# Various Approaches to Fourier-Transform Infrared Spectroscopy (FTIR) for Bioanalytical and Biotechnological Applications in Marine Algae

Elando Fréda Zamanileha, Julie Tantely Mitantsoa, Picardino Frienduc

Vaonalamihanta, Andriamanarivosoa Rija Razafintsalama, Fara Arimalala

Andrianony, Ravelonandro Pierre Hervé

► To cite this version:

Elando Fréda Zamanileha, Julie Tantely Mitantsoa, Picardino Frienduc Vaonalamihanta, Andriamanarivosoa Rija Razafintsalama, Fara Arimalala Andrianony, et al.. Various Approaches to Fourier-Transform Infrared Spectroscopy (FTIR) for Bioanalytical and Biotechnological Applications in Marine Algae. Mazumder Nirmal; Zhuo Guan-Yu. Recent Advances in Infrared Spectroscopy and Its Applications in Biotechnology, IntechOpen, In press, 978-0-85014-921-0. 10.5772/intechopen.1006723. hal-05117841

HAL Id: hal-05117841

https://hal.science/hal-05117841v1

Submitted on 18 Jun 2025

HAL is a multi-disciplinary open access
archive for the deposit and dissemination of sci-
entific research documents, whether they are pub-
lished or not. The documents may come from
teaching and research institutions in France or
abroad, or from public or private research centers.

L'archive ouverte pluridisciplinaire HAL, est
destinée au dépôt et à la diffusion de documents
scientifiques de niveau recherche, publiés ou non,
émanant des établissements d'enseignement et de
recherche français ou étrangers, des laboratoires
publics ou privés.

<figure><img src="image_1.png" /></figure>

Distributed under a Creative Commons Attribution 4.0 International License
# Various Approaches to Fourier-Transform Infrared Spectroscopy (FTIR) for Bioanalytical and Biotechnological Applications in Marine Algae

Elando Fréda Zamanileha, Julie Tantely Mitantsoa,

Picardino Frienduc Vaonalamihanta,

Andriamanarivosoa Rija Razafintsalama,

Fara Arimalala Andrianony and Pierre Hervé Ravelonandro

## Abstract

In recent decades, infrared spectroscopy has proved to be an effective tool for studying biological molecules, thanks in particular to advances in Fourier transform infrared (FTIR) spectroscopy. This method is widely used to characterize molecules, providing precise data on samples and finding extensive applications in biological and biotechnological research. Macroalgae, alongside marine crustaceans, are now recognized as primary sources of biomass, offering nutritional benefits and promising bioactive molecules. Hydrocolloids extracted from algae play a crucial role as food additives, improving the texture and quality of food products. This chapter explores in detail the fundamentals, data analysis methods, spectrum interpretation, and associated aspects of characterizing hydrocolloids present in the cell wall of algae using FTIR spectroscopy. It provides an in-depth overview of the methodologies used to extract these hydrocolloids from various types of algae while optimizing analytical techniques and interpreting infrared results. By enriching our understanding of these compounds, this chapter aims to promote their use in diverse fields such as food and biotechnology, highlighting their growing potential as sustainable and biologically beneficial resources.

**Keywords:** FTIR, hydrocolloids, infrared, macroalgae, polysaccharides, spectra

## 1. Introduction

The evolution of chemical analysis has been driven by Fourier transform infrared (FTIR) spectroscopy, which uses light to analyze samples [1]. By scattering all infrared wavelengths simultaneously, the sampling speed has increased significantly.
FTIR is commonly used to analyze the biochemical properties of biological samples such as proteins, cellular materials, and tissues [2], providing unbiased and detailed data on the vibrational fingerprint of organic compounds [3]. This method has been widely used in biomedical and biotechnology research. FTIR can be used to study a wide variety of materials at macro and micro levels. It is crucial to choose the right spectra acquisition geometries and sample preparation methods, especially for cells and tissues. The attenuated total reflection (ATR) mode facilitates analysis by avoiding sample preparation, thanks to a stationary evanescent wave [1]. In catalysis, FTIR is essential for identifying compounds in complex mixtures [4], monitoring catalytic activity in real time, and determining the active phases of catalysts [5, 6]. It analyzes catalyst surfaces and identifies chemisorbed species [7]. Historically, transmission-mode IR was used to study the acid-base properties of materials using probe molecules [8]. This technique is still preferred for obtaining detailed information on the composition and mechanisms of catalysts [9, 10].

Over 70% of the Earth's surface is covered by the ocean [11], home to a vast array of marine organisms producing numerous natural products [12]. Among them, marine algae stand out for their unique functional compounds [13], often absent from land plants. These compounds, with their antioxidant, anti-inflammatory, antibacterial, and antiviral properties, are of great interest for pharmaceutical, cosmetic, and food applications [14, 15]. The economic importance of seaweed farming has risen sharply in recent years [16], becoming the second most important form of aquaculture after fish farming [16]. The main drivers are the health benefits of seaweed and its potential as a sustainable source of bioenergy and biomaterials. Intensively cultivated macroalgae sequester carbon and help combat climate change [17, 18]. They create jobs in coastal communities [19], reduce pressure on wild fish stocks, and improve water quality by absorbing excess nutrients [19, 20]. The production of hydrocolloids, gelling, and thickening substances extracted from algae, such as agar-agar, carrageenan, and alginate, is also booming. Used in the food, medical, and even construction industries, these hydrocolloids are environment-friendly alternatives to synthetic products, contributing to a more sustainable economic cycle and a reduced carbon footprint.

Full characterization of the FTIR spectra of marine algae is fundamental to exploring their complex chemical composition and potential applications in various sectors [3]. This spectroscopic technique uses Fourier transform infrared (FTIR) spectroscopy to identify and quantify the biochemical components of algae based on their distinctive spectral signatures [21–23]. The basic mechanisms rely on the absorption of infrared (IR) energy by specific molecular bonds, such as the O-H bonds of polysaccharides and the C=O bonds of lipids, generating characteristic peaks in the infrared spectrum [24, 25]. Before FTIR analysis, algae samples are prepared by drying and powdering to ensure homogeneous distribution during measurement [26]. This allows rapid, non-destructive assessment of algae composition, essential for research into their biodiversity, nutritional properties, and potential industrial applications [26]. Indeed, it is used in the development of algae-based food, cosmetics, and pharmaceutical products, guaranteeing the quality and safety of finished products [27]. Although FTIR offers significant advantages such as sensitivity and speed, it requires expertise to correctly interpret spectra and optimize sample preparation.

This chapter presents the fundamental principles, data analysis process, spectral interpretation, and related information for characterizing algal cell wall hydrocolloids via Fourier transform infrared (FTIR) spectroscopy. It provides an overview of the essential data needed to understand hydrocolloid extraction methods from various
algal phyla, as well as to optimize analytical techniques and interpretations of IR spectra. The aim is to provide an in-depth understanding of the methods studied, facilitating the characterization and effective use of algal hydrocolloids.

## 2. Principle of IR spectroscopy

### 2.1 Fundamentals of FTIR

The Fourier transform infrared spectroscopy (FTIR) is an essential method for the analysis of materials, whether organic, polymeric, or inorganic [15]. This technique measures the absorption of infrared radiation by a sample at different wavelengths, producing a distinct spectrum that reveals the sample's molecular fingerprint. As infrared radiation passes through the sample [26], some frequencies are absorbed by chemical bonds, while others pass through. To modulate the IR radiation, a Michelson interferometer is used, generating an interferogram containing all the necessary spectral data [28]. This interferogram is then Fourier-transformed into an absorption spectrum.

An FTIR spectrometer consists of an infrared source, an interferometer, a sample holder, a detector, and a computer for data processing. Depending on the state of the sample, preparation varies—solids are prepared with KBr pellets or thin films, liquids with cells equipped with IR-transparent windows, and gases with special cells. Analysis of FTIR spectra enables functional groups and molecular structures to be identified by observing characteristic peaks. The fingerprint region, which lies between 400 and 1500 cm⁻¹, is particularly useful for accurate compound identification. The FTIR has several advantages, including rapid spectral acquisition, high sensitivity, and non-destructive analysis requiring little sample preparation [29]. The method is widely used for qualitative and quantitative analysis, material characterization, and environmental monitoring, making it a versatile tool in a variety of scientific fields [26].

### 2.2 Basic principles of FTIR

Infrared (IR) electromagnetic radiation does not possess sufficient energy to excite electrons in molecules, unlike UV-visible radiation or X-rays (Figure 1). However, it is capable of intensifying the molecular and rotational vibrations of chemical bonds. This ability to interact with molecular vibrations is exploited in infrared (IR) spectroscopy. IR spectroscopy covers two main regions of the spectrum, depending on photon energy (Figure 1) [30]:

*   **Far-IR:** Corresponding to longer wavelengths (lower energy, around 400,000 nm), wavelength region (400 to 10 cm⁻¹), this region mainly quantifies changes induced by rotational vibrations of molecules.

*   **Near-IR:** Located at shorter wavelengths (around 700 nm), the wavelength region is 4000 to 700 cm⁻¹. This region encompasses both the vibrational and rotational vibrations of molecules.

