06
MICROMORPHOLOGICAL, HISTOLOGICAL AND PHYTOCHEMICALCHARACTERIZATION OF SEAGRASSES CYMODOCEA SERRULATA ANDSYRINGODIUM ISOETIFOLIUM OF PALK BAY REGION, TAMILNADU
Jaya Durga Jaisankar1 and Arumugam Ramasubramanian2*
1Research student, PG & Research Department of Botany,
Alagappa Government Arts College, Karaikudi, Tamil Nādu, India
2Assistant Professors, PG & Research Department of Botany,
Alagappa Government Arts College, Karaikudi, Tamil Nādu, India*Corresponding Author: Email: drmugam@yahoo.co.in

How to cite this paper:
Jaya Durga Jaisankar andArumugam Ramasubramanian,(2022)

Micromorphological,

histological and phytochemical
characterization of seagrasses
Cymodocea serrulata and
Syringodium isoetifolium of
Palk Bay region, Tamil Nādu,
Journal of Global Resources,
Vol. 08 (02)DOI:
10.46587/JGR.2022.v08i02.006

Received: 18 April 2022
Reviewed: 01 May 2022
Revised: 21 May 2022
Final Accepted: 25 June 2022

Abstract: Rapidly expanding scientific knowledge on
seagrasses has led to a growing awareness that seagrasses
area valuable coastal resource. The present study is aimed
to validate the micro-morphometric parameters andphytochemical profiling of two sea grasses Cymodocea
serrulataandSyringodium
isoetifolium
throughhistochemical localization and molecular analysis. The
preliminary phytochemical studies of both the seagrasses
explored variety of biologically active compounds such as
flavonoids, terpenoids, alkaloids, phenols, tannins,
glycosides and coumarin. The micro-morphometricparameters like fiber bundles, air lacunae, branching patternand hairs in root have significant role in taxonomic
identification and delimitation of Cymodocea & Syringodium
from other sea grass genera of Palk Bay. The histochemical
localization coincided with qualitative phytochemical
analysis and also confirmed the sites of production or
accumulation of secondary metabolites in the tissues.
Antioxidant activity of 88.4 percent was observed in the leaf
extract of C. serrulata with the IC50 value of 115.7 µg/ml and86.3 percent in S. isoetifolium with IC50 value of 122.42µg/ml. The present findings on the strong antioxidant
properties of both the seagrasses significantly correlatedwith the total phenolic content. The FTIR analysis showedabsorption peaks assigned to various functional groups
related to flavonoids, phenols and coumaric acid. Further
research studies are needed to characterize thephytochemicals of these seagrasses at molecular level using
HPLC and GCMS technique and those results can beextrapolated to clinical studies.
Key words: Seagrass, Anatomy, Histochemistry, FTIR,
DPPH, Phytochemistry

Journal of Global Resources, July 2022, Volume 08 (02) J. D. Jaisankar and A. Ramasubramanian P a g e | 48

Introduction
Seagrasses are marine angiosperms found in all coastal areas of the world except in Antarctic(Duarte and Gattuso, 2010). Seagrasses continue to decline at a global rate of approximately
7 percent (Waycott et al. 2009), and the losses may be more acute in the Southeast Asian
region due to the pressure from increasing coastal populations and developments, and thelack of data on seagrass resources (Thangaradjou et al., 2008). Seagrass flora of India isrepresented by 6 genera and 14 species, Gulf of Mannar and Palk Bay harbour the maximum
species followed by Andaman and Nicobar and Lakshadweep islands. Seagrasses provide awide range of ecosystem services that directly or indirectly benefit human needs. But recent
research has shown that there are still many gaps in our comprehension of seagrassecosystem service provision (Nordlund et al., 2016). Despite being a key component of marine
ecosystems, seagrass meadows are a prime example of a habitat that is largely understudied
and under documented in the Southeast Asian region (Ooi et al., 2011). The previous works
reflects only the conditions of seagrass meadows, seagrass phenology, morphology and theirdistribution but not the importance of seagrasses and their great nutritional value.
Seagrass of the Gulf of Mannar biosphere were found to be endowed with nutraceuticalcharacteristics and proved to be a novel source of antioxidants and literature revealed lack ofadequate scientific investigation on vitamins and minerals, amino acid and fatty acid profile ofIndian seagrass (Jeevitha et al., 2013). Among the 72 species of seagrass present worldwide,
only a few have been explored for their natural products and their determinant role (Subhashini
et al., 2013). The acquisition and allocation patterns of biochemicals have been extensively
studied in many terrestrial plants, but little has been done on the submerged aquatic vegetationsuch as sea grasses (Mbaga and Mvungi, 2019). Recent research increases the search on
novel compounds from seagrass, since they have high contents of antioxidants likepolyphenols, terpenoids, flavonoids, tannins, and saponins (Ansari and Ghanem, 2019). The
compounds present in seagrasses are highly antioxidative in nature, and under stressconditions, they produce secondary metabolites as defense mechanism (Amudha et al., 2017;
Subhashini et al., 2013). The search for medicinal plants with therapeutic utility is a very old
practice, which currently plays an important role in modern medicine. Though, few studies havebeen conducted to ensure the quality of the raw material, the microscopic study is essential to
the standardization of plants used as medicines (Moreira et al., 2014). According to Upton etal. (2011), the preliminary set of pharmacogenetic tools used for quality assessment of
medicinally valuable plant parts is macro- and micro-anatomy. In view of bioprospecting
coastal resources, the present study was aimed to study the anatomical, histological andphytochemical characteristics of two dominant sea grasses Cymodocea serrulata (R. Brown)
Ascherson & Syringodium isoetifolium (Ascherson) Dandy from the Palk Bay coastal zone.
Materials and Methods
Monthly sea grass samplings were carried out from different hotspots along the Palk Bay viz.,
Kattumavadi (10° 13‘ N; 79° 23‘ E), Kodiyakarai (10° 16‘ N; 79° 49‘ E), Manamelkudi (10°02‘
N; 79°15‘ E), Kottaipattinam (09° 59‘ N; 79°13‘ E), Mimisal (09° 54‘ N; 79° 08‘ E), S. P. Pattinam(09° 51‘ N; 79° 05‘ E), Karankadu (09° 64‘ N; 78° 96‘ E) and Thondi (Lat.09°44‘N;
Long.79°01‘E) from April 2020 to March 2021. Seagrass biomass was assessed from the sites
during low tides by the use of 1m2 quadrate and taxonomic identification was carried out withthe help of seagrass manual (Kannan and Thangaradjou, 2006). Collected sea grasses were
thoroughly washed, separated into root, rhizome, leaves and then dried under shade at 30 ± 2ºC for about 10 days.

