPs within the SOM generated with the whole DNP dataset; (c) SOM generated with the dataset of marine NPs, after scaffold tree generation; (d) 1‐ring molecules (244) contained in the marine NPs, obtained through scaffold tree generation, 14% coverage of marine NPs; (e) 2‐ring molecules (399) contained in the marine NPs, obtained through scaffold tree generation, 41% coverage of marine NPs; (f) 3‐ring molecules (378) contained in the marine NPs, obtained through scaffold tree generation, 62% coverage of marine NPs; (g) 4‐ring molecules (229) contained in the marine NPs, obtained through scaffold tree generation, 54% coverage of marine NPs; (h) 5‐ring molecules (149) contained in the marine NPs, obtained through scaffold tree generation, 35% coverage of marine NPs; (i) 6‐ring molecules (72) contained in the marine NPs, obtained through scaffold tree generation, 20% coverage of marine NPs; and (j) bigger than 6‐ring molecules (82) contained in the marine NPs, obtained through scaffold tree generation, 13% coverage of marine NPs. The term marine NP refers here to all 2013 reported marine NPs.

Figure 10.14 Coverage of the total structural diversity by subsets of the marine NPs, divided according to their respective ring count.

Figure 10.15 Comparison of marine NPs, processed with the scaffold tree algorithm, with fragments derived from the DNP dataset according to the respective ring count employing SOMs. (a) SOM generated with 7365 non‐flat fragment‐sized natural products according to Pascolutti

et al

.; (b) distribution of marine NPs within the SOM generated with the whole DNP dataset, 18% coverage; (c) SOM of 1‐ring‐containing DNP‐derived fragments (2636), 60% coverage; (d) SOM of 1‐ring structures generated by scaffold tree analysis of marine NP (244), 5% coverage; (e) SOM of 2‐ring‐containing DNP‐derived fragments (2822), 80% coverage; (f) SOM of 2‐ring structures generated by scaffold tree analysis of marine NP (399), 14% coverage; (g) SOM of 3‐ring‐containing DNP‐derived fragments (1621), 44% coverage; (h) SOM of 3‐ring structures generated by scaffold tree analysis of marine NP (378), 19% coverage; (i) SOM of 4‐ring‐containing DNP‐derived fragments (273), 30% coverage; (h) SOM of 4‐ring structures generated by scaffold tree analysis of marine NP (229), 17% coverage; (k) SOM of 5‐ring‐containing DNP‐derived fragments (13), 3% coverage; and (l) SOM of 5‐ring structures generated by scaffold tree analysis of marine NP (149), 12% coverage. The term marine NP refers here to all 2013 reported marine NPs.

Figure 10.16 Coverage of structural diversity by subsets of the marine NPs compared with fragments derived from the DNP, divided according to their respective ring count.

Figure 10.17 Comparison of structural diversity of marine NPs processed with the scaffold tree algorithm and through the scaffold network approach within the SOM of the whole marine NP dataset processed with the scaffold tree algorithm (Figure 10.13c). The SOM of the scaffold tree algorithm shows a 5% coverage, while with the scaffold network approach 12% of the cells are occupied by at least one structure. *The term marine NP refers here to all 2013 reported marine NPs.

Figure 10.18 Unique marine‐derived scaffolds.

Chapter 11

Figure 11.1 Worldwide occurrence of tetrodotoxin (TTX) primarily from puffer fish in marine environments. Red dots indicate origin of puffer fish from which human deaths have been recorded; black dots indicate where human TTX intoxications have occurred from toxic local fish origins, but with no recorded deaths; green dots indicate where TTX has been found in marine fauna, but no human intoxications have been associated [11–31].

Figure 11.2 Worldwide occurrence of PSP events associated with

saxitoxin

(

STX

) and analogs.

Figure 11.3 Origin and fate of tetrodotoxin (TTX) via the food web, parasitism or symbiosis, and links to

puffer fish poisoning

(

PFP

) in the marine environment.

Figure 11.4 Origin and fate of paralytic shellfish toxins (PSTs) in the marine environment indicating food web transfer vectors.

Figure 11.5 Tetrodotoxin and major naturally occurring and

a

synthetic analogs.

Figure 11.6 Saxitoxin and major naturally occurring analogs. Chemical structure of saxitoxin and analogs produced by marine dinoflagellates. Structures marked with (

a

) were not fully characterized by NMR in previous studies but their probable structures were inferred based upon mass spectrometry analysis [71].

Figure 11.7 Biochemical scheme for tailoring reactions rendering GTX 2/3, B1, and C1/2 toxins from STX.

