Phytochemical Constituents, Antimicrobial Properties and Bioactivity of Marine Red Seaweed (Kappaphycus alvarezii) and Seagrass (Cymodocea serrulata)

The present work was performed to evaluate the levels of phytochemical constituents and the antioxidant and antibacterial properties of marine red seaweed (Kappaphycus alvarezii) and seagrass (Cymodocea serrulata). Quantitative phytochemical analysis, antioxidant activity and antimicrobial activity against 5 potential pathogenic bacteria was investigated. In both cases were found presence of flavonoids, tannins, phenolic compounds, glycosides, steroids, carbohydrates and ashes. Alkaloids were only found in K. alvarezii, buy not in C. serrulata. The antimicrobial properties of both K. alvarezii and C. serrulata chloroform extracts were found to be antagonis-tically effective against the gram-positive bacterium Bacillus subtilis and the gram-negative bacterium Vibrio parahaemolyticus, Vibrio alginolyticus, Vibrio harveyi and Klebsiella pneu-moniae. GC‒MS analysis revealed the presence of 94 bioactive compounds in K. alvarezii and 104 C. serrulata, including phenol, decane, dodecane, hexadecane, vanillin, heptadecane, diphenyla-mine, benzophenone, octadecanoic acid, dotriaconate, benzene, phytol, butanoic acid, and 2-hydroxyl-ethyl ether, which played a vital role in antioxidant and antibacterial activities. Thus, in view of the results, both K. alvarezii and C. serrulata could be considered as sources of ingre-dients with appreciable nutritional and medicinal value.

Keywords:

1. Introduction

Seaweeds are a group of autotrophic, halophytic and complex communities that live in marine environments that have potential renewable resources [1,2]. Biologically, they have been classified as Phaeophyta (Brown Algae), Rhodophyta (Red Algae) and Chlorophyta (Green Algae) [3]. Seaweeds grow in salt water, mainly in shallow coastal waters, and can be obtained for human consumption from both wild or cultivated form [4]. Their proximate and nutritional composition varies and is affected by a large variety of factors, including the seaweed species, geographic areas of origin, solar intensity or the seawater temperature [3].

Although in recent years the use of seaweeds as food has been increased in other parts of the world [3], seaweeds are still mostly consumed in Asian countries, such as Japan, China or Korea [3]. Seaweeds consumption has numerous advantages for human health, due to its content in dietary fibers, proteins, essential fatty acids, vitamins or essential minerals [5]. Seaweeds are also between the richest sources of bioactive primary and secondary metabolites, which are characterized by beneficial biological activities [1]. Between advantages for human health, seaweeds are known for their potential natural antioxidant, antiviral, antiobesity, antitumor and antimicrobial properties [5,6,7]. Additionally to their use as foods, other uses of seaweeds have been widely increased in recent decades and nowadays, seaweeds are also used as fertilizers, cosmetics, and their extracts were used in pharmaceutical industries as a fresh source of bioactive compounds with a wide range of medicinal properties [7].

Among seaweeds, Kappaphyus sp. is a commercially important red seaweed and is cultivated in tropical countries such as the Philippines, Indonesia, and Malaysia, as well as many countries in Eastern Africa [7], because it is relatively easy to cultivate, has short production cycles and has low production costs [7,8]. It is also a common food for the local people and is believed to have various beneficial effects. In India, the southeast coast has a unique marine habitat with a great variety of seaweed species spread around the intertidal zone and shallow- and deep-water regions of the ocean. Specifically, Kappaphycus alvarezii (commercially known as “cottoni”) grows well in the shores of the Kanyakumari and Ramanthapuram Districts of Tamil Nadu, India [9]. K. alvarezii has high economic value since is the principal source of the commercial hydrocolloid κ-carrageenan, and it also contains various inorganic and organic compounds that are beneficial for human health [7,12]. k-carrageenan is used in pharmaceuticals, cosmetics, textiles, organic fertilizers and in the food industry [12].

Other marine source of bioactive compounds with a broad spectrum of beneficial activities for human health are seagrasses [13]. Seagrasses are submerged flowering marine angiosperms living their full lifecycle submerged in marine environments, and they are the primary producers. They are the only flowering plants to recolonize the seaband, are highly productive and play an important ecological role in marine environments (Kim et al., 2021). Seagrasses are found in all coastal areas around the world except in Antarctica [14, 15]. Seagrass biomass are used in some countries as human food, and they are rich sources of secondary metabolites, such as alkaloids, flavonoids, terpenoids, tannins, steroids, and especially phenolic compounds [16]. The phenolic compounds present in seagrasses contribute to pigmentation, growth, reproduction, and resistance against pathogens, and they also act as defensive mechanisms against other aquatic lives as well as protection [17]. They have been used for traditional medicine, such as treating infections caused by pathogenic microbes, fever, inflammation, muscle pain, skin disease, viruses, diarrhea, diabetes, wound healing, sedation, and cancer [15]. Between the different genus forming seagrasses, Cymodocea, from Potamogetonaceae family, is represented worldwide by four species: Cymodocea rotundata, Cymodocea serrulata, Cymodocea angustata and Cymodocea nodosa [18]. C. serrulata is commonly found in the coastal area of the tropical Indo-West Pacific region. C. serrulata can be differentiated from other seagrass species by their shoots with distinctive open leaf scars, triangular, flat leaf sheath fibrous roots on the shoot and serrated leaf tips [14].

Hence, the aim of the present study was to evaluate the phytochemical constituents, antioxidant activity and antibacterial activity present in seaweeds and seagrass. The chemical compounds present in both K. alvarezii and C. serrulata were also determined.

2. Materials and Methods

2.1. Collection, identification and processing

The red seaweed K. alvarezii and the seagrass C. serrulata were obtained from Thondi coastal waters (Latitude: 9° 44” N and Longitude: 79° 00” E), Palk Bay, southeast coast of India. Freshly collected seaweed (K. alvarezii) and seagrass (C. serrulata) were cleaned thoroughly in seawater and transported to the laboratory within 1 h after collection. The epiphytes, necrotic parts, muds, dust and other debris were removed by washing thoroughly with fresh water and double distilled water. Then, they were shade dried at 25 ± 2°C for one week, ground into fine powder, and stored at room temperature in an airtight container (Tarsons, Chennai, India) for further analysis. The collected seaweed and seagrass were identified according to those established in the standard manual of Rao [19].

2.2. Preparation of extracts

The seaweed and seagrass extracts were prepared by adding 5 g of dried seaweed or seagrass powder into 50 mL of three different solvents, chloroform, ethanol and distilled water, in a conical flask and placed in a dark bottle in light agitation for 7 days. After that, the extracts were filtered through Whatman No. 1 filter papers and sterile cotton wools, and the supernatants were stored at 4°C for future use [20,21].

2.3. Determination of alkaloids

The alkaloid content of K. alvarezii and C. serrulata was determined by the method proposed by Hikino et al. [22]. One milliliter of test extract phosphate buffer (5 ml, pH 4.7) was added to 5 ml of bromocresol green solution, and the mixture was shaken vigorously added with 4 ml of chloroform. The extracts were collected in a 10 ml volumetric flask. The absorbance of the complex in chloroform was measured at 470 nm in UV‒Vis spectrophotometer (Shimadzu, Kyoto, Japan) against a blank prepared as described above but without extract. Atropine (Sigma-Aldrich, St. Louis, MO, USA) was used as a standard material, and the assay was compared with atropine equivalents.