Journal of Global Resources, July 2022, Volume 08 (02) J. D. Jaisankar and A. Ramasubramanian P a g e | 49

Anatomical Studies and Histological Localization
Free hand cross sections of root, stem and leaf were taken from the fresh material fixed in FAA
50 percent (Johansen, 1940), washed in distilled water and stained according to the techniquedescribed by Bukatsch (1972), with safranin. Similarly, the sections were also treated withrespective reagents used in qualitative phytochemical analysis to localize specific components,
viz alkaloids, flavonoids, phenol, lignin and tannin in the tissues. Presence of cutin was
confirmed by the appearance of red colour when fresh sections were treated with saturated
solution of Sudan IV prepared in 70 percent ethanol (Margolena, 1932). Presence of starchgrains were detected by appearance of blue colour when treated with potassium iodide
solution, which was made by dissolving 3 gm of iodine and 1.5 gm of potassium iodide in 100
ml of water. Lignin was tested by occurrence of red colour after treating lignified portion withphloroglucinol solution followed by 1-2 drops of 2.5 percent hydrochloric acid (Sass, 1951). To
test the presence of tannin, fresh sections were placed in 10 percent aqueous ferric chloride
solution with a pinch of sodium carbonate and blue colour of tannin substances was observed
under the microscope. Dragondorff’s reagent was used to visualize alkaloids and toluidine blue
was used for localizing various polysaccharides (O’Brien et al., 1964). The sections weretreated with ferric chloride and Sudan III to localize phenolic (Reeve, 1951) and lipophiliccompounds (Foster, 1949). These reagents were chosen based on their specific application inhistochemistry and chromatography techniques for the detection of the respective compounds.
All sections were viewed under the light microscope (Zeiss Axioscope 2plus ® Germany) andphotographed (ULTRA Scope image 9.0V).
Screening of Phytochemicals
The air-dried plant parts were pulverized to a coarse powder in a mechanical grinder, passed
through a 40-mesh sieve and extracted in a soxhlet extractor with ethanol, water and ethyl
acetate (Sunita et al., 2012). The extract was decanted, filtered with Whatman No. 1 filter paperand concentrated at reduced pressure below 40◦C through rota-vapor to obtain dry extract thensubjected to further analysis. The extracts were subjected to qualitative phytochemical analysis
for the presence of tannins, flavonoids, alkaloids, terpenoids, coumarin, glycosides andphenols using standard procedures. The preparations of the reagents are given in Table 1.
Chlorophyll content (mg g-1 fresh weight) was determined through organic solvent (80 percent
acetone) extraction method as described by Arnon (1949). Total phenols were quantitatively
estimated using the method of Lin and Tang (2007).
Anti-Oxidative Radical Scavenging Activity
The anti-oxidative activity was assessed on the basis of the radical-scavenging effect of thestable DPPH (1-Diphenyl-2-Picryl Hydrazine)free radical. The concentration of DPPH will be
kept at 300 μM in MeOH. 1 mg of the extracts dissolved separately in 1 mL of MeOH, fromwhich different concentrations (0-200 μg/mL) will be prepared. 10 μL of each extract solution
will be allowed to react with 200 μL DPPH at 37°C for 30 min in a 96-well microliter plate. Afterincubation, decrease in absorption for each solution will be measured at 490nm using a
microplate reader. Ascorbic acid will be used as reference (Mensor, 2001).
FTIR Analysis
Crude sample for Fourier Transform Infrared spectroscopy (FT-IR) analysis was prepared by
KBr pellet technique. IR analysis of crude ethanolic extract of C. serrulata and S. isoetifolium
was carried out in 400–4,000 cm−1 range and spectra were recorded.
RESULTS & DISCUSSION
Diversity, Growth and Biomass
In the present study, a preliminary survey was made to document the distribution of available
sea grasses in the sampling sites over the seasons. Five seagrass species under 3 genera
were recorded in the sampling sites and the major groups of seagrasses found in all the
Journal of Global Resources, July 2022, Volume 08 (02) J. D. Jaisankar and A. Ramasubramanian P a g e | 50

sampling sites are Cymodocea serrulata and Syringodium isoetifolium (Gokulakrishnan and
Ravikumar, 2016). C. serrulata was found to be dominant in Kodiyakarai whereas rarely
Halophila ovalis was recorded. Seagrass biomass values varied highly with respect to thespecies in different seasons, whereas within the sites the values are insignificant. The biomassof S. isoetifolium varied between 106.2 and 441.7g FWT m-2 registering the minimum at Thondi
and the maximum at Kodiyakarai. Similarly, the minimum biomass of 204.7g FWT m-2 of C.
serrulata was recorded at Karankadu and the maximum of 744.3g FWT m-2 was recorded at
Kodiyakarai during post monsoon (Jan – Feb. 2021). Generally, the below -ground biomass
was higher than above–ground biomass, which might indicate the survival strategies ofseagrasses to minimize anthropogenic pressure, exposure to desiccation at low tide and toincrease stability when exposed to high tides. Similar to our results, Govindasamy et al. (2013)
reported maximum biomass for C. serrulata (1429 g FWT m-2) during post monsoon in the Palk
Bay region. Earlier reports found the distribution of seagrasses up to 3-meter depth and thepresent study found the distribution even beyond that, extending up to 5-meter depths.
Temporal variability in C. serrulata and S. isoetifolium abundance has been found to occur in
response to either anthropogenic or natural disturbances (Green and Short 2003). Coles et al.
(2011) reported that many environmental biophysical parameters, the availability of seeds andvegetative fragments, and anthropogenic inputs can determine the presence of seagrasses
along coastal areas and regulate the growth and morphology of seagrasses.
Anatomical Characterization
Seagrasses are monocotyledonous angiosperms, generally have a similar external
morphology of other flowering plants with a well-developed creeping rhizome. The findings ofthe present study was supported by another study, which showed that Thalassia testudinum
had the highest number of leaves per shoot, high shoot density and the greatest biomass due
to nutrient and sediment input from the nearby mangrove community (Arrelano-Mendez et al.,2011).The ability of seagrass to occupy an area is also inseparable from the rhizome ability todevelop its root system on different types of substrates and indicates morphological plasticity(Balestri et al. 2015). The expansion of the rooting system and its penetration into the substrate
are essential for the growth of new seagrasses to overcome physical disturbance and the low
availability of nutrients.
In C. serrulata, at rhizome nodes, there is a short erect shoot bearing several foliage
leaves with serrate margin, each with a sheath at the base. Each leaf consists of a basal leaf
sheath and a distal leaf blade, these leaves usually are long linear and have a similar adaxial
and abaxial surface in cross section whereas S. isoetifolium has terete leaves (Kuo et al.,2018). The leaf cuticle was very thin (0.1 – 0.2µm) in both the species but C.serrulata showed
subcuticular cavities. A fiber bundle usually consists of several fiber cells, associated only withthe hypodermis adjacent to the longitudinal vascular bundles are observed in C. serrulata andthey are absent in S. isoetifolium. The fiber bundles and the thickened walls of the epidermal
cells provide tensile strength but retain a high degree of flexibility that allows the strap-like leaf
blades to withstand vigorous wave action (de los Santos et al., 2016). The strap-like leaves of
C. serrulata have several parallel longitudinal vascular bundles, which are connected bysmaller transverse bundles. The longitudinal vascular bundles in both species are separated
by prominent regularly arranged air lacunae of varying size. The wall of bundle sheath cells isthin, lignified in C. serrulata and thick, suberized in S. isoetifolium. The number and size ofxylem elements in the vascular bundles of seagrasses, asin other aquatic plants, are much
reduced in comparison with those in vascular land plants (Kuo et al., 2018).