*   **Mid-IR:** Located at wavelengths of 2–25 µm. Wavenumber region 4000 to 400 cm⁻¹. Analysis of the fundamental vibrations of chemical bonds, enabling precise identification of functional groups in organic and inorganic molecules.
<figure><img src="image_2.png" /><figcaption>**Figure 1.**
Infrared region of the electromagnetic spectrum [30].</figcaption></figure>

The frequency of absorption in the IR spectrum depends on the different modes of molecular vibration, such as symmetrical and asymmetrical stretching, bending, and so on. The intensity of absorption peaks depends on the efficiency with which molecules absorb infrared energy, which in turn depends on the variation in the molecule's dipole moment [26]. Symmetrical molecules such as N₂, H₂, and O₂, which are diatomic gases, do not possess a permanent dipole moment and, therefore, do not show a characteristic spectral fingerprint in the IR spectrum [26]. This makes IR spectroscopy particularly useful for identifying specific compounds with functional groups with distinct dipole moments, such as OH bonds in alcohols or amide groups in proteins. IR spectroscopy is a valuable technique for analyzing molecular vibrations and characterizing chemical structures due to its sensitivity to subtle variations in molecule vibrational modes [31].

Molecular vibrations in the infrared (IR) spectrum are essential for analyzing molecular structure, falling into two main types: stretching and bending. Stretching vibrations include symmetric stretching, where atoms move symmetrically for the bond axis, altering its length without changing its angle, and asymmetric stretching, where one atom moves toward the central atom while the other moves away, creating an asymmetric variation in bond length (Figure 2) [26].

The bending vibrations are divided into several modes: rocker bending, scissor bending, oscillation bending, and torsion bending. Rocker bending occurs when atoms move in a plane perpendicular to the central bond, altering the angle of the bond without changing its length. Scissor bending resembles the movement of a pair of scissors, altering the angle between bonds. Oscillation bending involves an oscillating movement in the plane of the molecule. Torsional bending is characterized by a twisting motion of the atoms outside the plane of the molecule, resulting in rotation
<figure><img src="image_3.png" /><figcaption>**Figure 2.**
Schematic diagram of molecular vibration modes [26].</figcaption></figure>

around the central bond. These vibrational modes provide valuable information about the bonds and structure of molecules, making IR spectroscopy a powerful tool for molecular analysis (Figure 2) [28].

These molecular vibrations are essential for IR spectroscopy, as they manifest themselves as specific absorptions at distinct frequencies (wave numbers) in the IR spectrum [26]. By analyzing these absorption frequencies and their intensities, we can identify the specific functional groups present in a molecule, enabling the precise characterization of chemical compounds [31]. Vibrational stretching frequency follows Hooke's law, where it is proportional to bond strength and inversely proportional to atomic mass. Thus, a triple bond vibrates at a higher frequency than a double or single bond (Table 1) [26].

<table><caption>**Table 1.**
IR bands are used more in catalytic studies [26].</caption><thead><tr><th>Functional group</th><th>Absorption band(s) (cm<sup>-1</sup>)</th><th>Functional group</th><th>Absorption band(s) (cm<sup>-1</sup>)</th></tr></thead><tbody><tr><td>Alcohol O—H stretch</td><td>3550–3200</td><td>Aromatic C—H bending</td><td>860–680</td></tr><tr><td>Water O—H stretch</td><td>3700–3100</td><td>Aromatic C—C bending</td><td>1700–1500</td></tr><tr><td>Carboxylic acid O—H stretch</td><td>3000–2500</td><td>Nitrile C=N stretch</td><td>2260–2220</td></tr><tr><td>Alkyl C—H stretch</td><td>2950–2850</td><td>N—H stretch</td><td>3500–3350</td></tr><tr><td>Alkenyl C—H stretch</td><td>3100–3010</td><td>Aldehyde C=O</td><td>1740–1715</td></tr><tr><td>Alkenyl C—C stretch</td><td>1680–1620</td><td>NOx stretch</td><td>1600</td></tr><tr><td>Alkynyl C—H stretch</td><td>≈3300</td><td>Ester C—O</td><td>1300–1000</td></tr><tr><td>Alkynyl C=C stretch</td><td>2260–2100</td><td>C—O—C stretch</td><td>1250–1050</td></tr><tr><td>Aromatic C—H stretch</td><td>≈3030</td><td></td><td></td></tr></tbody></table>
# 3. Macroalgae

## 3.1 Seaweeds classification

The seaweeds are photosynthetic organisms essential to marine ecosystems, grouped into three main categories: red algae (Rhodophyceae), brown algae (Phaeophyceae), and green algae (Chlorophyceae) [32]. Each category is distinguished by its pigments, cell structure, and specific habitats [33]. Red algae, rich in phycoerythrin, phycocyanin, and chlorophyll, have complex cell walls and thrive in a variety of marine habitats, including those of species such as *Porphyra*, *Chondrus*, and *Eucheuma*. Brown algae, characterized by chlorophyll a and c as well as fucoxanthin, have walls reinforced with alginic acid and sulfated fucans and include species, such as *Laminaria*, *Fucus*, and *Sargasum*, inhabiting mainly temperate and cold zones. Green algae, containing chlorophyll a and b, can be unicellular or multicellular and are found in marine and freshwater habitats, with species, such as *Ulva*, *Caulerpa*, and *Codium*. They play a crucial role in producing oxygen and stabilizing coastal ecosystems, and they are also economically important for their applications in the food, pharmaceutical, and cosmetics industries [34]. Understanding and preserving these algae is essential to maintaining marine biodiversity and supporting industries dependent on these resources.

## 3.2 Marine algae reproduction cycle

Marine algae, or macroalgae, present a remarkable diversity in their reproductive cycles, varying according to their classification into red algae (Rhodophyta), brown algae (Phaeophyceae), or green algae (Chlorophyta), each with distinct characteristics that influence their adaptation to marine environments.

### 3.2.1 Rhodophyta

Red algae are characterized by a complex life cycle dominated by the gametophyte phase [35]. This cycle alternates between a diploid sporophyte phase and a haploid gametophyte phase [36]. Asexual reproduction is common, often occurring through fragmentation where pieces of alga can develop into new individuals. Sexual reproduction involves the release of male and female gametes into the water, followed by fertilization [35].

### 3.2.2 Phaeophyceae

Brown algae have a heteromorphic life cycle, alternating between a diploid sporophyte phase and a haploid gametophyte phase [37, 38]. The sporophyte is generally larger and more complex than the gametophyte [39]. Asexual reproduction occurs through fragmentation or the formation of specialized spores [37]. Sexual reproduction involves the release of gametes into the water, where fertilization occurs.

### 3.2.3 Chlorophyta

Green algae show great variability in their reproductive cycles [40]. Some species alternate between haploid and diploid generations. Asexual reproduction can occur
through fragmentation or the formation of specialized reproductive structures [41].
Sexual reproduction also involves the release of gametes into the water for fertilization.

These reproductive cycles are influenced by environmental factors, such as light, temperature, and nutrient availability [42]. They are crucial to the survival and diversity of marine algae, contributing significantly to marine ecosystems by recycling nutrients and providing essential habitats for many marine species. In-depth study of these cycles provides a better understanding of the adaptation and population dynamics of marine algae, essential for the management and conservation of marine resources and coastal habitats.

### 3.3 Cell wall of seaweeds

Seaweeds have cell walls rich in polysaccharides such as cellulose, alginates, and carrageenans [43], which play crucial roles in both cell structure and the industrial applications of hydrocolloids [44]. Cellulose, predominant in green algae, confers essential rigidity to cell walls, ensuring mechanical resistance against marine environmental stresses [45]. Alginates, found mainly in brown algae, are polysaccharides capable of forming gels in the presence of calcium, thus contributing to cell wall stability and regulating osmotic balance. Carrageenans, found in red algae, are sulfated polysaccharides that add elasticity and flexibility to cell walls, in addition to possessing gelling properties used in various industries [25, 46]. These cell wall polysaccharides are directly linked to the functional properties of hydrocolloids extracted from seaweed. For example, agar, carrageenan, and alginate are widely used for their gelling, thickening, and stabilizing capacities in food, cosmetics, pharmaceuticals, and biotechnological products. This close relationship between cell walls and hydrocolloids demonstrates not only the importance of marine resources for sustainable industrial applications but also the efficient exploitation of algae's natural properties. The complex interactions between polysaccharide components of cell walls and hydrocolloid applications highlight the potential of marine algae as a source of biocompatible and ecologically sustainable materials in various technological and industrial fields.

## 4. Description of the different hydrocolloid types and extraction processes

### 4.1 Hydrocolloids in Chlorophyceae

Hydrocolloids are hydrophilic polymers widely used in the food and pharmaceutical industries for their thickening, gelling, and stabilizing properties [47]. Among the various sources of these polymers, algae of the Chlorophyceae class, or green algae, are of particular interest due to their unique chemical composition and abundance in aquatic environments [47].