2.4. Determination of flavonoids

Total flavonoid content was determined by the aluminum chloride method [23] using catechin (Sigma-Aldrich) as a standard. One milliliter of test sample and 4 ml of water were added to a volumetric flask (10 ml volume). After 5 min, 0.3 ml of 5% sodium nitrite and 0.3 ml of 10% aluminum chloride (Sisco Research Laboratories, Mumbai, India) were added. After 6 min of incubation at room temperature, 2 ml of 1 M sodium hydroxide (Sisco Research Laboratories) was added to the reaction mixture. Immediately, the final volume was brought to 10 ml with distilled water. The absorbance of the reaction mixture was spectrophotometrically measured at 510 nm against a blank in UV‒Vis spectrophotometer (Shimadzu). The results were expressed as catechin equivalents (mg catechin/g dried extract).

2.5. Determination of tannins

The total tannin content extracts were determined according to the Julkunen-Titto [24] method. 50 µl extracts were mixed with 1.5 ml of 40% vanillin (Sisco Research Laboratories) (prepared with methanol), and then 750 µl of HCl was added. The solution was shaken vigorously and left at room temperature for 20 min in darkness. The absorbance of the mixtures was measured at 500 nm using a spectrophotometer (Shimadzu). Catechin (Sigma-Aldrich) in the range of 20-200 mg/L was used to construct a calibration curve.

2.6. Determination of phenolic compounds

The total phenolic content in different solvent extracts was determined with Folin-Ciocalteu’s reagent proposed by Sangeeta and Vrunda [25]. In the procedure, different concentrations of the extracts were mixed with 0.4 ml Folin-Ciocalteu’s reagent (Sigma-Aldrich) (diluted 1:10 v/v). After 5 min, 4 ml of sodium carbonate solution was added. The final volume of the tubes was brought to 10 ml with distilled water and left for 90 min at room temperature. The absorbance of the sample was measured against the blank at 750 nm using a spectrophotometer (Shimadzu). A calibration curve was constructed using 1,2-dihydroxybenzen (catechol) (Sigma-Aldrich) solutions as standards, and the total phenolic content of the extract was expressed in terms of mg of catechol per g of dry weight.

2.7. Determination of cardiac glycosides

The cardiac glycoside content in the samples was evaluated using Buljet’s reagent as described by El-Olemy et al. [26]. One gram of the fine powder of K. alvarezii and C. serrulata was soaked in 10 ml of 70% alcohol for 2 hr. and then filtered. The extract obtained was then purified using lead acetate and Na2HPO4 solution before the addition of freshly prepared Buljet’s reagent (containing 95 ml aqueous Picric acid + 5 ml 10% aqueous NaOH) (Sigma-Aldrich). The difference between the intensity of colors of the experiment and blank samples gives the absorbance at 217 nm and is proportional to the concentration of the glycosides.

2.8. Determination of steroids

The steroid content was determined by Ejikeme et al. [24]. One milliliter of test extract of steroid solution was transferred into 10 ml volumetric flasks. Sulfuric acid) (Sisco Research Laboratories) (4 N, 2 ml and iron (III) chloride (Sisco Research Laboratories) (0.5% w/v, 2 ml) were added, followed by potassium hexacyanoferrate (III) solution) (Sisco Research Laboratories) (0.5% w/v, 0.5 ml). The mixture was heated in a water bath maintained at 70 ± 20°C for 30 minutes with occasional shaking and diluted to the mark with distilled water. The absorbance was measured at 780 nm spectrophotometer (Shimadzu) against the reagent blank.

2.9. Determination of carbohydrates

Carbohydrate content was estimated by the phenol‒sulfuric acid method [27]. Briefly, 200 mg of powdered sample was hydrolyzed by adding 5 ml of 2.5 N HCl. The sample was kept in a boiling water bath, and after 3 h of incubation, the solution was neutralized with solid Na2CO3 until effervescence ceased. The solution was made up to 50 ml and centrifuged at 8000 rpm for 10 min in a centrifuge (Remi Lab World, Mumbai, India). The supernatant was aliquoted and brought up to 1 ml with deionized water, to which 1 ml of phenol and 5 ml of 96 % sulfuric acid (Sisco Research Laboratories) were added. After mixing the solution, it was kept in a water bath at 25 ± 1°C for 20 min. The absorbance was measured at 490 nm using a UV‒Vis spectrophotometer (Shimadzu).

2.10. Ash content

The ash content was determined by the method of Yemm and Willis [27]. Two g of each sample was taken and weighed accurately in a clean silica dish. The dish was first heated over a low burner flame. After that, the dish was transferred to a SNOL muffle furnace (Utena, Lithuania) maintained at 3000°C-4500°C for 3-5 h. The ash residue obtained was then cooled in a desiccator and weighed. The percentage of total ash content was calculated by the formula as follows.

Total Ash Percent of plant sample (%) = [Weight of dry ash residue (g) ÷ Weight of plant sample (g)] x 100.

2.11. Hydrogen peroxide radical scavenging activity

The antioxidant activity of seaweed and seagrass extracts was evaluated by the hydrogen peroxide radical scavenging activity as described by Ebrahimzadeh et al. [28]. The seaweed and seagrass extracts (100 µg/mL) were reacted with 0.6 mL of 40 mM H2O2 solution prepared in phosphate buffer (pH 7.4) (Sisco Research Laboratories). After incubation at 37°C for 10 min, absorbance was measured at 230 nm using a UV‒Vis spectrophotometer (Shimadzu). Phosphate buffer was used as the corresponding blank solution. A similar procedure was repeated with distilled water instead of the extract, which served as a control. Ascorbic acid (Sigma-Aldrich) (20-100 µg/mL) was used as a standard.

2.10. In vitro antibacterial activity of seaweed and seagrass against human pathogenic bacteria

The antibacterial activity of seaweed and seagrass extracts was evaluated by the well diffusion method on Muller-Hinton agar (Hi-Media, Mumbai, India). Approximately 100 µl of 105 CFU/ml diluted inoculum of bacterial culture was applied on the surface of Muller-Hinton agar plates. The Muller-Hinton agar well was made with a well borer under aseptic conditions and filled with K. alvarezii and C. serrulata extracts and methanol served as positive controls. The plates were incubated at 37°C for bacterial growth, and the antibacterial activity of the seaweed and seagrass samples was evaluated by measuring the zone of inhibition (mm) against the tested pathogenic bacteria. All experiments were performed in triplicate, and the data are expressed as the mean values of the experiments.

2.11. Characterization of the active compound by gas chromatography‒mass spectrometry (GC‒MS)

The crude extracts of K. alvarezii and C. serrulata were loaded onto a silica gel (Hi-Media) packed column (20 cm length and 2 cm diameter) and eluted with n-hexane: ethyl acetate (50: 50 v/v) (Sigma-Aldrich). The fractions were characterized by gas chromatography GC-2010 interfaced with a quadrupole mass spectrometer QP-2010 (Shimadzu, Japan) analyzer to determine their chemical constituents using an Rtx-PCB capillary column (60 m x 0.25 mm i.d., 0.25 mm film thickness, Resteck, Bellefonte, PA). Helium with a purity of 99.99% was used as the carrier gas at a flow rate of 1 ml/min. One mil of extract was injected in spilt mode using an autosampler. The injector port, interface and ion source temperature were set at 250, 270 and 230°C, respectively. The GC temperature was programmed as follows: 50°C (1 min), 10°C (1 min) lamp to 320°C (10 min hold). The mass spectrometer was operated in electron ionization (EI) mode at 70 eV and at an emission current of 60 mA. Full scan data were obtained in a mass range of m/z 50-500. Interpretation of mass spectrum analysis was performed using the National Institute Standard and Technology (NIST) database. The spectrum of the unknown components was compared with the spectrum of known components stored in the NIST library.