Journal of Global Resources, July 2022, Volume 08 (02) J. D. Jaisankar and A. Ramasubramanian P a g e | 51

Compound
Alkaloids

Coumarin
Flavonoids
Phenols

Tannins

Terpenoids
Anthraquinone

Cardiacglycoside

Sterols,
Triterpenoids

Table 01: Preparation of Reagents and Qualitative Tests for Phytochemicals
Reagents
Test
Appearance
Mayer’s reagent
1ml of the extract was measured into a Yellow colour
watch glass and little amount of dilute Orange colour
Dragendroff’s test
HCl and reagents were added
Wagner’s test
Reddish brown
Sodium Hydroxide
1.5 ml of the extract was mixed with few Yellow colour
drops of alcoholic sodium hydroxide
Magnesium ribbon
1.3 ml of the extract was mixed with 0.5 Orange to red colour
g of magnesium turnings
Ferric chloride (1 percent) A few drops of ferric chloride solution Bluish green colourin 50 percent aqueous
were added to 2ml of the extract
methanolFerric chloride (0.1
To 2 ml of extract, 2-3 drops of 0.1 Blue black colour
percent)
percent ferric chloride solution was
added
Chloroform
5 ml of each extract mixed in 2 ml of
A reddish brown
Chloroform and 3 ml conc. H2SO4
colour at the interface
10
percent
Ammonia 5 ml of chloroform to 0.5 g of extract in a Formation of a rosesolution
clean dry test tube. This was shaken for pink color in the
5 min and the extract thereafter filtered. ammoniacal
layer
The filtrate was then mixed with equal (lower layer)
volume of 10 percent ammonia solution.
Glacial acetic acid
0.5 g of extracts dissolved in 2 ml of Brownring
wasglacial acetic acid, 1 drop of ferric formed at interphasechloride, 2ml of conc. sulphuric acidindicated the presenceof cardiac glycoside.
Chloroform
The extract added with a few drops of Appearance of red ring
chloroform and few drops of acetic at the junction of two
anhydride and mixed well. 1 ml of con. layers- triterpenoids.
H2SO4 was added from the sides of the Appearanceofatest tube
brown ring at thejunction of two layers sterols.

References
Sofowara, 1993

Yves-Alain et al.,
2007
Sofowara, 1993
Cook andSamman, 1996
Chung et al., 1998

Aryakrishna et al.,2016
Sorescu et al., 2018

Harborne, 1998

Bruneton, 2009

Journal of Global Resources, July 2022, Volume 08 (02) J. D. Jaisankar and A. Ramasubramanian P a g e | 52

The rhizomes of C. serrulata and S. isoetifolium are herbaceous, cylindrical with scars
at irregular interval (Kendrick et al., 2012). The internodes of rhizomes and erect shoots appear
similar in transverse section. The outermost layer is the epidermis covered by a distinct cuticle
contained tannin. The bulk of rhizome and stem is cortical tissue, the structure of which showed
great variations. In S. isoetifolium the cortex has large lacunae with parenchymatous septa. In
both the species, the cortex has distinct thick-walled hypodermis, the outer corticalparenchyma cells with thick, lignified walls and the inner one having thin, non-lignified walls.
The innermost layer of cortical cells is the endodermis that have thickened and lignified walls
which surrounds the central stele. A central stele contains one or more larger central
protoxylem elements surrounded by sieve tubes; the pericycle is not distinct.
The roots of both seagrasses are adventitious as in all monocotyledons, and arise fromthe ventral side of the rhizome nodes. The external morphology of roots often has characteristicfeatures in different taxa, with adaptation to different substrates, though the internal anatomy
is generally similar (Kiswara et al., 2009). C. serrulata and S. isoetifolium have at each rhizome
node, one or more moderately branched roots with few root hairs. The branching pattern ofroot and the presence of root hairs have sometimes been used for the taxonomic delimitationof Cymodocea & Syringodium from other genera, i.e., Amphibolis (no root hairs) and
Thalassodendron (Rarely branched) (Kuo et al., 2018). The mature root has an epidermis,
overlaying cortical parenchymatous tissue that encloses air lacunae and a central stele. The
epidermal cells usually have thin, undignified walls and below the epidermis there is a distinct
exodermis one or more cells thick, each of which has thickened but undignified walls. The
cortex of root showed an outer layer with larger cells, large air lacunae separated by radial
partitions at the middle cortex and an inner cortex consisting of compact, small cells in a
regular, concentric arrangement. As similar to monocotyledons, the stele is surrounded by alayer of small, compact endodermal cells with thin to moderately thick walls, non-lignified (delos Santos et al., 2016). Anatomical characterization is an inherent part of practically all
pharmacopoeias and is one of the primary identification tests necessary for pharmacopoeia
compliance. The individual structural elements are comparatively frequent within the sameplant organs, but the ways in which the tissues, elements and cells are set with in a plant organ
permits diagnosis to be performed and lend support to the quality assessment of herbal drugs(Upton et al., 2011).