#### 4.1.1 Hydrocolloid types in Chlorophyceae

Chlorophyceae produce several types of hydrocolloids, the most notable of which are polysaccharides such as ulvan and rhamnan. The ulvan backbone is made up of repeating disaccharide units: ulvanobioronic acid (types A and B) and ulvanobiose (type U). Type A contains β-D-glucuronic acid linked (1,4) to α-L-rhamnose
3-sulfate, while type B replaces β-D-glucuronic acid with α-L-iduronic acid [48]. Type U has β-D-xylose (1,4) linked to α-L-rhamnose 3-sulfate. These polysaccharides are extracted mainly from the *Ulva* and *Enteromorpha* genera.

### 4.1.2 Properties and applications

Hydrocolloids derived from Chlorophyceae have a wide range of functional properties, making them useful in many applications:

* **Thickeners and gelling agents:** Green algae polysaccharides can form gels and viscous solutions, which is crucial for the food industry in the manufacture of products such as sauces, jams, and gelled desserts.

* **Stabilizers:** By stabilizing emulsions and suspensions, these hydrocolloids improve the texture and consistency of food and cosmetic products.

* **Film formers:** Some Chlorophyceae polysaccharides can form films, used in biodegradable food packaging and edible films (Figure 3).

### 4.1.3 Extraction and production

Hydrocolloid extraction from Chlorophyceae generally involves physical and chemical methods to isolate the polysaccharides of interest. Algae are harvested, washed, dried, and ground before being subjected to hot-water extraction processes, followed by ethanol precipitation or other methods to purify the polysaccharides.

### 4.1.4 Research and development

Chlorophyceae hydrocolloid research focuses on optimizing extraction techniques, characterizing chemical structures, and assessing functional properties for

<figure><img src="image_4.png" /><figcaption>**Figure 3.**
Chemical structure of a typical ulvane [48].</figcaption></figure>
specific applications. The exploration of new species of green algae and the chemical modification of polysaccharides are also active areas of research. In addition, hydrocolloids from the Chlorophyceae represent a valuable natural resource with a wide range of industrial applications. Their ability to improve the textural and functional properties of food and non-food products makes them the ingredients of choice for industries seeking to exploit sustainable, environment-friendly materials.

## 4.2 Hydrocolloids in Phaeophyceae

Hydrocolloids from the Phaeophyceae, or brown algae, are natural polymers that play a crucial role in a variety of industrial applications, notably in the food, pharmaceutical, and cosmetics sectors. Brown algae are particularly rich in unique polysaccharides, such as alginate, fucoidane, and laminarin, which possess remarkable functional properties.

### 4.2.1 Hydrocolloid types in Phaeophyceae

*   **Alginate:** The best-known and most widely used hydrocolloid of brown algae. It is mainly extracted from the Laminaria, Macrocystis, and Ascophyllum genera. Alginate is composed of mannuronate and guluronate blocks, giving it a unique ability to form solid gels and films in the presence of divalent cations such as calcium.

*   **Fucoidan:** This sulfated polysaccharide is found mainly in the cell walls of brown algae. Fucoidan has interesting biological properties, including anticoagulant, antiviral, and anti-inflammatory activities, making it promising for medical applications.

*   **Laminarin:** Another polysaccharide found in brown algae, laminarin is a glucan that serves as an energy reserve for these algae. It is being studied for its immunomodulatory and anti-tumor properties (Figure 4).

<figure><img src="image_5.png" /><figcaption>**Figure 4.**
*Chemical structure of the Fucoidan from (A) Fucus vesiculosus and Ascophyllum nodosum and (B) Laminaria saccharina; Alginate base structure (C) and laminarin base structure (D).*</figcaption></figure>
### 4.2.2 Properties and applications

Hydrocolloids from Phaeophyceae have many applications due to their unique properties:

* *Thickeners and gelling agents:* Alginates are used to thicken and gel a wide variety of food products, including ice creams, sauces, and baked goods.

* *Stabilizers:* By stabilizing emulsions and suspensions, these hydrocolloids improve the texture and consistency of food and cosmetic products.

* *Bioactives:* Fucoidanes are used for their biological properties, notably in dietary supplements and pharmaceutical formulations.

* *Coatings and films:* Alginates can form protective films and coatings used in food packaging and biomedical applications.

### 4.2.3 Extraction and production

Hydrocolloid extraction from Phaeophyceae generally involves both physical and chemical processes. Algae are harvested, washed, crushed, and subjected to chemical treatments to isolate the polysaccharides of interest. Alginates, for example, are often extracted using alkaline solutions followed by acid precipitation.

### 4.2.4 Research and development

Current research focuses on optimizing extraction processes, chemically modifying hydrocolloids to enhance their functional properties, and exploring new applications. Detailed characterization of the structures and properties of brown algal hydrocolloids is also an active area of research. Phaeophyceae hydrocolloids represent a valuable resource with a wide range of industrial applications. Their ability to improve the textural and functional properties of food products, as well as their unique biological properties, make them ingredients of choice for a variety of industries seeking to exploit sustainable, environment-friendly materials.

## 4.3 Hydrocolloids in Rhodophyceae

Hydrocolloids from the Rhodophyceae, or red algae, are natural polysaccharides of considerable importance in the food, pharmaceutical, and cosmetics industries. Red algae are particularly well known for producing two main types of hydrocolloids: agar and carrageenan, each with distinct properties and varied applications.

### 4.3.1 Hydrocolloid types in Rhodophyceae

#### 4.3.1.1 Agar

Agarans, polysaccharides found in algae, differ from carrageenans in their more diverse functional groups. They generally come in two fractions (agarose and agaropectin), which differ in their ionic charges, with the neutral agar fraction (known as agarose and agaropectin) serving as the charged fraction [49].
*   **Composition and extraction:** Agar is mainly extracted from the *Gelidium* and *Gracilaria* genera. It is composed of two main fractions: agarose, which forms solid gels, and agaropectin, which contains sulfated groups and other substituents.

*   **Properties:** Agar forms heat-reversible gels at low concentrations and is used for its gelling, stabilizing, and thickening properties.

*   **Applications:** It is widely used in the food industry (as a gelling agent for jams, jellies, and dairy products), in microbiology laboratories (as a culture medium), and in the pharmaceutical industry (as an excipient and controlled-release agent) (Figure 5).

#### 4.3.1.2 Carrageenan

Carrageenan, extracted from red macroalgae such as *Chondrus crispus*, is composed of galactose and 3,6-anhydrogalactose linked by glycosidic bonds (Figure 6). It is divided into kappa, iota, lambda, mu, and nu types, each with specific properties for a variety of industrial applications [49].

*   **Composition and extraction:** Carrageenan is extracted mainly from the *Chondrus*, *Eucheuma*, and *Kappaphycus*. It exists in three main forms: kappa, iota, and lambda, each with distinct gelling and thickening properties.

*   **Properties:** Kappa and iota carrageenans form gels in the presence of specific cations (such as potassium and calcium), while lambda carrageenan acts primarily as a thickener.

<figure><img src="image_6.png" /><figcaption>**Figure 5.**

*Chemical structure of agaran from Gymnogongrus tenuis [48, 49].</figcaption></figure>

<figure><img src="image_7.png" /><figcaption>**Figure 6.**

*Chemical structure of μ-, κ-, ν-, ι-, and λ-carrageenan with the transformation of carrageenan precursors into κ- and ι-carrageenan under alkaline conditions [49].</figcaption></figure>
* *Applications*: Carrageenans are used as gelling agents, thickeners, and stabilizers in a variety of food products, including dairy products, processed meats, and desserts. They are also used in cosmetic and pharmaceutical formulations for their textural and stabilizing properties.

### 4.3.2 Properties and applications

The hydrocolloids of the Rhodophyceae possess functional properties that make them valuable for a variety of industrial applications.

* *Thickeners and gelling agents*: Agaroses and carrageenans are used for their ability to form solid gels and viscous solutions.

* *Stabilizers*: They improve the texture and stability of emulsions and suspensions in food and cosmetic products.

* *Texturizers*: These hydrocolloids modify the texture of food products to improve mouthfeel and palatability.

### 4.3.3 Extraction and production

The extraction of hydrocolloids from Rhodophyceae generally involves both mechanical and chemical processes. The algae are harvested, cleaned, and crushed before being subjected to hot water treatments to extract the polysaccharides. The solutions are then filtered and precipitated to isolate agar or carrageenan.

### 4.3.4 Research and development

Current research focuses on improving extraction techniques, chemically modifying hydrocolloids to enhance their functionality, and exploring new applications in the biomedical and environmental industries. Characterizing the physico-chemical properties of agaroses and carrageenans is also an active area of research. Rhodophyceae hydrocolloids are valuable ingredients thanks to their unique gelling, thickening, and stabilizing properties. Their use in a variety of industrial sectors bears witness to their versatility and economic importance. Ongoing innovation in the extraction and application of these natural polysaccharides promises to further expand their uses and improve their performance in a variety of contexts.

## 5. FTIR characterization of hydrocolloids

Fourier transform infrared spectroscopy (FTIR) is a leading analytical method for characterizing hydrocolloids, particularly those derived from marine sources such as algae. This technique enables in-depth exploration of the molecular structure and functional properties of the polysaccharides that make up these materials, offering a wide range of applications in various industrial sectors.