2.12. Statistical analysis

All determinations were given in terms of the mean ± standard deviation (SD). The results obtained were compared by one-way analysis of variance (ANOVA). The significance of the difference between means was determined by Duncan’s multiple range test (P < 0.05) using SPPS version 14 (Chicago, IL, USA).

3. Results and discussion

In the present work, red seaweed K. alverazii and seagrass C. serrulata were tested for their phytochemical, antioxidant and antibacterial activities. Phytochemical analysis of K. alvarezii and C. serrulata revealed the presence of alkaloids (only in the case of K. alverazii) flavonoids, tannins, phenolic compounds, glycosides, steroids, carbohydrates and ashes. Among the six phytochemicals present in K. alvarezii, higher contents were found for phenolic compounds (3.39 ± 0.41 mg/g) and tannins (2.94 ± 0.41 mg/g). Among the five phytochemicals present in C. serrulata, the higher contents were found for glycosides (2.47 ± 0.41 mg/g) and flavonoids (2.11 ± 1.40 mg/g) (Table 1). These constituents significantly contributes to the biological activity of seaweeds and seagrass [29]. Similar observations were also made by other works [30,31,32], in which tannins, flavonoids, phenolic compounds, carotenoids and polysaccharides were found in both seaweed and seagrasses.

In the present study, K. alvarezii showed a higher tannins content (2.94 ± 0.41 mg Catechin equivalent (CAE)/g) than C. serrulata (1.94± 0.85 mg CAE/g). Similarly, Deyad and Ward [33] reported a similar tannins content in the brown seaweed Dictyota dichotoma (2.12 ± 0.45 mg CAE/g), whereas Domettila et al. [34] reported a higher presence of tannins in the red seaweed Sargassum wightii (27.54 ± 0.54 mg CAE/g). In previous studies [13], it was reported the presence of tannins in C. serrulata (264.71 mg/ml tannic acid equivalence). Similarly, another work reported the presence of tannins in seagrass Syringodium isoetifolium (80.65 ± 5.64 mg CAE/g [35]. Tannins are polyphenols which have a large influence on the nutritive value of humans and animals, due to its antimicrobial, anti-inflammatory, and astringent activities [13].

Flavonoids content was similar in both K. alvarezii and C. serrulata, although in global term it was found in lower amounts than in previous work. Vaghela et al. [35] found a much higher content (15.26 ± 0.95 mg CAE 100 g^-1). Similarly, Smadi et al. [36] reported the flavonoid content of C. nodosa as 3.98 ± 0.03 mg CAE/g, which is comparatively much higher than the results of the present study.

K. alvarezii showed an alkaloids content of 1.91 ± 0.58 mg CAE/g. Similarly, Domettila et al. [34] showed the alkaloids content of 1.32 ± 0.02 mg CAE/g in Ulva reticulata. Previously, Kubbat et al. [38] also reported alkaloids content in two brown algae Cystoseira compressa and Sargassum hornschuchii of 4125.00 ± 180.28 mg/g DW and 3708.33 ± 152.75 mg/g DW. Alkaloids are proved to have antiplasmodic, antimicrobial, and cytotoxic properties [38].

The phenolic compounds content in seaweeds are in part responsible of their scavenging activity, which protects against lipid oxidation [39]. In this work, K. alvarezii showed a higher phenolic content (3.39 ± 0.45 mg gallic acid equivalents (GAE)/g) than C. serrulata (1.01 ± 0.39 mg GAE/g). Previously, other authors reported a significantly higher content of phenolic compounds of both K. alvarezii (3.14 ± 0.14 mg GAE/g) [40] and Kappaphycus striatum (7.24 ± 0.21 mg GAE/g) [41]. Regarding C. serrulata, the results obtained in the current work were also significantly lower than those reported by Libin et al. [17] for C. serrulata (2.98 ± 0.12 mg GAE/g), and those reported for Cynodocea rotundata (2.04 ± 0.1) [42]. The phenolic content of seaweed and seagrass depends on the solvent used for the extraction, environment, habitat and biomass.

The presence of steroids in seaweed K. alvarezii (2.51 ± 0.15 mg/g) was higher than in seagrass C. serrulata (1.60 ± 0.24 mg/g). Previous, study also showed that the presence of steroids in seaweed C. elongata is 2.27 ± 0.26 mg/g [43]. Kumar et al. [44] also reported the presence of steroids in Champai parvula (24.30 ± 0.11 mg/g). Previously, it was also reported the presence of steroids Himanthalia elongata (2.64 ± 2.21 mg/g) [45]. In previous studies, Kannan et al. [46] reported the presence of steroids in C. rotundata (2.37 ± 1.27 mg/g). Similarly, Tango et al. [47] also reported the presence of steroids in seagrass Haludole pinifolia (5.62 ± 0.76 mg/g). Steroids isolated from seaweed and seagrass have medicinal values such as antihelmintic, antioxidant, antimicrobial and antiviral [45].

K. alvarezii showed a glycosides content of 1.88 ± 011 mg/g and in C. serrulata of 2.47 ± 0.28 mg/g. Previously, Kumar et al. [44] also reported the presence of glycosides in seaweed Cymodocea parvula (35.33 ± 0.14 mg/g). Similarly, Prabakaran et al. (2018) also reported the presence of glycosides in Chorella vulgaris (5.75 ± 0.23 mg/g). Deyad et al. [33] also reported the presence of glycosides in seaweed D. dichotoma (2.14 ± 0.15 mg/g). In a previous work done by Regalado et al. [50] showed the presence of glycosides in Thalassia testudinum (4.61 ± 1.60 mg/g). Glycosides are well known to lower the blood pressure in humans [44].

With respect to carbohydrates and ash content, the carbohydrate content of K. alvarezii was 2.57 ± 1.89 mg/g DW, and that of C. serrulata was 1.44 ± 1.75 mg/g DW. The wide variation in the carbohydrate content observed in seaweed and seagrass might be due to the influence of different factors, such as salinity, temperature, and sunlight intensity. Moreover, carbohydrate content is also influenced by biomass, which reveals the link between growth and carbohydrate content. Regarding ashes, K. alvarezii had a higher ash content (8.5 ± 0.95 g/100 g) than C. serrulata (6.9 ± 0.49 g/100 g). High ash content showed the presence of appreciable amounts of diverse minerals found in both seaweed and seagrass.