Fig 1. Anatomy and histochemical localization (a) C. serrulata leaf (b-c) C. serrulata stem showing central stele and air lacunae (d) C.
serrulata rhizome showing hypodermal fiber bundles (e) S. isoetifolium leaf (f) S. isoetifolium rhizome central stele and air lacunae (g)Suberized endodermis of S. isoetifolium rhizome (h) Cutin layer of S. isoetifolium rhizome (i) lignified walls of fiber bundles of C. serrulata
rhizome (i-k) Phenols & Tannin in blade epidermis and rhizome of C. serrulata (l) Lipid droplets in the cortex of S. isoetifolium leaf

Journal of Global Resources, July 2022, Volume 08 (02) J. D. Jaisankar and A. Ramasubramanian P a g e | 53

Histochemical Localization
The thin-walled mesophyll parenchyma cells and the cortical parenchyma cells of rhizome of
C. serrulata showed the presence of small starch grains (Kuo et al., 2018). Tannin wasobserved in the cuticle of rhizome and walls of blade epidermal cells (Fig. 1k). The Sudan IIIstained endodermal cells at the stellar region and bundle sheath cells showed the presence oflignin in both the species. The cell walls of hypodermis and cortical parenchyma cells of therhizome and stem stained with T blue showed purple colour which confirmed the presence ofpolysaccharides. Sudan III-stained lipids (Fig. 1l) which were observed as reddish granules in
the upper mesophyll cells of leaf, in the outer cortical cells of rhizome of C. serrulata and very
few in the inner cortical region of rhizome of S. isoetifolium. Alkaloids were noticed in the crosssections of rhizome of C. serrulata, but not in leaf and root. Thin layer of cutin was observed
as red layer in the rhizome of S. isoetifolium (Fig. 1h). Cutin is the main component of the
cuticle and isa lipophilic polymer that is deposited in and the top of the Outerwall epidermal
cells (Upton et al., 2011). In the present study, phenolic compounds reacted positively withferric chloride and found as brownish or bluish black in the hypodermal and cortical regions of
the rhizome in both the species.
The histochemical test was possible to demonstrate the sites of production or
accumulation of some metabolites of the plant. The phenolic compounds in epidermal cells
may help the plant in different ways, as against pathogens attacks and even protection against
ultraviolet radiation (Simões et al., 2010). Lipids and fatty acid composition of seagrass might
be responsible for their thermal stability (Goncharova et al., 2000). Studies on histochemical
localization of both the sea grasses indicated that tannins are copiously present in rhizome
and leaf and were localized as dark brown contents in parenchyma cells bordering the vascularbundles. In the sub-hypodermal region of rhizome, flavonoids were located as a distinct yellow
layer. Similar results on histochemical localization are also confirmed in Excocaria agallocha
and in Barleria lupulina (Mandal and Mandal, 2014)
Phytochemical Screening
The study of the phytochemical properties of seagrasses is essential because they werereported to have significant nutraceutical values (Mani et al., 2012). In the present study, thepreliminary phytochemical tests were done in ethanol, ethyl acetate and water extracts of leaf
and C. serrulata. Both ethanol and ethyl acetate extracts confirmed the presence of important
active chemical constituents such as flavonoids, tannin, terpenoids, lignin and phenols in
different parts of both the seagrass that reportedly have antibacterial (Ragupathi Raja Kannan
et al., 2010), antialgal, antifungal, antiviral (Rowley et al., 2002), antiprotozoal (Orhan et al.,2006), anti-inflammatory (Hua et al., 2006), and antidiabetic (Gokce et al., 2008) activities.
Tannins and phenols were dominantly recorded in both leaf and rhizome of C. serrulata
whereas, alkaloids and polyols were traced in leaf only. The results of the phytochemical
analysis are compiled in Table.1. The presence of phenolics and flavonoids in the leaf extractsof C. serrulata and S. isoetifolium extract could be attributed to the defense mechanism of theplant against the epiphytes (Pradheeba et al., 2011). Similarly, coumarin was detected in theethanolic extracts of leaf and rhizome of both the sea grasses but anthraquinone was detected
only in the leaf extract of C. serrulata. These results were supported by the earlier reports ofErgene et al.(2006) and Sangeetha & Asokan (2016) who revealed the presence of tannins,saponins, proteins, resins, reducing sugar, acidic compounds, alkaloids, cardiac glycosides
and terpenoids in the phytochemical analysis of C. rotundata, Cymodocea serrulata, Halophilaovalis and Halodule pinifolia. The present results on the phytochemical screening were similar
to that of Saranya et al. (2017) who reported that Thalassia hemprichii methanolic extract
showed the presence of alkaloids, flavonoids, saponins, phenolics, tannins, steroids andtriterpenoids.
Journal of Global Resources, July 2022, Volume 08 (02) J. D. Jaisankar and A. Ramasubramanian P a g e | 54

Table 02: Phytochemical Localization of Secondary Metabolites in Leaf, Rhizome andRoot of C. serrulata and S. isoetifolium
Phytochemicals

StarchLigninTanninAlkaloids
Polysaccharides
LipidsCutin
Terpinoids
Phenolics
Coumarin
Steroids