Figure 11.8 Modified interpretation of Sako's [129] proposed STX biosynthesis pathway with incorporation of known elements of the

sxt

gene cluster in cyanobacteria.

Figure 11.9 Proposed biosynthetic pathway for STX and analogs in cyanobacteria. Dotted lines indicate additional but not essential tailoring reactions; dotted box indicates reaction only present in

Aphanizomenon

sp. NH‐5. See text for detail.

Figure 11.10 Interpretation of the tailoring reactions, leading to synthesis of sulfated carbamoyl and

N

‐sulfocarbamoyl derivatives based upon comparison of cyanobacterial elements of the

sxt

gene cluster in cyanobacteria.

Figure 11.11 Comparison

sxt

gene cluster in cyanobacteria, indicating differences in structural elements and toxin analogs between

R. brookii

D9 and

C. raciborski

T3.

Figure 11.12 Schematic representation of a nerve cell, where Na

v

channels are abundant.

Figure 11.13 Schematic representation of the α‐subunit of a Na

v

channel protein. Roman numbers (I–IV) indicate the pore domains. Each domain consists of 6 segments (indicated 1–6). Segments S5 and S6 are the pore‐forming segments. P‐loops are indicated.

Figure 11.14 Schematic representation of the α‐subunit of a Na

v

channel. In this representation, the pore and selectivity filter are formed by the segments S5 and S6 and the P‐loops (in red). The binding site 1 for guanidinium toxins is indicated among the P‐loops at the pore entrance.

Figure 11.15 Docking simulations of the benzoyl GC5b toxin coupled interactions with the Na

v

1.4 sodium channel. The ribbons show the outer vestibule of the Na

v

protein. The toxin is represented with colored sticks: light blue, carbon atoms; dark blue, nitrogen atoms; red, oxygen atoms; yellow, sulfur atoms; white, hydrogen atoms.

Figure 11.16 Different snapshots of neurotoxin I retrieved from the

Protein Data Bank

(

ID PDB

): two SH1 conformations were obtained experimentally from NMR spectra that could be obtained using MD simulations.

Figure 11.17 Representation of a di‐sulfated STX analog accessing the Na

v

channel outer vestibule.

Chapter 12

Figure 12.1 Flow chart illustrating the production of semi‐refined (a) and refined (b) carrageenans. (Adapted from McHugh 2003 [24] and Therkelsen

et al

. 1993 [136].)

Figure 12.2 Structures of the idealized disaccharide repeating units in the three main commercial carrageenans (κ‐, ι‐, and λ‐carrageenans), in their biological precursors (μ‐ and ν‐carrageenans), and in their C4 desulfated derivatives (β‐ and α‐carrageenans). Desulfation of carrageenan through the use of sulfurylases and sulfatases is described in Section 12.4.1.

Figure 12.3 Influence of cations on the gelification process of the κ‐ and the ι‐carrageenans. (Reproduced from [74], with permission of The Royal Society of Chemistry.)

Figure 12.4 500 MHz

1

H NMR spectra of κ‐ and ι‐carrageenans in D

2

O recorded at 25 °C and 70 °C clearly illustrate the effect of temperature on the spectral resolution.

1

H assignments are given in Table 12.5.

Figure 12.5 Comparison of 500 MHz

1

H (a) and

13

C (b) NMR spectra of

D

‐galactose and

D

‐galactose 4‐sulfate sodium salt in D

2

O at 25 °C. In

1

H NMR spectra (a), sulfation causes a deshielding of 0.6 ppm for the geminal proton and its effect depends on the orientation (axial/equatorial) of both groups [137]. In

13

C NMR spectra (b), substitution with a sulfate leads to a deshielding (8–11 ppm) for their linked carbon (α effect), whereas the neighboring carbons are slightly shielded (β effect) [138].

Figure 12.6 HMBC

1

H–

13

C NMR of κ‐carrageenan in D

2

O at 70 °C. In this spectrum, a correlation exists between the proton 4 of the DA unit and the carbon 1 of G unit (

3

J), indicating that these two units are linked via a 1 → 4 linkage.

Figure 12.7 Origin of characterized carrageenan‐modifying enzymes.