### 5.1 FTIR for hydrocolloid analysis: functional groups and interactions

FTIR is an essential tool for the analysis of hydrocolloids, which are substances capable of forming gels or viscous solutions when in contact with water. Using FTIR,
we can identify the functional groups present in hydrocolloids, such as hydroxyl (O-H) and carbonyl (C=O) groups, which play a crucial role in their gelling and solubility properties [50]. This method also enables us to examine interactions between hydrocolloids and other components, such as texturizing agents, by observing hydrogen bonds and associated structural changes. FTIR is particularly useful for analyzing changes in hydrocolloid structure during gel formation or in response to thermal or chemical treatments. Although direct quantification of hydrocolloid concentrations can present challenges, spectral-based calibration methods can provide rough estimates. By combining FTIR with other techniques such as rheology or microscopy, a more complete understanding of the properties and behavior of hydrocolloids in various formulations can be obtained.

## 5.2 Application of FTIR spectroscopy in the analysis of marine hydrocolloids

FTIR spectroscopy is essential for analyzing marine hydrocolloids, providing critical information about their composition and properties. It detects and identifies functional groups, revealing the molecular structure of these compounds, including the types of glycosidic linkages in polysaccharides, which influence their physical and functional properties [47]. This technique is also vital for quality control and authentication, as it compares spectral signatures with reference samples to ensure purity and conformity. Furthermore, FTIR allows for the study of complex interactions between hydrocolloids and other substances, such as salts and proteins, which is crucial for optimizing formulations and understanding their properties. It also enables quantitative analysis by measuring peak intensities, facilitating accurate determination of component concentrations in hydrocolloid mixtures. Marine hydrocolloids are valued for their durability, biocompatibility, and unique functional properties, making FTIR analysis an indispensable tool in their exploration for various applications in the food, pharmaceutical, cosmetics, and biotechnology sectors [47]. In short, FTIR spectroscopy plays a vital role in characterizing marine hydrocolloids and fully exploiting their potential in numerous industrial and technological fields, offering precise detection and analysis of their functional groups.

## 5.3 Computer optimization of FTIR analysis of marine hydrocolloids

The integration of IT tools into Fourier transform infrared (FTIR) spectroscopy has revolutionized the management and analysis of spectral data, particularly for marine hydrocolloids such as polysaccharides from algae. Software like FTIR Essential, OMNIC, and OriginPro facilitates instrument control, automating data acquisition and allowing precise adjustments of parameters such as resolution and wavelength range. Pre-processing algorithms enhance data quality by correcting baseline deviations, smoothing spectra, and normalizing data. During interpretation, specialized software compares acquired spectra with reference libraries to identify functional groups and employs advanced chemometric techniques, such as Principal Component Analysis (PCA), to extract significant patterns and classify samples [51, 52]. Tools like Unscrambler X 10.4, XLSTAT, Excel, and Minitab are commonly used for efficient analysis. Quantitative analysis, based on peak intensity measurements, helps determine the concentration of different compounds within samples. Integrated databases facilitate the rapid verification of sample authenticity, while automated workflows ensure swift and efficient analysis, catering to industrial and scientific needs. These IT tools also produce detailed reports, guaranteeing the
traceability and reproducibility of results. This makes FTIR analyses more precise and efficient, enhancing the application of marine hydrocolloids in sectors, such as pharmaceuticals, food, and cosmetics.

## 5.4 Infrared spectrum analysis of hydrocolloids: methodology and applications

Analyzing the infrared (IR) spectra of hydrocolloids is an essential systematic method for revealing their chemical composition and functional properties, particularly useful in the food, pharmaceutical, cosmetics, and biomedical industries. The process begins with the acquisition of spectral data using FTIR spectroscopies, tuned by specialized software to define spectral range and resolution, ensuring accurate and reproducible measurements. The raw spectra are then pre-processed by baseline correction, smoothing, and normalization, which improves the accuracy of results by eliminating background noise and facilitating comparison between samples. IR spectra can be used to determine the functional groups present in hydrocolloids, such as hydroxyl, carbonyl, and sulfate bonds. Each functional group absorbs infrared energy at specific frequencies, producing distinct peaks in the spectrum. For example, O-H stretches occur between 3200 and 3600 cm⁻¹ [53–55], C=O stretches between 1700 and 1750 cm⁻¹ [50], and C-O stretches between 1000 and 1300 cm⁻¹ [50]. The positions of these peaks may vary slightly depending on the chemical structure and environment of the hydrocolloids. Then, spectral analysis software facilitates interpretation by comparing the spectra obtained with reference libraries, helping to determine the chemical composition and conformity of hydrocolloids. Quantitative analysis of IR spectra enables the relative quantities of components to be estimated, while chemometric methods such as PCA reveal important information for sample classification and process optimization. IR combined with PCA distinguishes food polysaccharides, identifying specific spectral bands for components such as starch, β-glucan, galactan, and carrageenans [50]. These techniques highlight the differences between families of purified polysaccharides, crucial for guaranteeing product quality and safety in various industrial sectors.

# 6. A few illustrations of IR spectra and their interpretation for hydrocolloids

## 6.1 Preparing samples for FTIR analysis

FTIR spectra of polysaccharides extracted from various algae species, including *Caulerpa taxifolia*, *Uva fasciata*, *Sargassum latifolium*, *Laminaria digitata*, *Sargassum aquifolium*, *Cystoseira crinita*, *Gelidium pusillum*, *Gracilaria corticata*, *Kappaphycus striatus*, and *Eucheuma denticulatum*, were recorded using a FTIR spectrometer (model 6300, JASCO, Japan). The functional groups were analyzed in the range of 4000–400 cm⁻¹ with a resolution of 4 cm⁻¹ using KBr pellets. Dried powder samples were evaluated and compared to previous research data. Data processing was carried out using OMNIC 9.2.86 software. To ensure accurate results, a background measurement was performed before each sample analysis.

## 6.2 FTIR spectra of ulvan from green algae

In the FTIR analysis of *Caulerpa taxifolia*, a green alga, referring to the study of Bayro et al. [56]. The FTIR spectrum of *C. taxifolia* (Figure 7A) shows strong
absorption at 3340 cm⁻¹, compared with 3433 cm⁻¹ for ulvan, mainly due to stretching of the -OH groups and, to a lesser extent, the N-H group. Peaks at 2923 cm⁻¹ for *C. taxifolia* and 2939 cm⁻¹ for ulvan correspond to stretching of the C-H groups. The peak at 1645 cm⁻¹ indicates a conjugated aromatic C=O group. Bands at 1235 cm⁻¹ in *C. taxifolia* and between 1000 and 1200 cm⁻¹ in ulvan are associated with C-O vibrations of sugar rings and glycosidic bonds. Peaks at 1149, 1077 and 1015 cm⁻¹ for *C. taxifolia*, and at 1164, 1085' and 1034 cm⁻¹ for ulvan, confirm a pyranose configuration, indicating that *C. taxifolia* is a polysaccharide similar to ulvan.

The study by Barakat et al. [57], using *Uva fasciata* as the alga of interest (Figure 7), reveals that the FTIR spectra of ulvan show a band at 3375–3380 cm⁻¹ for O-H stretching, a weak peak at 2925–2928 cm⁻¹ for aliphatic C-H stretching, and a band at 1076 cm⁻¹ for C-O stretching of the sugar rhamnose. The presence of sulfate groups is confirmed by a band at 820–856 cm⁻¹ and a shoulder at 1225–1226 cm⁻¹, while the band at 748–750 cm⁻¹ corresponds to sugar rings, showing a diversity of sulfate groups in the ulvan of *Uva fasciata*.

## 6.3 FTIR spectra of alginate from brown algae

The FTIR profile of sodium alginate extracted from *Sargassum latifolium* (Figure 8A) reveals several important functional groups [58]. A broad absorption band at 3465 cm⁻¹ is attributed to the stretching of the hydroxyl group (-OH). Peaks in the 4000–3400 cm⁻¹ range correspond to alcohol and acid groups. The bands between 1628 and 1428 cm⁻¹ are associated with the vibrations of carboxylate anions (COO⁻), providing key insights into the structure of alginates and their interactions

<figure><img src="image_8.png" /><figcaption>**Figure 7.** FTIR spectrum of the extracted ulvan: from (A) Caulerpa taxifolia and (B) Uva fasciata.</figcaption></figure>

<figure><img src="image_9.png" /><figcaption>**Figure 8.** FTIR spectrum of sodium alginate from (A) Sargassum latifolium and (B) Laminaria digitata.</figcaption></figure>
with metals. The band at 2923–2854 cm⁻¹ is linked to aliphatic -CH stretching and C-H vibrations. Peaks in the 1090–1030 cm⁻¹ range reflect the saccharide structure, while the band at 1734 cm⁻¹ indicates the stretching of the C=O group. Bands at 848 cm⁻¹ and around 600 cm⁻¹ confirm the presence of sulfate groups and O=S=O deformations, respectively [58].