Leaf
E
EA
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
-

C. serrulata
Rhizome
E
EA
+
+
+
+
+
+
+
+
+
+
+
+
-

Root
E
EA
+
+
+
+
+
+
+
+
+
+
-

Leaf
E
EA
+
+
+
+
+
+
+
+
+
+
+
-

S. isoetifolium
Rhizome
Root
E
EA
E
EA
+
+
+
+
+
+
+
+
+
+
+
+
-

E – Ethanol extract; EA – Ethyl acetate extract

The qualitative phytochemical screening of C. serrulata was in agreement with theworks of Amudha et al. (2017). Rengasamy et al., (2013) tested six seagrass species such as
Syringodium isoetifolium, Enhalus acoroides, Halodule pinifolia, Thalassia hemprichii,
Cymodocea serrulata, Cymodocea roundata for their antioxidant property and found highcontent of phenolic, tannin, and vitamin E content in S. isoetifolium and high flavonoid levels
in C. serrulata. Subhashini et al. (2013) and Arsianti et al. (2017) reported that phenolic acids
such as p-coumaric acid and gallic acid are predominantly found in Halophila ovalis, Thalassia
hemprichii, Halodule spp., Cymodocea spp., Enhalus acoroides, Syringodium isoetifolium andother seagrass species. The array of phenolic acids produced by seagrasses is dependent
upon the species, population, and tissue examined.
Photosynthetic Pigments
Table 2 indicated that C. serrulata growing in sandy shoreline of Kodiyakarai showedsignificantly higher chlorophyll contents as compared to other sampling sites; the chlorophyll abeing significantly higher than chlorophyll b. The chlorophyll a content was found maximum(0.357 mg g-1) in the leaves of C. serrulata and (0.313 mg g-1) in the leaves of S. osietifolium
during early summer (March) while it was lesser in pre-monsoon (August) (0.331 mg g-1).
Chlorophyll b was found less during all the seasons in the range of 0.218 to 0.292mg g-1. The
magnitude of chlorophyll a/b ratio was found higher during summer and it has very low
carotenoid content (0.076 mg g-1). Bharatharathna and Santhanam (2019) have recorded avery low content of chlorophyll a (6.78µg/g FWT) and chlorophyll b (5.13 µg/g FWT) in S.
isoetifolium from Gulf of Mannar during wet season. The nutrient composition of sea water has
an important influence on the production of photosynthetic pigments compounds by marine
species. The present results are in agreement with earlier reports on other marine angiosperms(Enhalus acoroides, Halophila ovalis, H. beccarii, Cymodocea) which indicated that there is aclear seasonal variation in pigments concentration by registering the minimum value during
the monsoon season when the water column was more turbid and the light availability to theseagrasses was limited causing light stress (Kolsi et al., 2017). In general, the pigments
contents can be affected by several environmental factors largely influenced by the availabilityof the light and morphology of the seagrass leaves and the depth in which the plants are
growing (Pradheepa et al., 2011).
Total Phenol Content
Phenolic compounds present high level in S. isoetifolium which has high antioxidant activity
that scavenges the toxic free radicals and reactive oxygen species such as superoxide radical(O2), hydroxyl radical (OH), peroxide radical (ROO) and nitric oxide (NO) radicals (Girija et al.,Journal of Global Resources, July 2022, Volume 08 (02) J. D. Jaisankar and A. Ramasubramanian P a g e | 55

2013). Seasonal changes had significant effects on the concentrations of total phenol in C.
serrulata than S. isofolium, being higher during the dry season than in wet season. Likewise,significant effect of geography (sampling sites) on the concentration of total phenol was evident
where the highest concentration (4.36 mg g-1 dry wt.) was recorded at Kodiyakarai during
summer. Even though, there was no significant effect in allocation of phenol between the leaf
and rhizome, higher concentrations were frequently observed in leaves with mean values
ranging from 2.96 to 4.36 mg/g-1 dwt in C. serrulata, and from 2.56 to 4.27 in S. isoefolium.
Table 03: Photosynthetic Pigment Contents and Total Phenol Content (TPC) of C.
serrulata and S. isoetifolium
Sampling
SW
Monsoon(June-Sep)
NE Monsoon(Oct-Dec)
Winter(Jan-Feb)
Summer(Mar-May)

C. serrulata

S. isoetifolium

Chlorophyll a (mg g-1)

Chlorophyllb (mg g-1)

Carotenoid (mg g-1)

TPC
(mg g-1)

Chlorophylla (mg g-1)

Chlorophyllb (mg g-1)

Carotenoid (mg g-1)

TPC
(mg g-1)

0.337 ±
0.04

0.218 ±
0.09

0.062 ±
0.12

4.17 ±
0.03

0.257 ±
0.04

0.208 ±
0.09

0.058 ±
0.12

4.03 ±
0.06

0.326 ±
0.07
0.341 ±
0.06
0.357 ±
0.11

0.232 ±
0.11
0.257 ±
0.07
0.292 ±
0.13

0.058 ±
0.11
0.052 ±
0.09
0.076 ±
0.05

2.96 ±
0.11
3.11 ±
0.09
4.36 ±
0.07

0.286 ±
0.09
0.301 ±
0.05
0.317 ±
0.12

0.227 ±
0.12
0.277 ±
0.03
0.283 ±
0.13

0.052 ±
0.13
0.053 ±
0.11
0.066 ±
0.07

2.56 ±
0.13
2.91 ±
0.05
4.27 ±
0.11

Values are Mean ± SE

The presence of phytoconstituents such as phenols, flavonoids, tannin, coumarin in C.
serrulata and S. isoetifolium indicated a possible role that its extracts have antioxidant activity.
Similarly, some earlier reports reveal that seagrasses especially their polyphenols have theantioxidant activity (Gokce et al., 2008; Ragupathi Raja Kannan et al., 2010). Athiperumalsami
et al. (2010) have reported highest antioxidant activity in the methanolic extract of Halophilaovalis than H. pinifolia tested by the NO scavenging method. Ragupathi Raja Kannan et al.
(2010a) have reported antioxidant activity of 11.77 mg ascorbic acid equivalent/g of the leaf ofEnhalus acoroides collected from Chinapallam, Gulf of Mannar, India.
Figure 02: Antioxidant activity of C. serrulata Leaf Extract

Figure 03: Antioxidant activity of S. isoetifolium leaf extract

Journal of Global Resources, July 2022, Volume 08 (02) J. D. Jaisankar and A. Ramasubramanian P a g e | 56