Figure 12.8 Carrageenan‐modifying enzymes isolated from

P. carrageenovora

(A and B) and from

P. atlantica

(C and D). (A) Biodegradation of κ‐carrageenan. In this scheme, the sulfatase catalyzes the desulfation of the κ‐carrageenase end products (DP2 and DP4). As it is an

exo

‐enzyme, it acts only on the sulfate located at the non reducing ends [111]. (B and C) Bioconversion of the ι‐carrageenan into α‐carrageenan through the action of the 4S‐endosulfatase from

P. carrageenovora

(B) [112] or from

P. atlantica

(Q15XH3) (C). As

endo

‐enzymes, these sulfatases are able to catalyze the removal of any sulfate located on the C4 position of the G units in ι‐carrabiose moieties. Note that Q15XH3 from

P. atlantica

is also able to use the oligo‐ι‐carrageenans as substrates [113]. (D) Conversion of κ‐ into hybrid κ‐/β‐carrageenan through the action of the sulfatases Q15XH1 and Q15XG7 from

P. atlantica

T6C. *These enzymes have an

endo

‐mode of action and can also use κ‐/μ‐ and oligo‐κ‐carrageenan as substrates [114].

Figure 12.9 Bioconversion of ι‐carrageenan into α‐carrageenan using the ι‐carrageenan sulfatase from

P. carrageenovora

. The conversion was followed as a function of time by

1

H NMR. The ι‐carrageenan (top spectrum) is completely converted into α‐carrageenan (bottom spectrum) after 120 h of incubation with the 4S‐ι‐carrageenan sulfatase [112].

Figure 12.10 Uses of oligo‐carrageenans for their biological properties. Note that this list is not exhaustive, as the subject is vast and could be the topic of a paper by itself.

Chapter 13

Figure 13.1 Structure glycopeptides: vancomycin and teicoplanin.

Figure 13.2 Structure of lipopeptides: daptomycin and ramoplanin.

Figure 13.3 Structure of new lead antibiotic peptides [4].

Figure 13.4 Structure of andrimide A [56].

Figure 13.5 Structures of solonamides, unnarmicins, and ngercheumicins [58].

Figure 13.6 Structure of kahalalide F [59].

Figure 13.7 Structure of mojavensin A [63].

Figure 13.8 Structure of bacillistatins [64].

Figure 13.9 Structure of thiopeptide TP‐1161 [73].

Figure 13.10 Structure of kocurin [74].

Figure 13.11 Structure of marthiapeptide A [76].

Figure 13.12 Structure of peptidolipins [78].

Figure 13.13 Structure of Sungsanpin [79, 80].

Figure 13.14 Structure of ohmyungsamycins [79, 80].

Figure 13.15 Structures of pitiprolamide and pitipeptolides [83, 84].

Figure 13.16 Structure trichorzianine peptaibols [86].

Figure 13.17 Structure of trichoderin [87].

Figure 13.18 Structures of new compounds derived from genome mining, genome scanning, and genomisotopic approach.

Chapter 14

Figure 14.1 Important

Conus

spp. present in the Indian coast.

Figure 14.2 Distribution of Cone snails in the Indian coast.

Figure 14.3 (a) The morphological and (b) venom duct anatomical features of molluscivorus cone snail

Conus araneosus

, Gulf of Mannar, India.

Figure 14.4 Basic scheme showing the workflow for the isolation of peptide and screening of bioactive compounds from

Conus

spp.

Chapter 15

Figure 15.1 Examples of some typical chemical reactions on polysaccharides.

Figure 15.2 1,3‐Huisgen cycloaddition reaction (“click” reaction).

Figure 15.3 The “grafting from” and the “grafting onto” techniques (Adapted from Riva

et al

. [56]. Reproduced with permission of Springer.)

Figure 15.4 Schematic representation of water/gel interactions.

Figure 15.5 Diffusion of a drug from a hydrogel.

Figure 15.6 Schematic representation of the methods described in the literature for gel particle technology: (a) dropping the polysaccharide solution into a solution of small ions, (b) via a water‐in‐oil emulsification technique, and (c) complexation of oppositely charged polyelectrolytes. (Borgogna

et al

. [63]. http://www.mdpi.com/1660‐3397/9/12/2572/htm. Used under CC BY 3.0 https://creativecommons.org/licenses/by/3.0/.)

Figure 15.7 Schematic representation of a bioactive hydrogel dressing.

Figure 15.8 Structures of the three main types of carrageenans.

Figure 15.9 Mechanism of gelation of carrageenans. The associated counterions are required to induce sol/gel transition. λ‐Carrageenan has a structure that does not allow double‐helix formation. (Chuna and Grenha [69]. http://www.mdpi.com/1660‐3397/14/3/42/htm. Used under CC BY 4.0 https://creativecommons.org/licenses/by/4.0/.)

Figure 15.10 Possible arrangements of the M and G monomers in sodium alginate: homogeneous G–G sequence, homogeneous M–M sequence, and heterogeneous M–G sequence. (From Ref. [92].)

Figure 15.11 Gelation mechanism of alginate with the “egg‐box” model.