Antioxidant effectiveness is measured by monitoring the inhibition of oxidation of a suitable substrate [16]. In biological systems, antioxidant effectiveness is classified into two groups: evaluation of lipid peroxidation and measurement of free radical scavenging ability [28]. The in vitro antioxidant activity of K. alvarezii and C. serrulata extracts was evaluated by hydrogen peroxide radical scavenging activity K. alvarezii had higher scavenging activity (27.9 ± 0.1%) than C. serrulata (22.1 ± 0.1%). Regarding K. alvarezii, the results obtained were higher than those previously reported by other authors as Farah et al. [39], or Chew et al. [51], whose reported a lower (18.34 ± 0.57% and 11.8 ± 5.7%, respectively) 2,2-Diphenyl-1-picrylhydrazyl (DPPH) scavenging activity. Regarding C. serrulata, the DPPH scavenging activity obtained were lower than those obtained by Kannan et al. [52] (61.85 ± 0.95%) free radical scavenging activity from the same seagrass species, although higher than those reported by Rengasamy et al., 2013 (6.65 ± 0.12%) for other Cymodocea species, C. rotundata.

The antibacterial activity of both K. alvarezii and C. serrulata were investigated using chloroform extracts based on those reported by Pusparaj et al. [6], whose reported that best inhibitory effects of K. alvarezii was reported using chloroform extracts. The antibacterial activity of both K. alvarezii and C. serrulata depends on the presence of bioactive compounds, phenolic content and free radical scavenging activity [53]. In all case, it was detected inhibitory activity against the five pathogenic bacteria investigated (Table 2). The higher inhibitory activity was observed in K. alvarezii (26 ± 0.03 mm) against Bacillus subtilis, as well as in the case of C. serrulata exhibited maximum inhibitory activity (26 ± 0.08 mm) against Vibrio parahaemolyticus. The chloroform extract of K. alvarezii showed maximum activity of 26 ± 0.03 mm against B. subtilis at 100 µg/ml, and C. serrulata showed maximum activity of 26 ± 0.08 mm against V. parahaemolyticus at 100 µg/ml and minimum activity of 22±0.01 mm and 20±0.04 mm against Vibrio alginolyticus at 100 µg/ml in both K. alvarezii and C. serrulata, respectively (Table 2).

Jaswir et al. [53] reported the maximum inhibitory activity (12±1.02 mm) against B. subtilis using the methanolic extract of K. alvarezii. Similarly, Pusparaj et al. [6] reported the antibacterial activity of K. alvarezii against six human pathogens, Staphylococcus aureus, B. subtilis, Lactobacillus acidophilus, Pseudomonas aeruginosa, Escherichia coli and Proteus mirabillis, and he reported that the best activity was recorded in chloroform extracts. Kumar et al. [54] studied the antibacterial activity of C. serrulata against four fish-borne pathogens, namely, Bacillus cereus, B. subtilis, E. coli and Micrococcus luteus, and reported that C. serrulata was effective against several Bacillus sp.

The GC‒MS running time for the n-hexane:ethyl acetate (50:50 v/v) extracts of K. alvarezii and C. serrulata was 30 min. The target mass ions (m/z) and retention times of all identified compounds in K. alvarezii and C. serrulata are shown in Table 3 and Table 4. The results showed that K. alvarezii extracts contained 94 different bioactive compounds, including phenol, decane, dodecane, hexadecane, vanillin, heptadecane, diphenylamine, benzophenone, octadecanoic acid, dotriacontane and benzene (Table 3). In the other hand, C. serrulata was found to contain 104 different bioactive compounds, including tetradecane, dodecanal, dodecanal, diphenylamine, heptadecane, phytol, butanoic acid, 2-hydroxy-, ethyl ester, dodecane and benzene (Table 4). These compounds were responsible for the antioxidant and antibacterial activities of both K. alverazii and C. serrulata.

Datchanamurthy et al. [55] reported that red algae (Acoathophora deilei) contain major common components, such as hexadecanoic acid methyl ester, dibutyl phthalate, 2-ethyl butyric acid, octadecyl ester, 9-octadecanoic acid, methyl ester, and 1,2-benzendicarboxylic acid. Similarly, Manilal et al. [56] also reported that red algae (Asparagopsis taxiformis) contain components such as 4,5-dimethyl-1H-pyrrole-2carboxylic acid ethyl ester, chlorobenzene, 14-methyl-pentadecanoic acid methyl ester, octadec-9-enoic acid, 2,3-dihydroxy-propyl ester, 9-octadecanoic acid, pentadecanoic acid and octadecanoic acid, which might be involved in synergistic bioactivity. Anitha et al. [29] also studied the presence of phenols, hexadecanoic acid, n-hexadecanoic acid, tridecanoic acid, n-nonadecanoic acid, and benzene reported in red algae (Gracilaria cervicornis). Pushpabharathi et al. [13] reported 9 bioactive components in seagrass (C. serrulata): hexahydofarnesyl acetone, hexadecanoic acid, methyl ester, n-hexadecanoic acid, tetradecanoic acid, pentadecanoic acid, cholestesta 4,6 dien 3-ol and stigmasterol.

4. Conclusions

The red seaweed K. alvarezii and seagrass C. serrulata examined in the present study were found to possess rich sources of phytochemicals. The antioxidant properties of both seaweed and seagrass reveal that they have appreciable levels of protection against free radicals.

GC‒MS analysis revealed the presence of large active metabolites (94 in the case of K. alvarezii and 104 in the case of C. serrulate), such as phenol, decane, dodecane, hexadecane, vanillin, heptadecane, diphenylamine, benzophenone, octadecanoic acid, dotriacontane and benzene, in both red seaweed and seagrass. In view of the results obtained, both K. alvarezii and C. serrulata could be employed as potential marine drugs and may be used in the pharmaceutical and food processing industries as sources of ingredients with appreciable medicinal value. Since both red seaweed and seagrass were found to be good sources of essential phytochemicals, their commercial value can be enhanced by marketing them as value-added products.

Author Contributions

Conceptualization, S.P.; Formal analysis, D.D. and A.A.; Investigation, D.D. and S.P.; Data curation, S.P.; Literature data collection, A.L.S., and S.P.; Writing—original draft, A.A. and D.D.; Writing—review and editing, A.C.M. and A.L. Supervision, S.P and J.M.L. All authors have read and agreed to the published version of the manuscript.

Funding

The authors thank the European Regional Development Funds (FEDER), grant ED431C 2018/05, for covering the cost of publication.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

No new data were created or analyzed in this study. Data sharing is not applicable to this article.

Conflicts of Interest

The authors declare no conflicts of interest.