Similarly, sodium alginate extract from *Laminaria digitata* (**Figure 8B**) shows weak absorption bands at 1316.79, 1125.53, and 1094.66 cm⁻¹ [59]. These bands are respectively related to C-C-H and O-C-H deformations, C-O stretching, and C-O and C-C vibrations in the pyranose rings. In addition, the band at 1035.6 cm⁻¹ is also attributed to C-O stretching vibrations. The anomeric region (950–750 cm⁻¹) is the most debated in carbohydrates [60]. The spectrum shows a band at 948.2 cm⁻¹, which has been attributed to the C-O stretching vibration of uronic acid residues, and a band at 902.83 cm⁻¹ attributed to the C1-H deformation vibration of β-mannuronic acid residues. The band at 818.76 cm⁻¹ appears to be characteristic of mannuronic acid residues [59]. FTIR analyses of sodium alginates from different algae show specific characteristics that confirm the presence of distinct functional groups and their complex chemical structure.

## 6.4 FTIR spectra of Fucoidan from brown algae

FTIR spectra of fucoidans extracted from *Sargassum aquifolium* (**Figure 9A**) [61] and *Cystoseira crinita* (**Figure 9B**) [62] show distinct characteristics. For *Sargassum aquifolium*, the peak at 3425 cm⁻¹ corresponds to the O-H stretching of the polysaccharide, while that at 2960 cm⁻¹ indicates the C-H groups of the carbohydrate. Peaks at 1641 cm⁻¹ and 1420 cm⁻¹ are associated with C=O vibrations and those at 1260 cm⁻¹ and 1056 cm⁻¹ with S=O vibrations of sulfated polysaccharides. The peak at 802 cm⁻¹ confirms the presence of sulfate groups (C-O-S) [63].

For *Cystoseira crinita*, the band at 3427 cm⁻¹ is linked to O-H stretching of sugar residues [64]. The signal at 1611 cm⁻¹ is attributed to vibrations of ester groups (C=O) in acidic residues, indicating uronic acids [65]. The peak at 1412 cm⁻¹ could correspond to the stretching of -CH₂ groups in neutral monosaccharides and -CH₃ groups in fucosyl residues [58]. The band at 1135 cm⁻¹ is attributed to the stretching of pseudosymmetric sulfate groups (O=S=O) and hemiacetal groups [66, 67]. These results, in line with the literature, demonstrate the importance of FTIR spectra for identifying the functional groups of sulfated polysaccharides.

<figure><img src="image_10.png" /><figcaption>**Figure 9.** FTIR spectrum of Fucoidan from (A) Sargassum aquifolium and (B) *Cystoseira crinita*.</figcaption></figure>
## 6.5 FTIR spectra of agar from red algae

FTIR spectra of agar extracted from the algae *Gelidium pusillum* and *Gracilaria corticata* reveal several distinctive features essential for understanding the chemical composition of agar [68]. Analyses show peaks corresponding to various functional groups typical of agar, such as alcohols, amines, and alkanes [69]. The peaks observed at 3419.3 cm⁻¹ (Figure 10A) and 3421.9 cm⁻¹ (Figure 10B) are attributed to the stretching of O-H bonds, signaling the presence of hydroxyl groups, a fundamental characteristic of the polysaccharides present in agar [68]. The absorption band at 2900 cm⁻¹ is associated with the methoxyl group, another important component of agar's chemical structure. In addition, the band at 1600 cm⁻¹ is linked to CO and NH groups, which play a crucial role in conjugated peptide bonds, contributing to agar's structure and gelling properties. The bands were detected at 930 cm⁻¹, 1073.3 cm⁻¹ for *Gelidium pusillum* and 1072.3 cm⁻¹ for *Gracilaria corticata* are characteristic of 3,6-anhydrogalactose bridges, confirming the presence of this specific structure in extracted agar. These observations corroborate earlier findings by Pereira et al. [70], who also identified these features in agar samples, affirming the accuracy of the data obtained in the present study. The features identified by FTIR provide key information on the composition and structure of agar, essential for its scientific and industrial applications.

## 6.6 FTIR spectra of carrageenan from red algae

The FTIR spectral profile of polysaccharide extracted from the algae *Kappaphycus striatus* and *Eucheuma denticulatum* revealed absorption bands characteristic of

<figure><img src="image_11.png" /><figcaption>**Figure 10.** FTIR spectrum of agar from (A) *Gelidium pusillum* and (B) *Gracilaria corticata*.</figcaption></figure>

<figure><img src="image_12.png" /><figcaption>**Figure 11.** FTIR spectrum of red algal carrageenan from (A) *Kappaphycus striatus* and (B) *Eucheuma denticulatum*.</figcaption></figure>
<table><caption>**Table 2.**
Summary of FTIR spectra identified with the species studied.</caption><thead><tr><th>Absorbance bands (cm<sup>-1</sup>)</th><th>Vibrational molecule</th><th>Caulerpa taxifolia</th><th>Uva fasciata</th><th>Sargassum latifolium</th><th>Laminaria digitata</th><th>Sargassum aquifolium</th><th>Cystoseira crinita</th><th>Gelidium pusillum</th><th>Gracilaria corticata</th><th>Kappaphycus striatus</th><th>Eucheuma denticulatum</th></tr></thead><tbody><tr><td>3500–3000</td><td>Dimer OH [24]</td><td>3341</td><td>3400</td><td>3445</td><td>3421</td><td>3385</td><td>3435</td><td>3419</td><td>3442</td><td>3408</td><td>3332.87</td></tr><tr><td>3000–2800</td><td>C–H stretching, symmetric, asymmetric [24]</td><td>2923</td><td>2926</td><td>2899, 2835</td><td>2942</td><td>2898</td><td>+</td><td>2917, 2860</td><td>2929, 2855</td><td>2890</td><td>2889.81</td></tr><tr><td>1655–1645</td><td>C=O stretch (H-bond) [78]</td><td>1645</td><td>1645</td><td>1655</td><td>1645</td><td>1658</td><td>+</td><td>1655</td><td>1645</td><td>+</td><td>+</td></tr><tr><td>1500–1400</td><td>Ar C–C stretch [78]</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>—</td><td>—</td><td>+</td><td>+</td></tr><tr><td>1450–1350</td><td>S=O sulfate ester [24]</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>1420</td><td>1426</td><td>+</td><td>+</td></tr><tr><td>1450–1410</td><td>COO<sup>-</sup> symmetric [78]</td><td>+</td><td>1445</td><td>1448</td><td>1417</td><td>1485</td><td>1412</td><td>—</td><td>—</td><td>—</td><td>—</td></tr><tr><td>1300–1150</td><td>C–O stretch</td><td>1373</td><td>+</td><td>1170</td><td>1126</td><td>1275, 1252</td><td>+</td><td>1262, 1203</td><td>1262</td><td>1274, 1245</td><td>1156.601221.20</td></tr><tr><td>1080–1020</td><td>C=O-C Glycosidique [24]</td><td>1077</td><td>+</td><td>1080</td><td>1035</td><td>1020</td><td>1152</td><td>—</td><td>—</td><td>—</td><td>—</td></tr><tr><td>1070, 933–928 (shoulder)</td><td>3,6-anhydro-D-galactos</td><td>—</td><td>—</td><td>—</td><td>—</td><td>—</td><td>—</td><td>—</td><td>—</td><td>933</td><td>928</td></tr><tr><td>885–870</td><td>C–H out of plane</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td></tr><tr><td>850–800</td><td>Sulfate (S=O) [24]</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>+</td><td>845</td><td>839.37</td></tr><tr><td>850–840</td><td>D-galactose-4-sulphate [24]</td><td>—</td><td>—</td><td>—</td><td>—</td><td>845</td><td>845</td><td>+</td><td>+</td><td>+</td><td>+</td></tr><tr><td>950–930</td><td>Pyranoside</td><td>—</td><td>—</td><td>—</td><td>—</td><td>—</td><td>—</td><td>930</td><td>931</td><td>—</td><td>—</td></tr><tr><td>905, 805–800 (shoulder)</td><td>3,6-anhydro-D-galactose-2-sulphate [24]</td><td>—</td><td>—</td><td>—</td><td>—</td><td>—</td><td>—</td><td>+</td><td>+</td><td>+</td><td>+</td></tr><tr><td>900–800</td><td>C–H Mannuronic/ Guluronic</td><td>—</td><td>—</td><td>825</td><td>856</td><td>—</td><td>—</td><td>+</td><td>+</td><td>—</td><td>—</td></tr></tbody><tfoot><tr><td colspan="12">(–) Absence of peak; (+) Presence of peak (low intensity).</td></tr></tfoot></table>
sulfated carrageenan (Figure 11A and B). For *Kappaphycus striatus*, the bands at 930, 850, and 1260 cm⁻¹ indicate the presence of symmetrical C-O-C vibrations of 3,6-anhydro-D-galactose, the C4-O-S stretch and the O=S=O group of β-D-galactose-4-sulfate, respectively, confirming kappa carrageenan composition [71, 72]. The band at 1640 cm⁻¹ is related to the structural deformation of water [73]. In addition, bands at 1159, 1070, 2934, and 3411 cm⁻¹ are attributed to -O bridge stretching, C-O, C-H, and O-H stretching vibrations, respectively [74, 75]. These results confirm the presence of kappa carrageenan in *Kappaphycus striatus*.