Figure 15.12 Chemical structure of chitin. The main difference regarding cellulose is at carbon 2, where the hydroxyl group on cellulose is substituted by acetamide group (circled) in chitin.

Figure 15.13 Extraction process, preparation, and chemical structures of chitin and chitosan. (Adapted from Suginta

et al

. [138]. Reproduced with permission of American Chemical Society.)

Figure 15.14 A diagrammatic presentation of wound healing process.

Figure 15.15 (a) General principles of periodate oxidation of polysaccharides, where cleavage occurs between C2 and C3. (b) Periodate oxidation of chitosan. The nitrogen at C2 is released as ammonia.

Figure 15.16 Grafting of alkyl chains by reductive amination. (Adapted from Riva

et al

. [56]. Reproduced with permission of Springer.)

Figure 15.17 Synthesis of

N,N,N

‐methyl chitosan chloride. (Adapted from Riva

et al

. [56]. Reproduced with permission of Springer.)

Figure 15.18 Synthesis of acyl chitosan. (Adapted from Riva

et al

. [56]. Reproduced with permission of Springer.)

Figure 15.19 Synthesis of

O

‐succinyl chitosan. (Adapted from Zhang

et al

. [172]. Reproduced with permission of Elsevier.)

Figure 15.20 (a) Synthesis of chitosan–Pluronic copolymers. (Adapted from Park

et al

. 2008 [179]. Reproduced with permission of Elsevier.). (b) Hydrogel formation via hydrophobic interaction of Pluronic chains. (Adapted from Park

et al

. [180]. Reproduced with permission of Elsevier.)

Chapter 16

Figure 16.1

Haslea ostrearia

with marennine pigment accumulated in the apical areas of the cells. Scale = 50 µm.

Figure 16.2 Effect of pH on the color of intracellular marennine (IMn).

Figure 16.3 Clearance rate (CR) of oysters

Crassostrea gigas

exposed to EMn (green) compared with the control. Values are means ± standard error (

n

 = 6).

Figure 16.4 Scheme used for the greening experiments. Oysters (

Crassostrea gigas

) were maintained in chambers containing (a) filtered (0.2 µm) artificial seawater with

Skeletonema costatum

cells (control); (b) culture of

Haslea ostrearia

with purified and lyophilized extracellular marennine (EMn); (c)

H. ostrearia

supernatant (crude extract) containing EMn, without

H. ostrearia

cells, but with

S. costatum

cells; and (d) filtered artificial seawater containing

H. ostrearia

cells only, without any added marennine.

Figure 16.5 Effect of oyster (

Crassostrea gigas

) treatments on the coloration of gills. (a) Oyster fed with

Skeletonema costatum

as control; (b) oyster exposed to

Haslea ostrearia

and EMn containing supernatant; (c) oyster exposed to EMn containing supernatant; and (d) oyster exposed to

H. ostrearia

cells only. Scale = 1 cm.

Figure 16.6 Views of whole‐mount gills of oysters (

Crassostrea gigas

) showing marennine‐colored mucocytes (M) in ordinary filaments (O), which constitute gill plicae (P). Note that principal filaments are not visible (scale bars, 500 μm and 100 μm).

Figure 16.7 Raman spectra at 514.5 nm obtained on the different organs (labial palps, gills, digestive gland, mantle) of (a) control oysters and (b) green oysters.

Figure 16.8 Influence of pH on the wavelength of intracellular marennine (IMn) maximum absorbance (λ

max

).

Figure 16.9 Position of intracellular marennine (IMn) at pH 7 (squares), pH 2.5 (diamonds), extracellular marennine (EMn) pH 7 (dots), pH 2.5 (shuttle) in the CIE

L

*

a

*

b

* colorimetric system (standard observer CIE 10°), with different illumination: standard daylight D65 (filled symbols), tungsten lamp A (hatched), fluorescent lamp F2 (empty).

Figure 16.10 Color characteristics of green oyster gills. Projection in a (

a

*,

b

*) plane of CIE

L

*

a

*

b

* colorimetric system of the (

a

*

b

*) sets obtained for control (diamonds) and green (filled square) oyster gills under standard daylight D65 illumination.

Chapter 17

Figure 17.1 Outline of NRPS and PKS biosynthesis and associated tailoring modifications. Mechanism of one iteration of NRPS

(A)

or PKS

(B)

chain extension. A hypothetical compound, tethered to a thioesterase (TE), highlighting the structural diversity generated by NRPS and PKS tailoring domains

(C)

. MT: methyltransferase; E: epimerase; Cy: cyclisation‐condensation; Ox: oxidase; DH: dehydratase; KR: ketoreductase; ER: enoylreductrase; AMT: aminotransferase; Hal: halogenase.