References

Anjum, A.; Aruna, G.; Noorjahan, C.M. Phytochemical analysis and antibacterial activity of selected seaweeds from coast of Mandapam, Tamilnadu. Indian J. Appl. Microbiol. 2014, 17, 50–58. [Google Scholar]
Ragunath, C.; Santhosh Kumar, Y.A.; Kanivalan, I.; Radhakrishnan, S. Phytochemical screening and GC‒MS analysis of bioactive constituents in the methanolic extract of Caulerpa racemosa (Forssk.) J. Agardh and Padina boergesenii Allender & Kraft. Curr. Appl. Sci. Technol. 2020, 20, 380–393. [Google Scholar]
Lopez-Santamarina, A.; Miranda, J.M.; Mondragón, A.C.; Lamas, A.; Cardelle-Cobas, A.; Franco, C.M.; Cepeda, A. Potential use of marine seaweeds as prebiotics: A review. Molecules 2020, 25, 1004. [Google Scholar] [CrossRef]
Balachandran, P.; Anson, S.M.; Ajay, K.T.V.; Parthasarathy, V. Preliminary phytochemical analysis of the ethanolic extract of brown Seaweed Sargassum wightii. Int. J. Res. Pharm. Sci. 2016, 7, 154–156. [Google Scholar]
Manivannan, K.; Kathiga Devi, G.; Ananthararaman, P.; Balasubramanian, T. Antimicrobial potential of selected brown seaweeds from vedalia coastal waters, Gulf of Mannar. Asian Pac. J. Trop. Biomed. 2011, 1, 114–120. [Google Scholar] [CrossRef]
Pushparaj, A. Antibacterial activity of Kappaphycus alvarezii and Ulva lactuca extracts against human pathogenic bacteria. Int. J. Curr. Microbiol. Appl. Sci. 2014, 3, 432–436. [Google Scholar]
Chin, Y.X.; Mi, Y.; Cao, W.X.; Lim, P.E.; Xue, C.H.; Tang, Q.J. A pilot study on anti-obesity mechanisms of Kappaphycus alvarezii: The role of native κ-carrageenan and the leftover sans-carrageenan fraction. Nutrients 2019, 11, 1133. [Google Scholar] [CrossRef] [PubMed]
Adharini, R. , Setyawan, A.R.; Suadi, S.; Jayanti, A.D. Comparison of nutritional composition in red and green strains of Kappaphycus alvarezii cultivated in Gorontalo Province, Indonesia. J. Appl. Phycol. 2020, 31, 725–730. [Google Scholar] [CrossRef]
Prasad, M.P.; Shekhar, S.; Babhulkar, A.P. Antibacterial activity of seaweed (Kappaphycus) extracts against infectious pathogens. Afr. J. Biotechnol. 2013, 12, 2968–2971. [Google Scholar] [CrossRef]
Veeramani, R.; Paramasivan, K.; Rose, A.H.R.; Manimmehalai, N.; Subhasri, D. Development and characterization of biodegradable film from marine red seaweed (Kappaphycus alvarezzi). Pigment Resin Technol. 2023, 52, 478–489. [Google Scholar] [CrossRef]
Mayakrishnan, A.; Srinivasan, P.; Kumari, V.; Kanakarajan, S.; Kamalanathan, A. Antimycobacterial Activity of Kappaphycus alvarezii against Mycobacterium tuberculosis and in silico molecular docking of kappa-carrageenan against InhA enzyme. Int. J. Drug Res. Tech. 2015, 5, 35–46. [Google Scholar]
Rupert, R.; Rodrigues, K.F.; Thien, V.Y.; Lym Yong, W.T. Carrageenan from Kappaphycus alvarezii (Rodophyta, Soliariaceae): Metabolism, structure, production, and application. Front. Plant. Sci. 2022, 13, 859635. [Google Scholar] [CrossRef]
Pushpabharathi, N.; Jayalakshmi, M.; Amudha, P.; Vanitha, V. Identification of bioactive compounds in Cymodocea serrulata seagrass by Gas Chromatography–Mass Spectroscopy. Asian J. Pharm. Clin. Res. 2018, 11, 317–320. [Google Scholar] [CrossRef]
Govindasamy, C.; Arulpriya, M.; Anantharaj, K.; Ruban, P.; Srinivasan, R. . Seasonal variations in seagrass biomass and productivity in Palk Bay, Bay of Bengal, India. Int. J. Biodivers. Conserv. 2013, 5, 408–417. [Google Scholar] [CrossRef]
Zulkifli, L.; Muksin, Y.D.; Hartanto, P.; Desimarlina, Y.; Idrus, A.A.; Syukur, A. 2021. Phytochemical profiles and ethnomedicine preliminary studies on seagrass species in the Southern Coast of Lombok Island Indonesia. Environ. Earth Sci. 2021, 913, 012102. [Google Scholar] [CrossRef]
Bharathi, N.P.; Jayalakshmi, M.; Amudha, P.; Vanitha, V. Phytochemical screening and in vitro antioxidant activity of the seagrass Cymodocea serrulata. Indian J. Mar. Sci. 2019, 48, 1216–1221. [Google Scholar]
Libin, B.; Sankar, T.V.; Chandramohana, K.N. Changes in phenolic compounds in seagrasses against changes in the ecosystem. J. Pharmacogn. Phytochem. 2017, 6, 742–747. [Google Scholar]
Dilipan, E.; Arulbalachandran, D. Genetic diversity of seagrass Cymodocea species as an ecological indicator on the Palk Bay Coast, India. Ecol. Genet Genom. 2022, 23, 100119. [Google Scholar] [CrossRef]
Rao, M. Key for identification of economical important seaweeds. Central Marine Fisheries Research Institute Bulletin, 1987, 41, 116. [Google Scholar]
Harborne, J.B. Phytochemical methods-A guide to modern techniques of plant analysis. Chapman and Hall, London. 1998, 5, 21–27. [Google Scholar]
Arulkumar, A.; Thomas, R.; Paramasivam, S. Phytochemical composition, in vitro antioxidant, antibacterial potential, and GC-MS analysis of red seaweeds (Gracilaria corticata and Gracilaria edulis) from Palk Bay, India. Biocatal. Agric. Biotechnol. 2018, 15, 63–71. [Google Scholar] [CrossRef]
Hikino, H.; Kiso, Y.; Wagner, H.; Fiebig, M. Antihepatotoxic actions of flavonolignans from Silybum marianum. Planta Med. 1984, 50, 284–250. [Google Scholar] [CrossRef] [PubMed]
Ejikeme, C.M., Ezeonu, C.S., Eboatu, A.N. Determination of physical and phytochemical constituents of some tropical timbers indigenous to Niger Delta area of Nigeria. Eur. Sci. J. 2014, 10, 247–270.
Julkunen-Titto, R. Phenolic constituents in the leaves of northern willows. Methods for the analysis of certain phenolics. J. Agric. Food Chem. 1985, 33, 213–217. [Google Scholar] [CrossRef]
Sangeeta, S.; Vrunda, V. Quantitative and qualitative analysis of phenolic and flavonoid content in Moringa oleifera. Pharmacognosy Res. 2016, 8, 16–21. [Google Scholar]
El-Olemy, M.M.; Al-Muhtadi, F.J.; Afifi, A.F.A. Experimental phytochemistry: A laboratory manual. King Saud University Press, Saudi Arabia, 1994, 21-27.
Yemm, E.W.; Willis, J. The estimation of carbohydrates in plant extracts by anthrone. Biochem. J. 1954, 57, 508–514. [Google Scholar] [CrossRef] [PubMed]
Ebrahimzadeh, M.A.; Nabavi, S.M.; Nabavi, S.F.; Bahramian, F.; Bekhradnia, A.R. Antioxidant and free radical scavenging activity of H. officinalis L. var. angustifolius, V. odorata, B. hyrcana and C. speciosum. Pak. J. Pharm. Sci. 2010, 23, 29–34. [Google Scholar]