FTIR analysis of polysaccharide extracted from *Eucheuma denticulatum* revealed bands typical of carrageenans, according to Vandanjon et al. [76]. The sulfated ester at 1221.5 cm⁻¹ (S=O) is common to ¹-, κ- and ν-carrageenans [24, 77]. The C-O of 3,6-anhydrogalactose occurs at 1061.82 cm⁻¹ for ¹- and κ-carrageenans, while the band at 973.16 cm⁻¹ indicates the presence of galactose and these carrageenans. The band at 925.06 cm⁻¹ is specific to ¹-carrageenan, and at 905 cm⁻¹ the C-O-SO₄ group on the C2 of 3,6-anhydrogalactose is characteristic of ¹- and κ-carrageenans. The bands at 843 cm⁻¹ and 805 cm⁻¹ are associated with carrageenans and ¹-carrageenan, respectively [76]. The results therefore indicate that the nearest peaks in this spectrum correspond mainly to iota-carrageenan.

## 6.7 Overview of FTIR spectra of analyzed hydrocolloids

The FTIR spectra analyzed reveal distinctive features that allow for the identification and differentiation of hydrocolloids extracted from algae. The primary absorption bands observed indicate the presence of specific functional groups, showcasing the structural complexity of these polysaccharides. Analysis shows that the spectra obtained are more or less the same as those obtained in previous studies, with the absorbance bands slightly shifted. The studies listed above are summarized in Table 2.

For polysaccharides extracted from green algae, absorption bands around 3340–3433 cm⁻¹ are primarily attributed to -OH stretching, while peaks at 2923–2939 cm⁻¹ are associated with C-H stretching. The C-O vibrations in sugar rings and glycosidic bonds appear between 1000 and 1200 cm⁻¹, confirming the presence of pyranose configurations. These characteristics are crucial for understanding the structure and functional properties of ulvan. The spectra of alginates and fucoidans from brown algae display typical bands for carboxylate (COO⁻) and sulfate (S=O) groups. Bands around 1628–1428 cm⁻¹ are particularly informative for identifying alginates, indicating their ability to bind metal ions, while peaks between 820 and 856 cm⁻¹ confirm the presence of sulfate groups, essential for fucoidans. Finally, the spectra of agar and carrageenan from red algae reveal characteristic bands for 3,6-anhydrogalactose bridges, which are fundamental to their gelling properties. Peaks associated with O-H, C-H, and SO₄²⁻ groups are crucial for identifying the structure of carrageenans, enabling the distinction between kappa and iota forms. These FTIR analyses thus provide a deep understanding of the molecular structures and functional potentials of hydrocolloids extracted from different algae species.

## Acknowledgements

This chapter was written with contributions from the URGPGE team and in collaboration with a member of LCO-SM. The authors would like to thank Prof.
Ravelonandro Pierre for initiating such a project, and for allowing all collaborators to co-contribute to this work. We would also like to thank the University of Antananarivo for their support of this project from start to finish.

**Conflict of interest**

The author declares no conflict of interest.

**Author details**

Elando Fréda Zamanileha¹, Julie Tantely Mitantsoa¹, Picardino Frienduc Vaonalamihanta², Andriamanarivosoa Rija Razafintsalama¹, Fara Arimalala Andrianony¹ and Pierre Hervé Ravelonandro¹*

1 Université d'Antananarivo, Faculty of Sciences, Unité de Recherche en Génie de Procédés et Génie de l'Environnement (URGPGE), Antananarivo, Madagascar

2 Université d'Antananarivo, Faculty of Sciences, Laboratory of Organic Chemistry - Marine Substances (LCO-SM), Antananarivo, Madagascar

*Address all correspondence to: pierre.ravelonandro@urgpge-ua.mg; phravelona@yahoo.com

**IntechOpen**

© 2024 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/4.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. [CC BY](https://creativecommons.org/licenses/by/4.0)
**References**

[1] Hallinan DT Jr. Attenuated total reflectance mode for transport through membranes. In: El-Azazy M, Al-Saad K, Ahmed S. El-Shafie, editors. Infrared Spectroscopy-Perspectives and Applications. London, UK: IntechOpen; 2022. DOI: 10.5772/intechopen.107869

[2] Errico S, Moggio M, Diano N, Portaccio M, Lepore M. Different experimental approaches for Fourier-transform infrared spectroscopy applications in biology and biotechnology: A selected choice of representative results. Biotechnology and Applied Biochemistry. 2023;70:937-961. DOI: 10.1002/bab.2411

[3] Agatonovic-Kustrin S, Ramenskaya G, Kustrin E, Ortakand DB, Morton DW. A new integrated HPTLC - ATR/FTIR approach in marine algae bioprofiling. Journal of Pharmaceutical and Biomedical Analysis. 2020;189:113488. DOI: 10.1016/j.jpba.2020.113488

[4] Bertolotti M, Sibilia C, Guzman A. Evanescent waves. In: Bertolotti M, Sibilia CM, Guzman A, editors. Evanescent Waves Opt. Introd. Plasmon. Cham, Switzerland: Springer International Publishing; 2017. pp. 35-67. DOI: 10.1007/978-3-319-61261-4_2

[5] Markovic NMPNR, Ross PN. New Electrocatalysts for fuel cells from model surfaces to commercial catalysts. CATTECH. 2000;4:110-126. DOI: 10.1023/A:1011963731898

[6] Marković NM, Ross PN. Surface science studies of model fuel cell electrocatalysts. Surface Science Reports. 2002;45:117-229. DOI: 10.1016/S0167-5729(01)00022-X

[7] Gambaro LA, Fierro JLG, Tejuca LG, Agudo AL. Surface structure and NiO

dispersion for NiO/γ-Al2O3 catalysts. Surface and Interface Analysis. 1982;4:234-239. DOI: 10.1002/sia.740040604

[8] Yakerson VI, Lafer LI, Danyushevskii VYA, Rubinshtein AM. Infrared spectra of catalysts and adsorbed molecules. Russian Chemical Bulletin. 1967;16:2154-2159. DOI: 10.1007/BF00913298

[9] Tsyganenko AA, Pozdnyakov DV, Filimonov VN. Infrared study of surface species arising from ammonia adsorption on oxide surfaces. Journal of Molecular Structure. 1975;29:299-318. DOI: 10.1016/0022-2860(75)85038-1

[10] Aika K-I, Ozaki A. Kinetics and isotope effect of ammonia synthesis over a singly-promoted iron catalyst. Journal of Catalysis. 1970;19:350-352. DOI: 10.1016/0021-9517(70)90257-5

[11] Faulkner DJ. Interesting aspects of marine natural products chemistry. Tetrahedron. 1977;33:1421-1443. DOI: 10.1016/0040-4020(77)88001-0

[12] Wijesinghe WAJP, Jeon Y-J. Enzyme-assistant extraction (EAE) of bioactive components: A useful approach for recovery of industrially important metabolites from seaweeds: A review. Fitoterapia. 2012;83:6-12. DOI: 10.1016/j.fitote.2011.10.016

[13] Sadeghi A, Rajabiyan A, Nabizade N, Meygoli Nezhad N, Zarei-Ahmady A. Seaweed-derived phenolic compounds as diverse bioactive molecules: A review on identification, application, extraction and purification strategies. International Journal of Biological Macromolecules. 2024;266:131147. DOI: 10.1016/j.ijbiomac.2024.131147
[14] Bourgougnon N, Stiger-Pouvreau V. Chemodiversity and Bioactivity within Red and Brown Macroalgae along the French Coasts, Metropole and Overseas Departements and Territories. Hoboken, USA: John Wiley and Sons, Ltd.; 2011. pp. 58-105

[15] Fernando IPS, Sanjeewa KKA, Samarakoon KW, Lee WW, Kim H-S, Kim E-A, et al. FTIR characterization and antioxidant activity of water soluble crude polysaccharides of Sri Lankan marine algae. Algae. 2017;32:75-86. DOI: 10.4490/algae.2017.32.12.1

[16] Hayashi L, Reis RP, dos Santos AA, Castelar B, Robledo D, de Vega GB, et al. The cultivation of *Kappaphycus* and *Eucheuma* in tropical and subtropical waters. In: Hurtado AQ, Critchley AT, Neish IC, editors. Trop. Seaweed Farming Trends Probl. Oppor. Focus IKappaphycusi IEucheumai Commer. Cham, Switzerland: Springer International Publishing; 2017. pp. 55-90. DOI: 10.1007/978-3-319-63498-2_4

[17] Ji Y, Gao K. Chapter two - effects of climate change factors on marine macroalgae: A review. In: Sheppard C, editor. Adv. Mar. Biol., vol. 88. Academic Press; 2021. pp. 91-136. DOI: 10.1016/bs.amb.2020.11.001

[18] Kwan V, Fong J, Ng CSL, Huang D. Temporal and spatial dynamics of tropical macroalgal contributions to blue carbon. Sci Total Environ. 2022;828:154369. DOI: 10.1016/j.scitotenv.2022.154369