Figure 17.2 Overview of NRPS/PKS structures. Numbering of structures corresponds to the following respective sections (17.2.1.x), which provide an overview of each specialized metabolite.

Figure 17.3 Marine cyanobacterial ribosomally synthesised (RiPP) natural products.

10

patellamide A;

11

prochlorosin. The lanthipeptide (

11

) undergoes posttranslational modification by the ProcM tailoring system, which cyclises the peptide through the formation of thioether crosslinks (C‐S‐C).

Chapter 18

Figure 18.1 Protonation equilibria of ovothiol A. Shown in the box is the doubly zwitterionic form prevailing at physiological pH values.

Figure 18.2 Flowchart of ovothiol A purification from

P. lividus

eggs.

Figure 18.3 Proton NMR and mass spectra of ovothiol A.

Figure 18.4 Chemical synthesis of ovothiol A as pure enantiomer and racemate.

Figure 18.5 Chemical synthesis of enantiomeric pure ovothiol A.

Figure 18.6 Biosynthetic pathway of ovothiol formation.

Figure 18.7 Comparison between the reactions catalyzed by OvoA (a) and EgtB (b). OvoA and EgtB catalyze the formation of the respective sulfoxide intermediates, 5‐histidylcysteine sulfoxide and trimethylhistidylglutamylcysteine sulfoxide from histidine and cysteine (a) and trimethyl histidine and gamma‐glutamyl cysteine (b), respectively.

Figure 18.8 Proposed model for ovothiol function in sea urchins. (a) At fertilization ovothiol protects sea urchin eggs from the deleterious effects of H

2

O

2

, acting as a nonenzymatic glutathione peroxidase. (b) During development, ovothiol protects developing embryos from oxidative stress produced by environmental cues.

Figure 18.9 Synthesis of ovothiol derivatives.

Figure 18.10 Ovothiol diselenide derivatives.

Chapter 20

Figure 20.1 Underwater photograph taken in the waters of Stradbroke Island, Australia (May 2016), showing a wide diversity of pigmented organisms, including crinoids, soft corals, sponges, and algae. Although marine biodiversity is colorful, it is not always easily accessible, meaning that oceans are a treasure chest of untapped pigment diversity.

Figure 20.2 Compliance of photosynthetic pigments with Lipinski's rules of five. Predicted data were generated by ACD/Labs Percepta Platform – PhysChem Module and retrieved online from ChemSpider (http://www.chemspider.com). Information retrieved were (a) average mass (Da), (b) number of hydrogen bond donors (HB

Don

), (c) number of hydrogen bond acceptors (HB

Acc

), and (d) calculated partition coefficient between octanol and water (Log

P

value). A non‐exhaustive subset of marine pigments was analyzed containing 15 chlorophylls, 3 phycobilins, and 38 carotenoids, which can be detected notably in phytoplankton and cyanobacteria species. Compliance is marked by green area, while violation is represented in red area.

Figure 20.3 Applications of marine pigments. This figure provides a broad idea of the attrition rate between potential and industrial applications of pigments. However, this is subject to variations since policy can be different according to the countries. (1) Astaxanthin can be used in skin care against photoaging, (2) lutein and zeaxanthin found application against AMD, (3) some pheophorbides, chlorines, and bacteriochlorophylls are used for photodynamic treatments, (4) pigment‐enriched diet benefit to all stage of rearing, (5) seaweed or microalgae powder containing β‐carotene can be consumed as antioxidant, (6) C‐phycocyanin extract can be included in ointments for its antioxidant properties, (7) zeaxanthin can be used to prevent AMD, (8) R‐phycoerythrin is used for immunolabeling, (9) astaxanthin and canthaxanthin are used to color salmon flesh through pigment complementation diet, (10) some carotenoids and phycobiliproteins can be integrated to topical ointments and dairy products but chlorophylls are sometimes too labiles for industrial uses.

Figure 20.4 Flowchart for sourcing, production, and valorization of pigments. This simplified workflow for production of valuable natural ingredients/molecules from idea to commercialization is applicable to pigments and other natural products. Chemodiversity is source of valuable compounds or ingredients. Pigments projects are well adapted to follow this kind of scheme collaboration between academia and industry. Peer‐reviewed publications are essential for academic researchers and this kind of valorization is possible along the project and besides patents possibilities.

Chapter 21

Figure 21.1 Solar spectrum showing ranges of ultraviolet radiation (UVR), photosynthetically active radiation (PAR), and infrared (IR) radiation.

Figure 21.2 Photographs showing the filaments of

Scytonema

sp. with sheaths containing yellow‐brown UV‐protective pigment, scytonemin.