Anitha, K.G.; Arputha, G.; Muthubala, G.; Susithra, R.; Mullaivendhan, M.; Anandham, R. GC‒MS Analysis of bioactive compounds of seaweed extracts collected from seashore of Manalmelkudi (Pudukkottai dist., Tamilnadu), responsible for antifungal activity. Int. J. Curr. Microbiol. App. Sci. 2019, 8(9), 2319–7706. [CrossRef]
Sheela, D.; Uthayakumari, F. GC‒MS analysis of bioactive constituents from coastal sand dune taxon - Sesuvium portulacastrum (L.). Biosci. Dis. 2013, 4, 47–53. [Google Scholar]
Mahabaleshwara, K.; Chandrasekhar, N.; Govindappa, M. Phytochemical investigations of methanol leaf extracts of Randia spinosa using column chromatography, HPTLC and GC‒MS. Natural Products Chemistry & Research, 2016, 4 (2). [CrossRef]
Rodriguez, D.; Carro, A.M.; Cela, R.; Lorenzo, R.A. Microwave-assisted extraction and large-volume injection gas chromatography-tandem mass spectrometry determination of multiresidue pesticides in edible seaweed. Anal. Bioanal. Chem. 2010, 398, 1005–1016. [Google Scholar] [CrossRef]
Deyad, M.; Ward, F. Qualitative and quantitative analysis of phytochemical studies on brown seaweed, Dictyota dichotoma. Int. J. Eng. Develop. Res. 2016, 4, 674–678. [Google Scholar]
Domettila, C.; Joselin, J.; Jeeva, S. Phytochemical analysis on some south Indian seaweeds. J. Chem. Pharm. Res. 2013, 5, 275–278. [Google Scholar]
Kalaivani, P.; Kavitha, D.; Amudha, P. In vitro antioxidant activity and phytochemical composition of Syringodium isoetifolium. Res. J. Pharm. Technol. 2021, 14, 6201–6206. [Google Scholar] [CrossRef]
Vaghela, P.; Das, A.K.; Trivedi, K.; Vijay, K.G.; Shinde, P.; Ghosh, A. Characterization and metabolomics profiling of Kappaphycus seaweed extract. Algal Res. 2022, 66, 1022774. [Google Scholar] [CrossRef]
Smadi, A.; Civatta, M.L.; Bitam, F.; Carbone, M.; Villani, G.; Gavagnin, M. Flavonoids and phenolic compounds from the Cymodocae nodosa. Plant Med. 2018, 84, 704–709. [Google Scholar] [CrossRef]
Alghaweer, R.; Azwai, S.; Garbaj, A.M.; Amr, A.; Elghmasi, S.; Sidati, M.; Yudiati, E.; Kubbat, M.G.; Eskandrani, A.A.; Shamlan, G.; Alansari, W.S. Alkaloids rich extracts from brown algae against multidrug-resistant bacteria by distinctive mode of action. Arab. J. Sci. Eng. 2021, 47, 179–188. [Google Scholar] [CrossRef]
Chan, P.T.; Matanjun, P.; Yasir, S.M.; Tan, T.S. Antioxidant activities and polyphenolics of various solvent extracts of red seaweed, Gracilaria changii. J. Appl. Phycol. 2015, 27, 2377–2386. [Google Scholar] [CrossRef]
Farah, D.; Abdullah, A.; Shahrul, H.; Chan, K. Antioxidant activity of red algae Kappaphycus alvarezii and Kappaphycus striatum. Int. Food Res. J. 2015, 22, 1977–1984. [Google Scholar]
Araújo, P.G.; Nardelli, A.E.; Fujii, M.T.; Chow, F. Antioxidant properties of different strains of Kappaphycus alvarezii (Rhodophyta) farmed on the Brazilian coast. Phycologia 2020, 53, 272–279. [Google Scholar] [CrossRef]
Dumay, O.; Costa, J.; Desiobert, J.M.; Pergent, G. Variations in the concentrations of phenolic compounds in the seagrass Cymodocae rotundata under conditions of competition. Phytochemistry 2004, 65, 3211–3220. [Google Scholar] [CrossRef]
Devi, K.N. Antibacterial activity of seagrass species of Cymodocea serrulata against chosen bacterial fish pathogens. Ann. Biol. Res. 2011, 2, 88–93. [Google Scholar]
Kumar, V.; Murugesan, S.; Bhuvaneswari, S. Phytochemical analysis of red alga Champia parvula (C. Agardh) collected from Mandapam coast of Tamil Nadu, India. Int. J. Adv. Pharm. 2015, 4, 15–20. [Google Scholar] [CrossRef]
Sanchez, M.; Lopez, D.L.; Seiro, J.P.L. An HPLC method for the quantification of sterols in edible seaweeds. Biomed Chromatogram. 2004, 18(1), 183–190. [Google Scholar] [CrossRef] [PubMed]
Kannan, R.R.; Arumugam, R.; Lyapparaj, P.; Thangaradjou, T.; Anantharaman, P. In vitro antibacterial, cytotoxicity and haemolytic activities and phytochemical analysis of seagrasses from the Gulf of Mannar, South India. Food Chem. 2013, 136(34), 1484–9. [Google Scholar] [CrossRef]
Tango, E.; Canencia, O.; Del Rosario, R.M. Phytochemical screening and proximate composition of the seagrass Halodule pinifolia of the coastal waters of Carmen, Agusan Del Norte, Philippines. Int. J. Modern Pharm. Res. 2021, 5, 75–80. [Google Scholar]
Prabakaran, G.; Moovendhan, M.; Arumugam, A.; Matharasi, A., Dineshkumar, R.; Sampathkumar, P. Quantitative analysis of phytochemical profile in marine microalgae Chlorella vulgaris. Int. J. Pharm. Biol. Sci. 2018, 8, 562–565.
Mohammed, D.; Elkatony, T.; and Ward, F. Qualitative and Quantitative Analysis of Phytochemical Studies on Brown Seaweed, Dictyota dichotoma. International Journal of Engineering Development and Research 2016, 4(2), 674–678. [Google Scholar]
Regalado, E.L.; Menendez, R.; Valdés, O.; Morales, R.A.; Laguna, A.; Thomas, O.P.; Hernandez, Y.; Nogueiras, C.; Kijjoa, A. Phytochemical analysis and antioxidant capacity of BM-21, a bioactive extract rich in polyphenolic metabolites from the Sea Grass Thalassia testudinum. Nat. Prod. Commun. 2011, 7, 47–50. [Google Scholar] [CrossRef]
Chew, Y.L.; Lim, Y.Y.; Omar, M.; Khoo, K.S. Antioxidant activity of three edible seaweeds from two areas in South East Asia. LWT-Food Sci. Tecnol. 2008, 41, 1067–1072. [Google Scholar] [CrossRef]
Kannan, R. R.; Arumugam, R.; Lyapparaj, P.; Thangaradjou, T.; Anantharaman, P. In vitro antibacterial, cytotoxicity and haemolytic activities and phytochemical analysis of seagrasses from the Gulf of Mannar, South India. Food Chem. 2013, 136, 1484–1489. [Google Scholar] [CrossRef] [PubMed]
Jaswir, I.; Tawakalit Tope, A.H.; Raus, R.A.; Monsur, H.A.; Ramli, N. Study on antibacterial potentials of some Malaysian brown seaweeds. Food Hydrocoll. 2014, 42, 275–279. [Google Scholar] [CrossRef]
Kumar, C.S.; Sarada, D.V.L.; Gideon, T.P.; Rengasamy, R. Antibacterial activity of three South Indian seagrasses, Cymodocea serrulata, Halophila ovalis and Zostera capensis. World J. Microbiol. Biotechnol. 2008, 24, 1989–1992. [Google Scholar] [CrossRef]
Datchanamurthy, B.; Narayanamurthy, U.; Anandh, S.J.V. Preliminary phytochemical and GC‒MS analysis of marine seaweed Acathophora deilei (Red alga). Biomed. Pharmacol. J. 2022, 15(3), 1695–1707. [Google Scholar] [CrossRef]
Manilal, A.; Sujith, S.; Sabarathnam, B.; Seghal Kiran, G.; Selvin, J.; Shakir, C.; Lipton, A.P. Bioactivity of the red algae Asparagopsis taxiformis collected from the southwestern coast of India. Braz. J. Oceanogr. 2010, 58, 93–100. [Google Scholar] [CrossRef]