[19] Cinner JE, Caldwell IR, Thiault L, Ben J, Blanchard JL, Coll M, et al. Potential impacts of climate change on agriculture and fisheries production in 72 tropical coastal communities. Nature Communications. 2022;13:3530. DOI: 10.1038/s41467-022-30991-4

[20] Martínez-Vázquez RM, Milán-García J, De Pablo Valenciano J. Challenges of the blue economy: Evidence and research trends. Environmental Sciences Europe. 2021;33:61. DOI: 10.1186/s12302-021-00502-1

[21] Khan SA, Khan SB, Khan LU, Farooq A, Akhtar K, Asiri AM. Fourier transform infrared spectroscopy: Fundamentals and application in functional groups and Nanomaterials characterization. In: Sharma SK, editor. Handb. Mater. Charact. Cham, Switzerland: Springer International Publishing; 2018. pp. 317-344. DOI: 10.1007/978-3-319-92955-2_9

[22] Meskinfam M. 17 - polymer scaffolds for bone regeneration. In: Tanzi MC, Farè S, editors. Charact. Polym. Biomater. Woodhead Publishing; 2017. pp. 441-475. DOI: 10.1016/B978-0-08-100737-2.00017-0

[23] Herrero YR, Camas KL, Ullah A. Chapter 4 - characterization of biobased materials. In: Ahmed S, Annu, editors. Adv. Appl. Biobased Mater. Amsterdam, Netherlands: Elsevier; 2023. pp. 111-143. DOI: 10.1016/B978-0-323-91677-6.00005-2

[24] Knutsen SH, Myslabodski DE, Larsen B, Usov AI. A modified system of nomenclature for red algal Galactans. Botanica Marina. 1994;37:163-169. DOI: 10.1515/botm.1994.37.2.163

[25] Aguilan JT, Broom JE, Hemmingson JA, Dayrit FM, Montaño MNE, Dancel MCA, et al. Structural analysis of carrageenan from farmed varieties of Philippine seaweed. Botanica Marina. 2003;46:179-192. DOI: 10.1515/BOT.2003.018

[26] Guerrero-Pérez MO, Patience GS. Experimental methods in chemical
engineering: Fourier transform infrared spectroscopy—FTIR. Canadian Journal of Chemical Engineering. 2020;98:25-33. DOI: 10.1002/cjce.23664

[27] Roohinejad S, Koubaa M, Barba FJ, Saljoughian S, Amid M, Greiner R. Application of seaweeds to develop new food products with enhanced shelf-life, quality and health-related beneficial properties. Food Research International. 2017;99:1066-1083. DOI: 10.1016/j.foodres.2016.08.016

[28] Ismail AA, van de Voort FR, Sedman J. Chapter 4 Fourier transform infrared spectroscopy: Principles and applications. In: Paré JRJ, Bélanger JMR, editors. Tech. Instrum. Anal. Chem., vol. 18. Amsterdam, Netherlands: Elsevier; 1997. pp. 93-139. DOI: 10.1016/S0167-9244(97)80013-3

[29] Dutta A. Chapter 4 - Fourier transform infrared spectroscopy. In: Thomas S, Thomas R, Zachariah AK, Mishra RK, editors. Spectrosc. Methods Nanomater. Charact. Amsterdam, Netherlands: Elsevier; 2017. pp. 73-93. DOI: 10.1016/B978-0-323-46140-5.00004-2

[30] Beć KB, Grabska J, Huck CW. Chapter one - physical principles of infrared spectroscopy. In: Cozzolino D, editor. Compr. Anal. Chem., Vol. 98. Amsterdam, Netherlands: Elsevier; 2022. pp. 1-43. DOI: 10.1016/bs.coac.2020.08.001

[31] Berna F. Fourier transform infrared spectroscopy (FTIR). In: Gilbert AS, editor. Encycl. Geoarchaeology, Dordrecht: Springer Netherlands; 2017. pp. 285-286. DOI: 10.1007/978-1-4020-4409-0_15

[32] Baweja P, Kumar S, Sahoo D, Levine I. Chapter 3 - Biology of seaweeds. In: Fleurence J, Levine I, editors. Seaweed in Health and Disease

Prevention. San Diego, USA: Academic Press; 2016. pp. 41-106. DOI: 10.1016/B978-0-12-802772-1.00003-8

[33] Ushakiran MSB, Treasa MVM, Kaladharan P. Phycocelloid contents in certain economically important seaweeds of Kerala coast, India. Journal of the Marine Biological Association of India. 2021;63:5-11. DOI: 10.6024/jmbai.2021.63.2.2269-01

[34] Satpati GG, Sengupta S, Pal R. Seaweeds: The ecological roles, the economic benefits and the threats for changing the carbon cycle. In: Ranga Rao A, Ravishankar GA, editors. Sustain. Glob. Resour. Seaweeds Vol. 1 Bioresour. Cultiv. Trade Multifarious Appl. Cham: Springer International Publishing; 2022. pp. 295-311. DOI: 10.1007/978-3-030-91955-9_16

[35] Yoon HS, Nelson W, Lindstrom SC, Boo SM, Pueschel C, Qiu H, et al. Rhodophyta. In: Archibald JM, AGB S, Slamovits CH, editors. Handb. Protists. Cham: Springer International Publishing; 2017. pp. 89-133. DOI: 10.1007/978-3-319-28149-0_33

[36] Krueger-Hadfield SA, Shainker-Connelly SJ, Crowell RM, Vis ML. The eco-evolutionary importance of reproductive system variation in the macroalgae: Freshwater reds as a case study. Journal of Phycology. 2024;60:15-25. DOI: 10.1111/jpy.13407

[37] Cock JM, Godfroy O, Macaisne N, Peters AF, Coelho SM. Evolution and regulation of complex life cycles: A brown algal perspective. Current Opinion in Plant Biology. 2014;17:1-6. DOI: 10.1016/j.pbi.2013.09.004

[38] Bogaert KA, Zakka EE, Coelho SM, De Clerck O. Polarization of brown algal zygotes. Seminars in Cell &
Developmental Biology. 2023;134:90-102. DOI: 10.1016/j.semcdb.2022.03.008

[39] Bringloe TT, Starko S, Wade RM, Vieira C, Kawai H, De Clerck O, et al. Phylogeny and evolution of the Brown algae. Critical Reviews in Plant Sciences. 2020;39:281-321. DOI: 10.1080/07352689.2020.1787679

[40] Balar NB, Mantri VA. Insights into life cycle patterns, spore formation, induction of reproduction, biochemical and molecular aspects of sporulation in green algal genus *Ulva*: Implications for commercial cultivation. Journal of Applied Phycology. 2020;32:473-484. DOI: 10.1007/s10811-019-01959-7

[41] Sekimoto H. Sexual reproduction and sex determination in green algae. Journal of Plant Research. 2017;130:423-431. DOI: 10.1007/s10265-017-0908-6

[42] Pereira L. Macroalgae. Encyclopedia. 2021;1:177-188. DOI: 10.3390/encyclopedia1010017

[43] Fenoradosoa TA. Caractérisation Des Polysaccharides Extraits d'algues de Madagascar : *Halymenia Durvillei* (Rhodophyceae) et *Sargassum Turbinaroides* (Phaeophyceae). Phd Thesis. Université Blaise Pascal; 2009

[44] Zhang B, Gao Y, Zhang L, Zhou Y. The plant cell wall: Biosynthesis, construction, and functions. Journal of Integrative Plant Biology. 2021;63:251-272. DOI: 10.1111/jipb.13055

[45] Cassab GI, Varner JE. Cell Wall Proteins. Annual Review of Plant Biology. 1988;39:321-353. DOI: 10.1146/annurev.pp.39.060188.001541

[46] Anderson NS, Dolan TCS, Penman A, Rees DA, Mueller GP, Stancioff DJ, et al. Carrageenans. Part IV. Variations in the structure and gel properties of κ-carrageenan, and the characterisation

of sulphate esters by infrared spectroscopy. J Chem Soc C Org. 1968;0:602-606. DOI: 10.1039/J39680000602

[47] Pirsa S, Hafezi K. Hydrocolloids: Structure, preparation method, and application in food industry. Food Chemistry. 2023;399:133967. DOI: 10.1016/j.foodchem.2022.133967

[48] Kidgell JT, Magnusson M, de Nys R, Glasson CRK. Ulvan: A systematic review of extraction, composition and function. Algal Research. 2019;39:101422. DOI: 10.1016/j.algal.2019.101422

[49] Panggabean JA, Adiguna SP, Rahmawati SI, Ahmadi P, Zainuddin EN, Bayu A, et al. Antiviral activities of algal-based Sulfated polysaccharides. Molecules. 2022;27:1178. DOI: 10.3390/molecules27041178

[50] Hong T, Yin J-Y, Nie S-P, Xie M-Y. Applications of infrared spectroscopy in polysaccharide structural analysis: Progress, challenge and perspective. Food Chem X. 2021;12:100168. DOI: 10.1016/j.fochx.2021.100168