Antioxidant Activity
More recently, reports have revealed that seagrasses are rich sources of antioxidant
compounds (Gokce et al., 2008; Ragupathi Raja Kannan et al., 2010). The presence ofphytoconstituents such as phenols, flavonoids and tannin in seaweeds and seagrass indicateda possible role that its extracts may have antioxidant activity. The results of the in vitro
antioxidant activities of C. serrulata and S. isofolium are shown in Fig. 2 & 3. In the present
study, the DPPH scavenging ability of ethanolic extract of both the seagrasses wasconcentration dependent and it was comparable to the ascorbic acid. DPPH radical was
scavenged by antioxidant through donation of hydrogen to form a stable DPPH molecule. The
concentration of antioxidant needed to decrease the initial DPPH concentration by 50 percent(IC50) is a parameter widely used to measure antioxidant activity. As the IC50 value of theextract decreases, the free radical scavenging activity increases. In the present study, theinvestigated leaf extract of C. serrulata expressed the ability to scavenge the stable DPPH free
radical reaching 50 percent of reduction with an IC50 value 115.7µg/ml conformed to vit. C.
Similarly, S. isoetifolium showed a maximum antioxidant activity of 86.4 percent with
IC50 value of 122.2 µg/ml. According to Bharathi et al. (2016) Syringodium isoetifolium showsa wide range of pharmacological activities such as antibiotic, antihemolytic, cytotoxicity,
antibacterial, and antifungal activity. The present findings are in agreement with thosedescribed for other extracts from five species of seagrasses and six species of seaweeds
which were collected from the Gulf of Mannar, on the southeastern coast of India(Athiperumalsami et al., 2010). In contrast to our findings, Jayaprakash et al. (2016) reported
a high radical scavenging activity of ethyl acetate extract in S. isoetifolium, C. rotundata andH. uninervis
Molecular Characterization of Phytochemicals
The chemical nature and functional groups present in the dried leaf powder of C. serrulata and
S. isoetifolium was predicted using FTIR. The bonds were determined by interpreting theinfrared absorption spectra (Fig 4). Strong bonds were found at 3432 cm−1, 1631.92 cm−1
and 1419 cm−1, while the others varied from weak to medium. These results demonstrated
the presence of hydroxyl groups, lipids, alkanes, amino acids and phosphorus compounds.
The sharp absorption peak at 1631 – 1736 cm-1 are assigned to C=O stretching vibration incarbonyl compounds which may be characterized by the presence of high content of flavanoids
in the leaf of C. serrulata. The FTIR spectral analysis confirmed the presence of phenolic
compounds in the aqueous methanolic extracts of both the seagrasses investigated in thepresent study. The FTIR identified that the leaf extract contained strong C-H out-of-planebending (oop bend) vibration for substituted benzene ring, indicating the presence of phenols
and flavonoids in the crude S. isoetifolium extract also. The wavelength numbers of FTIR
spectra for tannic acid at 669, 855, 1163, 1518 were observed in both the seagrasses.
The peak around 1734–1745 cm−1 was assigned as C=O ester, which may be related
to chlorophyll (Li et al., 2018). The FTIR spectrum at 1654 cm−1 was assigned as the C=O
conjugate. The conjugated double bond in carotenoids has been reported as the structure
responsible for light absorption (Mezzomo and Ferreira, 2016). A previous study stated that apeak at 1654 cm−1 is due to chlorophyll and protein content (Kushwaha et al., 2014). Basedon the phytochemical compounds discovered in the FTIR analysis, C. serrulata has thepotential to be used for biomedicine applications, as phenolic compounds, tannins, flavonoids,
chlorophyll and carotenoids are known to have antioxidant activities (Lachowicz et al., 2018).
A strong broad absorption band around 3422 cm-1 was found in C. serrulata and 3439 cm-1
in S. isoetifolium may be due to the presence of hydrogen bond N-H stretching, characteristicof amino acids. The absorption band at 2921 cm-1 and 2920 cm-1 are corresponding to C-Hstretching of the CH2 groups respectively, indicates the presence of various amino acids, these
Journal of Global Resources, July 2022, Volume 08 (02) J. D. Jaisankar and A. Ramasubramanian P a g e | 57

bands may also be characteristic for the presence of aliphatic CH groups in these compounds.
Bands at 1518 cm-1 and 1523 cm-1 indicates the presence of benzene ring in aromaticcompounds. The absorption band at 1031 cm-1 and 619 cm-1 respectively are due to thestretching vibration of C-O group of esters and phenols (Valchos et al., 2006).
Figure 04: FTIR Spectral Analysis of Methanol Leaf extract of (a) C. serrulata (b) S. isoetifolium