Figure 21.3 Chemical structures of different forms of scytonemin. (a) Chemical structures of scytonin, reduced scytonemin, oxidized scytonemin and scytonemin‐3a‐imine. (b) Chemical structures of dimethoxy scytonemin, tetramethoxy scytonemin, and scytonemin A.

Figure 21.4 Shikimate pathway for MAAs biosynthesis and their possible interconversion. Broken line represents the putative biosynthetic connection between dehydroquinate, gadusols, and MAAs.

PEP

,

phosphoenolpyruvate

;

DAHP

,

3‐deoxy‐

D

‐arabinoheptulosinate‐7‐phosphate

; DHQ, dehydroquinate;

EPSP

,

5‐enolpyruvylshikimate‐3‐phosphate

.

Figure 21.5 Genes involved in MAAs biosynthesis.

Figure 21.6 Biosynthetic route for the scytonemin and corresponding gene products involved in each step. Functionally characterized gene products are represented by continuous arrow while gene products indicated by broken arrow are still to be functionally characterized for their involvement in corresponding step.

Figure 21.7 Structure of genome associated with biosynthesis of scytonemin in

N. punctiforme

. The hatch marks indicate a break in the distance scale.

Figure 21.8 Absorption spectrum of scytonemin having peaks at 252, 300, and 386 nm.

Figure 21.9 Absorption spectrum of MAAs having peak at 334 nm.

Figure 21.10 Various applications of UV‐screening compounds: MAAs and scytonemin.

Chapter 22

Figure 22.1 Phylogenetic tree of living organisms. (a) Position of the three groups of macroalgae, Ulvophyceae, Rhodophyta, and Phaeophyceae in the tree of life (modified from [2]). (b) Illustration of the diversity in forms, colors, and reproduction of diverse macroalgae all around the world.

Figure 22.2 Some typical polysaccharides structuring the cell wall of marine brown macroalgae: structural anatomy of the cell wall, chemical structure of both fucoidan and alginate together with the hydrogel formed by alginates.

Figure 22.3 Chemical structure of some particular mycosporine‐like amino acids (MAAs) and phlorotannins isolated from red and brown macroalgae. In the case of fuhalol, the additive hydroxy group is visible by the dotted square.

Chapter 23

Figure 23.1 Chemical structures of representative current local anesthetics: (

1

) lidocaine, (

2

) bupivacaine, and (

3

) procaine. Compounds

1

and

2

are amino amide analgesics, while

3

is an amino ester analgesic.

Figure 23.2 Chemical structures of major saxitoxin congeners.

Figure 23.3 Chemical structures of initial planar structure proposals for saxitoxin (

1

and

2

) and final structure

3

as determined by crystal structure analysis.

Figure 23.4 Members of the brevetoxin class of compounds and biosynthetically related antitoxin brevenal.

Figure 23.5 Representative members of other dinoflagellate ladder‐frame polyether toxins.

Figure 23.6 Representative compounds of cyclic imine toxins, in addition to the gymnodimine and spirolide toxins.

Figure 23.7 Chemical structures of known members of the gymnodimine family. (a) GYM A congeners; (b) GYM B and C congeners; and (c) di‐tetrahydrofuran congener GYM D.

Figure 23.8 Chemical structures of several members of the spirolide toxin family. (a) Tri‐spiroketal congeners; (b) free amine congeners; and (c) di‐spiroketal congeners. Δ

2,3

refers to dihydrolactone analogs.

Chapter 24

Figure 24.1 Four marine luciferins (top) and firefly luciferin (bottom). The latter is not of marine origin, but is shown because it is the most well‐known luciferin.

Figure 24.2 The immature precyclized protein (left) undergoes a fast autocatalyzed cyclization, followed by a slower oxidation to form the mature chromophore (right).

Figure 24.3 Structure of green fluorescent protein (GFP). The chromophore (green) is formed in the autocatalytic cyclization shown in Figure 24.2.

Figure 24.4 Phototransformable

fluorescent protein

s (

FP

s) can be divided into photoconvertible, photoactivatable, and photoswitchable FPs. (a) In the photoconvertible fluorescent proteins (e.g., EosFP), the photoinduced cleavage of the backbone extends the conjugation of His62. (b) Photoactivated fluorescent proteins such as PA‐GFP are irreversibly photoactivated when Glu222 is decarboxylated and the chromophore forms the fluorescent phenolate form. (c) In the photoswitching Dronpa protein, the chromophore can be reversibly switched between the dark

trans

and fluorescent

cis

forms.