Table 1. Steroids, tannins, flavonoid, glycosides, alkaloids and phenolic compounds of Kappaphycus alvarezii and Cymodocea serrulata.

Parameters	K. alverazii	C. serrulata
Alkaloids (ATE/g dry wt)	1.91±0.58*	-
Flavonoids (CAE/g dry wt)	1.63±2.73	2.11±1.40
Tannins (CAE/g dry wt)	2.94±0.41*	1.94±0.85
Phenolic Compound (GAE/g dry wt)	3.39±0.45*	1.01±0.39
Glycosides (mg/g dry wt)	1.88±0.11	2.47±0.28*
Steroids (mg/g dry wt)	2.51±0.15*	1.60±0.24
Carbohydrates (% DW)	2.57±1.89	1.44±1.75
Ash (% DW)	8.5±0.95	6.9±0.49
Antioxidant activity	27.9±0.1	22.1±01

Values are means of three analyses of the extracts ± standard deviation (n=3); CAE: Catechin equivalent; GAE: Gallic acid equivalents; ATE: Atropine equivalent.

Table 2. Antibacterial activity of K. alvarezii and C. serrulata in chloroform extract against human pathogenic bacteria.

Human Pathogenic Bacteria	Concentration (µg/ml)	Seaweed Extract	Seagrass Extract
Human Pathogenic Bacteria	Concentration (µg/ml)	Zone of inhibition (mm)
Bacillus subtilis	100	26±0.03	25±0.16
Klebsiella pneumoniae	100	23±0.01	22±0.20
Vibrio alginolyticus	100	22±0.01	20±0.04
Vibrio parahaemolyticus	100	24±0.02	26±0.08
Vibrio harveyi	100	24±0.10	22±0.01

Date were expressed as the mean ± SD values of triplicates (n=3).

Table 3. List of compounds identified from the purified extracts of K. alvarezii using GC-MS analysis.

No.	Name	Retention Time	Base m/z
1	Phenol	4.119	94.05
2	Cyclopropyl methyl carbinol	4.170	58.05
3	Decane	4.347	57.05
4	Butanoic acid, 2-hydroxy-, ethyl ester	4.429	59.05
5	2-Methylpentyl formate	4.457	56.05
6	Benzene, 1,4-dichloro-	4.530	145.95
7	Cyclopentane, 1,2-dimethyl-, cis-	4.568	70.10
8	Dodecane, 2,6,11-trimethyl-	5.158	57.05
9	Undecane, 5-methyl-	5.237	57.10
10	Ethane, hexachloro-	5.395	116.90
11	Dodecane, 2,6,10-trimethyl-	5.803	57.05
12	3-Ethyl-3-methylheptane	5.890	57.05
13	Naphthalene	6.994	128.10
14	Dodecane	7.201	57.05
15	Benzaldehyde, 2,5-dimethyl-	7.450	133.10
16	Octadecanoic acid, phenyl ester	7.517	94.05
17	Benzene, 1,3-bis(1,1-dimethylethyl)-	7.986	175.15
18	Undecane, 2,4-dimethyl-	8.093	57.10
19	Dodecane, 4,6-dimethyl-	8.317	57.05
20	Hexadecane	8.437	57.10
21	Formamide, N-phenyl-	8.884	121.05
22	Dodecane, 2,6,10-trimethyl-	8.942	57.10
23	Chloroxylenol	9.762	121.10
24	Benzene, 1-cyclobuten-1-yl-	9.844	129.10
25	Hexadecane	9.907	57.05
26	Vanillin	9.950	151.05
27	Heptadecane	10.186	57.05
28	Dodecane, 2,6,10-trimethyl-	10.690	57.10
29	1-Dodecanol	10.864	55.05
30	Nonadecane	11.005	85.10
31	Decane, 1-bromo-2-methyl-	11.044	57.05
32	Heneicosane	11.113	71.10
33	Nonadecane	11.159	57.05
34	2,4-Di-tert-butylphenol	11.351	191.15
35	Hexadecane	11.655	57.05
36	Hexadecane	12.350	57.05
37	Diphenylamine	12.664	169.15
38	Benzophenone	12.754	105.05
39	3-Hydroxydiphenylamine	13.216	185.10
40	Hexadecane, 2,6,10,14-tetramethyl-	13.328	57.10
41	Heptadecane	13.476	57.05
42	Dodecane, 2,6,10-trimethyl-	13.540	71.10
43	Heneicosane	13.590	71.10
44	Heneicosane	13.681	57.10
45	Decane, 1-iodo-	13.751	71.10
46	Heptadecane, 8-methyl-	13.940	71.10
47	Heneicosane	14.061	71.10
48	Tetradecanoic acid	14.148	57.05
49	Formamide, N,N-diphenyl-	14.487	168.10
50	Heneicosane	14.549	57.05
51	p-(Benzylideneamino)phenol	14.651	196.10
52	Isopropyl myristate	14.836	60.00
53	Carbamic chloride, diphenyl-	14.990	196.10
54	2-Pentadecanone, 6,10,14-trimethyl-	15.036	57.05
55	Phenoxazine	15.211	183.10
56	1,2-Benzenedicarboxylic acid, bis(2-methylpropyl) ester	15.305	149.05
57	Heneicosane	15.445	57.05
58	Hexadecane	15.569	57.05
59	Tetrapentacontane	15.735	71.10
60	7,9-Di-tert-butyl-1-oxaspiro(4,5)deca-6,9-diene-2,8-dione	15.837	57.05
61	3-Hydroxydiphenylamine	15.925	185.10
62	Benzoic acid, 2-benzoyl-, methyl ester	16.008	163.05
63	n-Hexadecanoic acid	16.227	73.05
64	7,9-Di-tert-butyl-1-oxaspiro(4,5)deca-6,9-diene-2,8-dione	16.414	57.05
65	Heneicosane	16.547	57.05
66	Cyclic octaatomic sulfur	16.953	63.95
67	Palmitic Acid, TMS derivative	17.018	117.05
68	Dotriacontane	17.125	57.05
69	Tetrapentacontane	17.472	57.05
70	Pentatriacontane	17.560	85.10
71	Octadecane, 3-ethyl-5-(2-ethylbutyl)-	17.635	71.10
72	Dotriacontane	17.723	57.05
73	Dotriacontane	17.820	71.10
74	Octadecanoic acid	18.063	73.05
75	Tetrapentacontane	18.120	71.10
76	Tetrapentacontane	18.193	71.10
77	Heneicosane	18.374	57.05
78	Tetracosane	19.231	57.10
79	Tetrapentacontane	19.605	71.10
80	1-Heptadecanamine	19.985	85.10
81	Heneicosane	20.056	57.05
82	Benzenemethanamine, N-hydroxy-N-(phenylmethyl)-	20.519	91.05
83	9-Octadecenenitrile, (Z)-	20.833	55.05
84	Bis(2-ethylhexyl) phthalate	21.292	149.05
85	13-Docosenamide, (Z)-	21.461	59.05
86	9-Octadecenamide, (Z)-	21.520	59.05
87	Dotriacontane	21.670	57.05
88	Tetracontane	22.656	57.05
89	13-Docosenamide, (Z)-	23.720	59.05
90	Squalene	24.338	69.05
91	Tetrapentacontane	25.429	57.05
92	13-Docosenamide, (Z)-	26.983	59.00
93	Dotriacontane	27.136	57.05
94	Cholesterol	28.552	386.35

Table 4. List of compounds identified from the purified extracts of C. serrulata using GC -MS analysis.