The hydroxyl groups both alcohol and carboxylic acids are observed at the 3392 cm-1
stretching band. Aromatic C-H stretching is identified by the following stretching bands 2921,
1031, and 1110. The vinyl carbons are identified by the stretching band around 1631 cm-1.
The aromatic C=C are assigned by the band around 1439, 1518, 1523 cm-1. The C-O
stretching or C-OH are assigned by the band at 1245 cm-1 was observed in C. serrulata and1253 cm-1 in S. isoetifolium. The methanolic extracts showed all the above bands which areevidence for the presence of p-coumaric acid in the seagrasses (Pharmawati and Wrasiati,2020). The identification of compounds via FTIR spectrophotometry supported the findings
from the phytochemical screening, which detected the presence of phenols and flavonoids.
The amines, imines, alkanes and phenols present were considered the major functional groupsof bioactive compounds (Pradeepa et al., 2015).
Conclusion
Seagrasses can be assertively recognized as one of the important coastal ecosystems and it
requires greater attention for its conservation, monitoring and management. The results of thepresent study may increase the search on novel compounds from seagrass, and they havehigh contents of antioxidants like polyphenols, terpenoids, flavonoid, tannins, and coumarin.
Generally, the trend observed in this study indicated that accumulation of phytochemicals
would differ depending on the plant organ, season, and the conditions of the environment
where they grow. Further work on the purification of individual groups of bioactive components
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may reveal the exact potential of the seagrass to develop a novel broad spectrum herbal
antimicrobial formulation in the future. Further purification of active compounds and structural
elucidation can be used for drug discovery.
References
1. Amudha, P., Vanitha, V., Bharathi, N. P., Jayalakshmi, M., Mohana Sundaram, S. (2017).
Phytochemical analysis and invitro antioxidant screening of sea grass-Enhalusacoroides.
International Journal of Research in Pharmaceutical Sciences, 8(2):251– 258.
2. Ansari, A. A., Ghanem, S. M. (2019). Growth attributes and biochemical composition of Padinapavonica (L.) from the Red Sea, in response to seasonal alterations of Tabuk, Saudi Arabia. The
Egyptian Journal of Aquatic Research, 45(2):139–144.
3. Arellano-Méndez LU, Herrera-Silveira JA, Montero-Muñoz JL, Liceaga-Correa M de los Angeles.
(2011). Morphometric trait variation in Thalassiatestudinum (Banks ex Konig) associated toenvironmental heterogeneity in a Subtropical Ecosystem. J Ecosys Ecograph. DOI: 10.4172/21577625.S1-001
4. Arnon, D.L. (1949). A copper enzyme is isolated chloroplast polyphenol oxidase in Beta vulgaris.
Plant Physiol., 24: 1-15.
5. Arsianti A, Astuty H, Fadilah F, Bahtiar A, Tanimoto H, Kakiuchi K. (2017). Design and screening ofgallic acid derivatives as inhibitors of malarial dihydrofolate reductase by in silico docking. Asian J
Pharm Clin Res;10(2):330-4.
6. Aryakrishna U. K, Vinayan R and Sudha Bai R., (20160. Phytochemical profiling and histochemical
localization in leaf and stem of Trichosanthes cucumerina (L) var. cucumerina with reference to the
influence of plant age and geographical variations, Journal of Pharmacy Research 10(10):660-665.
7. Athiperumalsami, T., Devi Rajeswari, V., Hastha Poorna, S., Kumar, V., & Louis Jesudass, L.
(2010). Antioxidant activity of seagrasses and seaweeds. Botanica Marina, 53, 251–257.
8. Balestri E, de Battisti D, Vallerini F, Lardicci C. (2015). First evidence of root morphological andarchitectural variations in young Posidonia oceanica plants colonizing different substrate typologies.
Estuar Coast Shelf Sci 154: 205-213.
9. Bharatharathna P., and P. Santhanam, (2019). Analysis of phytochemical, biochemical, pigments
and antioxidant activity of seagrass Syringodium isoetifolium.
10. Bharathi, N., Amudha, P. and Vanitha, V. (2016). Seagrass-novel marine nutraceuticals. Int. J.
Pharm. Biosci., 7(4): 86-94.
11. Bukatsch (1972). Bemerkungenzurdoppelfärbung Astrablau-Safranin. Mikrokosmos 61: 255.
12. Bruneton J, Pharmacognosie; phytochimie, plantes médicinales. 4ème édition Lavoisier, Paris,
2009.
13. Chung, K.T., Wong, T.Y., Huang, Y. W., Lin, Y., (1998). Tannins and human health: a review. Crit.
Rev. Food Sci. Nutr. 38, 421–464.
14. Coles R, Grech A, Rahseed M, McKenzie L, Unsworth R, Short F. (2011). Seagrass ecology andthreats in the tropical Indo-Pacific Bioregion. In: Pirog RS (ed) Seagrass: Ecology, Uses andThreats. Nova Scotia Publisher, Inc. New York.
15. Cook, N.C. and Samman, S. (1996). Flavonoids-Chemistry, Metabolism, Cardioprotective Effects,
and Dietary Sources. The Journal of Nutritional Biochemistry, 7, 66-76.
16. de los. Mar Ecol Prog Ser 551:81–94Santos CB, Onoda Y, Vergara JJ, Pérez-Lloréns JL, Bouma
TJ, La Nafie YA, Cambridge ML, Brun F.G, (2016). A comprehensive analysis of mechanical andmorphological traits in temperate and tropical seagrass species
17. Duarte CM and Gattuso, J.P., (2010). Seagrass meadows. In: Encyclopedia of earth. Eds. Cutler J.
Cleveland (Washington, D.C.: Environmental Information Coalition, National Council for Scienceand the Environment). http://www.eoearth.org/article/Seagrass_meadows.
18. Ergene, A., P. Guler, S. Tans, S. Miric, E. Hamzaoglu and A. Duran, (2006). Antibacterial andantifungal activity of Heracleum sphondylium subsp. artvinense, African J. Biotech., 5: 1087-1089.
19. Foster, A.S., (1949). Practical Plant Anatomy, 2nd ed. D. Van Nostrand, Princeton.
20. Girija, K., Parthiban, C., Hemalatha, A., Saranya, C., Anantharaman, P. 2013. Evaluation ofantioxidant activities and preliminary phytochemical analysis of seagrasses Halodule pinifolia,
Halophila ovalis and Syringodium isoetifolium. The J. Phytochem, 114:181–187.
21. Gokce G, Haznedaroglu MZ. (2008). Evaluation of antidiabetic, antioxidant and Vaso protective
effects of Posidonia oceanic extract. J Ethnopharmacol, 115, 122–130.
22. Gokulakrishnan R and Sundaram Ravikumar, (2016). Assessment of seagrass biomass and coastal
land forms along Palk Strait. Indian J of Geo-marine sciences, 45 (8): 1035 – 41.
23. Goncharova SN, Sanina NM, Kostetsky EY. (2000). Role of lipids in molecular thermoadaptation
mechanisms of seagrass Zostera marina. Biochem Soc Trans, 28(6):887-90.
24. Govindasamy, M. Arulpriya, K. Anantharaj, P. Ruban and R. Srinivasan (2013). Seasonal variations
in seagrass biomass and productivity in Palk Bay, Bay of Bengal, India International Journal of
Biodiversity and Conservation vol 5(7):408-417
Journal of Global Resources, July 2022, Volume 08 (02) J. D. Jaisankar and A. Ramasubramanian P a g e | 59