Figure 24.5 The crystal jellyfish (

Aequorea victoria

) (a) has about 300 photoorgans (b) on the bottom edge of its umbrella. These can emit green light using both aequorin and green fluorescent protein (GFP). (c) The green eel (

Kaupichthys hyoproroides

), found in the waters of the Bahamas, has some fluorescent proteins. (d) CaMPARI fluorescence in a larval zebrafish brain, showing active neurons (magenta) that were marked while the fish was swimming freely. (e) Rainbow fluorescent proteins highlighting Purkinje neurons that reach their arbor‐like dendrites into the molecular layer of the developing cerebellum of a mouse. (f) The portrait “Facing the Light” was first displayed at the “125 years of Albert Einstein” exhibition of the University of Ulm, which commemorated his 125th birthday. Jörg Wiedenmann and Franz Oswald created the images by taking a black‐and‐white image of Albert Einstein (top‐left Einstein) and then covering it with a nitrocellulose membrane that had EosFP immobilized on it (top‐right Einstein). EosFP, named after the goddess of dawn, is a photoconvertible fluorescent protein. Initially it fluoresces green (bottom‐left Einstein), but irradiation with UV light (390 ± 30 nm) induces cleavage between the amide nitrogen and the α‐carbon atom in the histidine adjacent to the chromophore, resulting in a red fluorescent form (bottom‐right Einstein).

Chapter 25

Figure 25.1 Number of citations in PubMed (https://www.ncbi.nlm.nih.gov/pubmed) containing the term “marine bacteria.”

Figure 25.2 Bacterial biofilm grown on a glass slide and observed from the top by scanning electron microscopy. The image was artificially colored, showing bacteria in blue and the matrix in yellow‐green (LBCM, Université de Bretagne‐Sud, France).

Figure 25.3 Effect of the

Pseudoalteromonas

3J6 culture supernatant (SN

3J6

) on the attachment of

Vibrio tapetis

onto glass and on biofilm formation by

Pseudomonas aeruginosa

,

Salmonella enterica

,

Escherichia coli

, and

V. tapetis

. Confocal laser scanning microscopy observations after the attachment step (top views) and biofilms (top, side, and 3D views) grown for 24 h (

V. tapetis

) or 48 h (

P. aeruginosa

,

S. enterica

, and

E. coli

) [52, 70].

Figure 25.4 (a) Oxidative deamination of

L

‐lysine by the antimicrobial enzyme LodA. CTQ is the cofactor of LodA; B represents an active site residue that participates in acid–base chemistry [108]. (b) Structure of the CTQ cofactor [109].

Chapter 26

Figure 26.1 Structural components of chitin are represented by glucosamine (C

6

H

13

NO

5

) (a), which consequently is to be found in the homopolymer chain of β‐1,4‐linked

N

‐acetylglucosamine (b). Such chains can be specially oriented when within 2 nm large crystalline unit of chitin (c).

Figure 26.2 Schematic view of the common structural concept of three‐dimensional demosponge chitin as a scaffold (a) that is naturally constructed from tubulous nanofibrous chitin (b).

Figure 26.3 Chitin as a structural component within the skeletal frameworks of glass sponges

Aspidoscopulia

sp. (a) and

Euplectella aspergillum

(c) can be initially visualized using fluorescence microscopy (b and d, respectively) after Calcofluor white staining.

Figure 26.4 Alkali‐based treatment of highly flexible anchoring spicules of glass sponge

S. hawaiicus

(e) led to the obtaining of an alkali‐resistant chitin‐based layer (a) with complex nanostructural organization (b). HR‐TEM investigations show crystalline 2‐nm large nanostructures (c, arrows) typical for crystalline subunits of chitin. It is suggested that nanophase of amorphous silica is localized between nanofibers of chitin (d).

Figure 26.5 The common freshwater

Spongilla lacustris

demosponge (a) contains chitin within both the gemmules (b) and the body skeleton (c, d).

Figure 26.6 Proteinaceous, mineral‐free spongin‐based skeletal fibers of bath sponges (subclass Ceractinomorpha) are monolithic and remain in a “spaghetti‐like” morphology (a). However, chitin‐based skeletal fibers of all sponges related to Verongida order are mineralized with silica and calcium carbonates and show inner tubulous “macaroni‐like” morphology (b).

Figure 26.7 Skeletal fibers (arrows) of Verongida sponges are up to 150 µm in diameter and can be observed even when fixed in ethanol or dried in air or when sponges have been cut or broken. Such structures are well seen on examples of

Suberea

sp. (a),

A. cavernicola

(b),

A. fulva

(c),

A. fistularis

(d), and

A. cauliformis

(e, f).