NO.	Name	R. Time	Base m/z
1	1-Trifluoroacetoxy-2-methylpentane	3.115	71.05
2	Propanoic acid, 2-hydroxy-2-methyl-	3.929	59.05
3	Cyclopropyl methyl carbinol	4.168	58.05
4	Carbamic acid, 2-(dimethylamino)ethyl ester	4.354	58.05
5	Butanoic acid, 2-hydroxy-, ethyl ester	4.427	59.05
6	2-Methylpentyl formate	4.455	71.05
7	1-Heptanol	4.565	70.10
8	Octane, 3,3-dimethyl-	4.656	71.10
9	3-Heptanol, 4-methyl-	4.780	59.05
10	Propane, 1,3-dichloro-	4.817	76.00
11	Dodecane, 2,6,10-trimethyl-	5.156	57.05
12	Dodecane, 4,6-dimethyl-	5.236	57.10
13	Ethane, hexachloro-	5.392	116.90
14	Dodecane, 2,6,10-trimethyl-	5.800	57.05
15	Naphthalene	6.993	128.10
16	Tetradecane	7.265	57.05
17	(Z),(Z)-2,4-Hexadiene	7.332	77.00
18	Decane, 2-methyl-	7.399	57.05
19	Benzaldehyde, 2,4-dimethyl-	7.455	133.10
20	Undecane, 4,8-dimethyl-	7.510	71.10
21	Tridecane	7.817	57.05
22	Tridecane	7.893	57.05
23	Benzene, 1,3-bis(1,1-dimethylethyl)-	7.983	175.15
24	Nonadecane	8.092	57.05
25	Dodecane, 4,6-dimethyl-	8.316	71.10
26	Nonadecane	8.431	57.05
27	Dodecane, 4,6-dimethyl-	8.507	71.10
28	Dodecane, 2,6,10-trimethyl-	8.615	71.10
29	Dodecane, 4,6-dimethyl-	8.940	71.10
30	Naphthalene, decahydro-1,4a-dimethyl-7-(1-methylethyl)-, [1S-(1.alpha.,4a.alpha.,7.alpha.,8a	9.200	57.05
31	Benzene, 1-cyclobuten-1-yl-	9.841	129.10
32	Hexadecane	9.903	57.05
33	Dodecanal	10.039	57.05
34	Heptadecane	10.184	57.05
35	Cyclotetrasiloxane, octamethyl-	10.471	281.05
36	Hexadecane	10.570	57.05
37	Dodecane, 2,6,10-trimethyl-	10.687	71.10
38	Heptane, 2,4-dimethyl-	10.720	85.10
39	Hexadecane, 1-bromo-	10.852	57.05
40	Undecane, 2,4-dimethyl-	11.005	85.10
41	Octane, 2-methyl-	11.039	71.10
42	Hexadecane	11.110	71.10
43	Tetradecane	11.157	57.05
44	Octadecane, 1-iodo-	11.220	57.05
45	2,4-Di-tert-butylphenol	11.349	191.15
46	Hexadecane	11.653	57.10
47	Dodecanoic acid	11.918	73.05
48	Octadecane	12.008	57.10
49	Hexadecane	12.346	57.10
50	1,4-Methanoazulen-9-ol, decahydro-1,5,5,8a-tetramethyl-, [1R-(1.alpha.,3a.beta.,4.alpha.,8a.	12.425	85.10
51	Heptadecane	12.515	57.05
52	Pentadecane, 4-methyl-	12.590	71.10
53	Diphenylamine	12.653	169.10
54	Heptadecane	12.749	57.05
55	Hexadecane, 2,6,10,14-tetramethyl-	13.057	57.05
56	Heneicosane	13.192	57.05
57	Heneicosane	13.249	57.05
58	Hexadecane	13.330	57.05
59	Dodecane, 1-iodo-	13.385	57.05
60	Heneicosane	13.483	57.05
61	Hexadecane	13.540	71.10
62	Heneicosane	13.588	71.10
63	Hexadecane	13.945	57.05
64	Heneicosane	14.058	71.10
65	Pentacosane	14.156	57.05
66	3,5-di-tert-Butyl-4-hydroxybenzaldehyde	14.250	219.15
67	Octadecane, 1-iodo-	14.320	57.05
68	Heneicosane	14.546	57.05
69	Octacosane	14.645	57.10
70	6-Octen-1-ol, 3,7-dimethyl-, acetate	14.966	68.05
71	Heneicosane	15.042	57.05
72	1-Tetradecanamine	15.151	59.05
73	Phytol	15.213	57.05
74	1,2-Benzenedicarboxylic acid, bis(2-methylpropyl) ester	15.302	149.05
75	Phytol	15.397	57.05
76	Hexadecane	15.445	57.05
77	Hexadecane	15.489	57.05
78	Tetracosane	15.650	57.10
79	Dotriacontane	15.695	71.10
80	Tetracosane	15.738	267.05
81	7,9-Di-tert-butyl-1-oxaspiro(4,5)deca-6,9-diene-2,8-dione	15.835	57.05
82	Benzoic acid, 2-benzoyl-, methyl ester	16.006	163.05
83	n-Hexadecanoic acid	16.202	73.05
84	Dibutyl phthalate	16.441	149.05
85	Heneicosane	16.544	57.05
86	Palmitic Acid, TMS derivative	17.010	117.10
87	Heneicosane	17.115	57.10
88	Dotriacontane	17.246	57.05
89	Heneicosane	17.476	57.05
90	Tetrapentacontane	17.564	57.05
91	Phytol	17.631	71.10
92	Tetracosane	17.725	57.05
93	Tetrapentacontane	17.817	71.10
94	Octadecanoic acid	18.056	57.05
95	Tetrapentacontane	18.116	71.10
96	Tetrapentacontane	18.190	71.10
97	Docosane	18.371	57.05
98	4-Morpholinepropanamine	18.495	100.05
99	3-Isopropyl-2,5-piperazine-dione	18.583	114.10
100	Heptadecane, 2-methyl-	18.945	57.05
101	Tetrapentacontane	18.997	57.10
102	Heneicosane	19.229	57.05
103	Tetracosane	19.450	57.05
104	Tetrapentacontane	19.529	71.10

Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

© 2023 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

Copyright: This open access article is published under a Creative Commons CC BY 4.0 license, which permit the free download, distribution, and reuse, provided that the author and preprint are cited in any reuse.

Alerts

MDPI Initiatives

Important Links

Choose an area of interest and we will send you notifications of new preprints at your preferred frequency.

Disclaimer