[51] Xiao N, Bock P, Antreich SJ, Staedler YM, Schönenberger J, Gierlinger N. From the soft to the hard: Changes in microchemistry during Cell Wall maturation of walnut shells. Frontiers in Plant Science. 2020;11:466. DOI: 10.3389/fpls.2020.00466

[52] Szymanska-Chargot M, Zdunek A. Use of FT-IR spectra and PCA to the bulk characterization of Cell Wall residues of fruits and vegetables along a fraction process. Food Biophysics. 2013;8:29-42. DOI: 10.1007/s11483-012-9279-7

[53] Wang S, Xu H, Luan H. Multiscale structures of starch granules. In: Wang S, editor. Starch Struct.
Funct. Appl. Foods. Singapore:
Springer; 2020. pp. 41-55.
DOI: 10.1007/978-981-15-0622-2_4

[54] Wang J, Wang Y, Xu L, Wu Q, Wang Q, Kong W, et al. Synthesis and structural features of phosphorylated *Artemisia sphaerocephala* polysaccharide. Carbohydrate Polymers. 2018;181:19-26. DOI: 10.1016/j.carbpol.2017.10.049

[55] Wang Y, Liu Y, Yu H, Zhou S, Zhang Z, Wu D, et al. Structural characterization and immuno-enhancing activity of a highly branched water-soluble β-glucan from the spores of *Ganoderma lucidum*. Carbohydrate Polymers. 2017;167:337-344. DOI: 10.1016/j.carbpol.2017.03.016

[56] Bayro AM, Manlusoc JK, Alonte R, Caniel C, Conde P, Embralino C. Preliminary characterization, antioxidant and Antiproliferative properties of polysaccharide from *Caulerpa taxifolia*. Pharm Sci Res. 2021;8:30-36. DOI: 10.7454/psrv.v8i1.1102

[57] Barakat KM, Ismail MM, Abou El Hassayeb HE, El Sersy NA, Elshobary ME. Chemical characterization and biological activities of ulvan extracted from *Ulva fasciata* (Chlorophyta). Rendiconti Lincei Sci Fis E Nat. 2022;33:829-841. DOI: 10.1007/s12210-022-01103-7

[58] Dalal SR, Hussein MH, El-Naggar NE-A, Mostafa SI, Shaaban-Dessuuki SA. Characterization of alginate extracted from *Sargassum latifolium* and its use in *Chlorella vulgaris* growth promotion and riboflavin drug delivery. Scientific Reports. 2021;11:16741. DOI: 10.1038/s41598-021-96202-0

[59] Fertah M, Belfkira A, Dahmane E, Montassir, Taourirte M, Brouillette F. Extraction and characterization of sodium alginate from Moroccan

*Laminaria digitata* brown seaweed.
Arabian Journal of Chemistry.
2017;10:S3707-S3714. DOI: 10.1016/j.arabjc.2014.05.003

[60] Mathlouthi M, Koenig JL. Vibrational spectra of carbohydrates. In: Tipson RS, Horton D, editors. Adv. Carbohydr. Chem. Biochem., vol. 44. Academic Press; 1987. pp. 7-89. DOI: 10.1016/S0065-2318(08)60077-3

[61] Lee Z-H, Lee M-F, Chen J-H, Tsou M-H, Wu Z-Y, Lee C-Z, et al. Fucoidan with three functions extracted from *Sargassum aquifolium* integrated rice-husk synthesis dual-imaging mesoporous silica nanoparticle. J Nanobiotechnology. 2022;20:298. DOI: 10.1186/s12951-022-01430-9

[62] Apostolova E, Lukova P, Baldzhieva A, Delattre C, Molinié R, Petit E, et al. Structural characterization and In vivo anti-inflammatory activity of Fucoidan from *Cystoseira crinita* (Desf.) Borry. Marine Drugs. 2022;20:714. DOI: 10.3390/md20110714

[63] Palanisamy S, Vinosha M, Manikandakrishnan M, Anjali R, Rajasekar P, Marudhupandi T, et al. Investigation of antioxidant and anticancer potential of fucoidan from *Sargassum polycystum*. International Journal of Biological Macromolecules. 2018;116:151-161. DOI: 10.1016/j.ijbiomac.2018.04.163

[64] Sellimi S, Benslima A, Barragan-Montero V, Hajji M, Nasri M. Polyphenolic-protein-polysaccharide ternary conjugates from *Cystoseira barbata* Tunisian seaweed as potential biopreservatives: Chemical, antioxidant and antimicrobial properties. International Journal of Biological Macromolecules. 2017;105:1375-1383. DOI: 10.1016/j.ijbiomac.2017.08.007
[65] Dammak M, Hadrich B, Miladi R, Barkallah M, Hentati F, Hachicha R, et al. Effects of nutritional conditions on growth and biochemical composition of *Tetraselmis* sp. Lipids in Health and Disease. 2017;**16**:41. DOI: 10.1186/s12944-016-0378-1

[66] Hentati F, Delattre C, Ursu AV, Desbrières J, Le Cerf D, Gardarin C, et al. Structural characterization and antioxidant activity of water-soluble polysaccharides from the Tunisian brown seaweed *Cystoseira compressa*. Carbohydrate Polymers. 2018;**198**:589-600. DOI: 10.1016/j.carbpol.2018.06.098

[67] Ermakova S, Men'shova R, Vishchuk O, Kim S-M, Um B-H, Isakov V, et al. Water-soluble polysaccharides from the brown alga *Eisenia bicyclis*: Structural characteristics and antitumor activity. Algal Research. 2013;2:51-58. DOI: 10.1016/j.algal.2012.10.002

[68] Rasheed I, Tabassum A, Khan U, Rehman A. Fourier transform infrared (FT-IR) spectroscopy of agar from red seaweeds of Karachi coast. International Journal of Biology and Biotechnology. 2019;**16**:53-63. DOI: 10.5772/53172

[69] Qari R, Haider S. Agar extraction, physical properties, FTIR analysis and biochemical composition of three edible species of red seaweeds *Gracilaria corticata* (J. Agardh), *Gracilaria dentata* (J. Agardh) and *Gracilariopsis longissima* (S. G. Gmelin) Steentoft, L. M; Irvine and Farnham. Pak J Sci Ind Res Ser B Biol Sci. 2021;**64**:263-273. DOI: 10.52763/PJSIR.BIOL.SCI.64.3.2021.263.273

[70] Pereira L, Gheda SF, Ribeiro-Claro PJA. Analysis by vibrational spectroscopy of seaweed polysaccharides with potential use in food, pharmaceutical, and cosmetic industries.

Int J Carbohydr Chem. 2013;**e537202**:7. DOI: 10.1155/2013/537202

[71] Santos GA. Carrageenans of species of *Eucheuma* J. Agardh and *Kappaphycus* Doty (Solieriaceae, Rhodophyta). Aquatic Botany. 1989;**36**:55-67. DOI: 10.1016/0304-3770(89)90091-0

[72] Mendoza WG, Montaño NE, Ganzon-Fortes ET, Villanueva RD. Chemical and gelling profile of ice-ice infected carrageenan from *Kappaphycus striatum* (Schmitz) Doty "sacol" strain (Solieriaceae, Gigartinales, Rhodophyta). Journal of Applied Phycology. 2002;**14**:409-418. DOI: 10.1023/A:1022178119120

[73] Kačuráková M, Wilson RH. Developments in mid-infrared FT-IR spectroscopy of selected carbohydrates. Carbohydrate Polymers. 2001;**44**:291-303. DOI: 10.1016/S0144-8617(00)00245-9

[74] Arman M, Qader SAU. Structural analysis of kappa-carrageenan isolated from *Hypnea musciformis* (red algae) and evaluation as an elicitor of plant defense mechanism. Carbohydrate Polymers. 2012;**88**:1264-1271. DOI: 10.1016/j.carbpol.2012.02.003

[75] Souza RB, Frota AF, Silva J, Alves C, Neugebauer AZ, Pinteus S, et al. *In vitro* activities of kappa-carrageenan isolated from red marine alga *Hypnea musciformis*: Antimicrobial, anticancer and neuroprotective potential. International Journal of Biological Macromolecules. 2018;**112**:1248-1256. DOI: 10.1016/j.ijbiomac.2018.02.029

[76] Vandanjon L, Burlot A-S, Zamanileha EF, Douzenel P, Ravelonandro PH, Bourgougnon N, et al. The use of FTIR spectroscopy as a tool for the seasonal variation analysis and for the quality control of polysaccharides
from seaweeds. Marine Drugs.

2023;21:482. DOI: 10.3390/md21090482

[77] Chopin T, Kerin BF, Mazerolle R. Phycocolloid chemistry as a taxonomic indicator of phylogeny in the Gigartinales, Rhodophyceae: A review and current developments using Fourier transform infrared diffuse reflectance spectroscopy. Phycological Research. 1999;47:167-188. DOI: 10.1046/j.1440-1835.1999.00170.x

[78] Emami-Karvani Z, Emtiazi G, Nahvi I, Ahadi A, Hashemol-Hosseini M. The combination of bacterial polymer and tragacanth to form antimicrobial biofilter for desalination. Desalination Water Treat. 2017;64:90-100.

DOI: 10.5004/dwt.2017.20134