Figure 26.8 Partial demineralization of selected fragments of Verongida sponges related to the Aplysinidae family (a, b) led to the disruption of the cells that are naturally located within mesohyl. After that, the distribution of skeletal fibers within the sponge body became visible (c, d).

Figure 26.9 Schematic view of the principal structure of the

Ianthella basta

sponge skeleton. Initially visible as two‐dimensional (a, b), the fiber‐based skeleton of this sponge is three‐dimensional (c).

Figure 26.10 A typical representative of Verongida sponges,

Verongula gigantea

(a), contains reticulate fibrous skeleton made of mineralized chitin. Acidic treatment of such fibers led to the dissolution of calcium carbonate (b). After those, acid‐resistant mineral layers that have been identified as silica–chitin composites became visible using SEM (c) and light microscopy (d, e). Chitin‐based skeletal fibers of verongiids are examples of naturally occurring multiphase biomineralized structures (f).

Figure 26.11 Schematic view of the step‐by‐step acidic/alkali treatment procedure for the isolation of chitinous scaffolds from skeletons of Verongida sponges.

Figure 26.12 Treatment of partially demineralized but still pigmented skeletal sponge fibers with 35% hydrogen peroxide during 15 min at room temperature (a) led to obtaining translucent chitinous fibers (b).

Figure 26.13 Very rigid

A. cauliformis

sponge fragment (3‐cm diameter) (a) has been apically demineralized and became discolorated after 120 h insertion in a 2.5 M NaOH solution. This mineral‐ and pigment‐free fragment (b) can be cut into pieces of desired size and shape for uses in tissue engineering and biomedicine as microporous 3D scaffolding (c, d).

Figure 26.14 Cross sections of Verongida sponge skeletal fibers confirm the presence of up to 50 µm in diameter large channels (a–c). This feature remains stable in purified sponge chitin scaffolds (d) obtained after demineralization of the natural skeleton.

Figure 26.15 Chitinase digestion of chitinous fiber (a) isolated from

Aplysina fistularis

. Microfibers became visible in light microscopy after 2 and 4 h of treatment with chitinase (b, c), respectively.

Figure 26.16 Multilayered structures of the verongid skeletal fibers can be observed using light microscopy (a) and SEM (b, c). Numerous alternating nanolayers of chitin are visible on cross section of demineralized individual fiber of

V. gigantea

sponge also using TEM (d).

Figure 26.17 SEM image of an alkali‐treated (a) and mechanical‐disrupted naturally occurring sample of Verongida sponge fiber (b) showing their nanofibrillar organization.

Figure 26.18

Aplysina aerophoba

sponge colony cultivated under marine ranching conditions in Kotor Bay, Montenegro.

Figure 26.19 Three‐dimensional, nondemineralized but cell‐free skeletons of different representatives of the Aplysinidae family (a–c) resemble the shape and morphology of the native sponges. These skeletons are rigid enough to be used for mechanical tests. The yellow‐brownish color of skeletons is determined by the presence of bromotyrosines that possess inhibitory activity against chitinases.

Figure 26.20 The flat 3D skeleton of

Ianthella basta

sponge (a) possesses characteristic macroporous membrane‐like fibrous morphology (b, c). The semisquare architecture of isolated chitinous scaffolds (d) is based on the mechanical stability of individual fibers (e) and nearly exactly resembles the shape and dimension of original

I. basta

skeleton.

Figure 26.21 The large size of naturally occurring

Ianthella basta

sponges (a) allows the isolation of corresponding large fragments of chitin‐based scaffolds (b). These were used in cultivation of hMSCs. The viability of hMSCs (c), as well as adipose tissue cells (d) on

I. basta

chitinous scaffolds, was very high.

Figure 26.22 Extreme biomimetically synthesized Chitin‐germanium oxide composite material was obtained via extreme biomimetic route. This material resembles the form and shape of

Aplysina cauliformis

chitinous scaffold. It is suggested that chitin interacts with germanium oxide nanoparticles only, by formation of hydrogen bonds [95].

Figure 26.23 Light microscopy image of the

Aplysina cauliformis

sponge chitin scaffolds inserted into a 47% NaOH solution at 85 °C shows that the outer layers of this chitin can be transformed into chitosan (a, arrows). No structural changes have been observed also using fluorescence microscopy (b).

Chapter 27

Figure 27.1 Major biogenic sources of marine biominerals, with age estimates of earliest fossil records. (a) carbonate‐based marine biominerals; (b) silica‐based marine biominerals; (c) heavy metal bioaccumulations.

Figure 27.2 Mollusc shells. Examples of diversity of shapes, textures, colors, and patterns. (a) Blue bubble shell (

Janthina janthina