25. Green E.P, Short F.T, (2003). World Atlas of Seagrasses. UNEP World Conservation Monitoring
Centre. University of California Press, Berkeley, USA.
26. Harborne AJ. (1998). Phytochemical methods a guide to modern techniques of plant analysis.
Springer Science and Business Media.
27. Hua K.F, Hsu H.Y, Su Y.C, Lin I.F, Yang S.S, Chen Y.M, Chao L.K. (2006). Study on theantiinflammatory activity of methanol extract from seagrass Zostera japonica. J Agric Food Chem,
54, 306–311.
28. Jeevitha, M., Athiperumalsami, T., Kumar, V. (2013). Dietaryibre, mineral, vitamin, amino acid andfatty acid content of seagrasses from Tuticorin Bay, Southeast coast of India. Phytochemistry.
29. Jeyapragrash, D., Subhashini, P., Abirami, K. and Thangaradjou, T. (2016). Evaluation of in-vitro
antioxidant activity of seagrass: Signal for potential alternate source. Free Rad. Antiox., 6(1): 77-89.
30. Johansen 1940 Plant micro technique. New York: McGraw-Hill, 523 p.
31. Kannan L and Thangaradjou T, (2006). Identification and assessment of biomass and productivityof seagrasses, In: National Training Workshop on marine and Coastal Biodiversity Assessment forConservation and Sustainable Utilization, SDMRI Publication, 10; 9-15.
32. Kendrick G.A, Waycott M, Carruthers T.J.B, Cambridge ML, Hovey RK, Krauss SL, Lavery PS, Les
DH, Lowe RJ, Mascaró O.V, (2012). The central role of dispersal in the maintenance andpersistence of seagrass populations. Bioscience 62:56–65
33. Kiswara W, Behnke N., van Avesaath P, Huiskes A.H.L, Erftenmeijer P.L.A, Bouma T.J. (2009).
Root architecture of six tropical seagrass species, growing in three contrasting habitats inIndonesian waters. Aquat Bot 90: 235-245.
34. Kolsi, R. B., Hichem Ben Salah, Saber Abdelkader Saidi, Noureddine Allouche, Hafedh Belghith
and Karima Belghith, (2017). Evaluation of nutritional value, characteristics, functional properties of
Cymodocea nodosa and its benefits on health diseases, Lipids in Health and Disease 16:238
35. Kuo J., M. L. Cambridge, and H. Kirkman, (2018). Anatomy and Structure of Australian Seagrasses© Springer International Publishing AG, part of Springer Nature, A. W. D. Larkum et al. (eds.),
Seagrasses of Australia, 4: 93 – 125 https://doi.org/10.1007/978-3-319-71354-0_4
36. Kushwaha, K., Saxena, J., Tripathi, B. K. and Agarwal, M. K. (2014). Detection of carotenoids inpsychrotrophic bacteria by spectroscopic approach. Journal of BioScience and Biotechnology, 3(3):253
37. Lachowicz, S., Oszmiański, J. and Wiśniewski, R. (2018). Determination of triterpenoids,
carotenoids, chlorophylls, and antioxidant capacity in Allium ursinumL. at different times of
harvesting and anatomical parts. European Food Research Technology, 244(7): 1269-1280.
38. Li, X., Zhou, R., Xu, K., Xu, J., Jin, J., Fang, H. and He, Y. (2018). Rapid determination of chlorophyll
and pheophytin in green tea using Fourier transform infrared spectroscopy. Molecules, 23: 1010.
39. Lin, Jin-Yuarn & Tang, Ching-Yin. (2007). Determination of total phenolic and flavonoid contents inselected fruits and vegetables, as well as their stimulatory effects on mouse splenocyte proliferation.
Food Chemistry. 101. 140-147. 10.1016/j.foodchem.2006.01.014.
40. Mani AE, Bharathi V, Patterson J, Devadason S. (2012). Antibacterial activity and preliminary
phytochemical analysis of seagrass Cymodocearotundata. Int J Microbiol Res (2):99-103.
41. Margolena, L. A. (1932). Feulgen's and some of its applications for botanical mate rial. Stain.
Tech.,7:9-16.
42. Mbaga WM and Esther F Mvungi, (2019). Effects of Seasons and Zonation on Biochemical Contentsof Two Seagrass Species along Dar es Salaam Coast, Tanzania Journal of Science 45(3): 490-501
43. Mensor L.L., Menezes F.S., Leitao G.G., Reis AS, Santos T.S. and Coube C.S. (2001). Screeningof Brazilian plant extracts for antioxidant activity by the use of DPPH free radical method.
Phytotherapy Research, 15, 127– 130.
44. Mezzomo, N. and Ferreira, S. R. S. (2016). Carotenoids functionality, sources, and processing by
supercritical technology: A review. Journal of Chemistry, 2016: 1-16.
45. Moreira dl, Teixeira ss, Monteiro Mhd, Oliveira Acax and Paumgartten Fjr. (2014). Traditional use
and safety of herbal medicines. Rev Bras Farmacogn 24: 248-257.
46. Nordlund, L.M., Koch, E.W., Barbier, E.B., Creed, J.C., (2016). Seagrass ecosystem services andtheir variability across genera and geographical regions. PLoS One 2016. http://dx.doi.org/10.1371/journal.pone.0163091.
47. O’Brien, T. , N. Feder , and E. Mcgully, (1964). Polychromatic staining of cell walls by Toluidine blue
O. Protoplasma 59 : 368 – 373.
48. Ooi, J.L.S., G.A. Kendrick, K.P. Van Viel and Y.A. Affendi. (2011). Knowledge gaps in tropicalSoutheast Asian seagrass systems. Estuar. Coast. Shelf. Sci. 92: 118–131.
49. Orhan I, Sener B, Atici T, Brun R, Perozzo R, Tasdemir D. (2006). Turkish freshwater and marine
macrophytes extracts show in vitro antiprotozoal activity and inhibit Fabl, a key enzyme of
Plasmodium falciparum fatty acid biosynthesis. Phytomedicine, 13, 735–739.
50. Pande J, Chanda S. (2017). Phyto-Physico-Pharmacognostic study of few medicinal plants ofGujarat. LAP LAMBERT Academic Publishing GmbH & Co. KG, Heinrich-Bocking-Straße 6-8,
66121 Saarbrucken, Germany.
Journal of Global Resources, July 2022, Volume 08 (02) J. D. Jaisankar and A. Ramasubramanian P a g e | 60

51. Pharmawati and Luh Putu Wrasiati (2020). Phytochemical Screening and FTIR Spectroscopy On
Crude Extract From Enhalus acoroides leaves Malaysian Journal of Analytical Sciences, Vol 24 No
1: 70 - 77
52. Pradheeba M, Dilipan E, Nobi EP, Thangaradjou T, Sivakumar K. (2011). Evaluation of seagrasses
for their nutritional value. Indian Journal of Marine Sciences; 40(1):105.
53. Pratheeba, T., Ragavendran, C. and Natarajan, D. (2015). Larvicidal, pupicidal and adulticidal
potential of Ocimumgratissimumplant leaf extracts against filariasis inducing vector. International
Journal of Mosquito Research, 2 (2): 1-8.
54. Ragupathi Raja Kannan R, Arumugam R, Anantharaman P. (2010). Antibacterial potential of three
seagrasses against human pathogens. Asian Pacific J Trop Med, 11, 890–893.
55. Reeve RM., (1951). Histochemical tests for polyphenols in plant tissues. Stain Technol 26, 91-96
56. Rengasamy, K. R., Arumugam, R., Palanisamy, I., Thangaradjou, T. (2013). In vitro antibacterial,cyto- toxicity and haemolytic activities and phytochemical analysis of seagrasses from the Gulf ofMannar, South India. Food Chemistry, 136(3-4):1484– 1489.
57. Rowley D.C, Hansen MS, Rhodes D, Sotriffer C.A, Ni H, McCammon J.A, Bushman F.D, FenicalW. (2002). Thalassiolins A-C: new marinederived inhibitors of HIV cDNA integrase. Bioorg Med
Chem, 10, 3619–3625.
58. Saranya, S., Padma, V.V. (2017). A review on medicinal plants used in cardioprotective remedies
in traditional medicine. Holistic Healthcare: Possibilities and Challenges, 209.
59. Sass JE. (1951). Botanical microtechnique. Ames: Iowa State College Press, 391 p.
60. Simões Cmo, Schenkel Ep, Gosmann G, Mello Jcp, Mentz La and Petrovick Pr. (2010).
Farmacognosia: da planta ao medicamento. 6th ed., Porto Alegre/Florianópolis: Editora da
Universidade UFRGS / Editora da UFSC, 1104 p.
61. Sofowora A. (1993). Medicinal plants and traditional medicines in Africa. Chischester John Willey &Sons New York.
62. Sorescu, A., Alexandrina Nuta, Rodica-Mariana Ion and Lorena Iancu, (2018). Qualitative analysisof phytochemicals from Sea Buckthorn and Gooseberry. DOI: 10.5772/intechopen.77365
63. Subhashini, P., Dilipan, E., Thangaradjou, T., Papenbrock, J. (2013). Bioactive natural products
from marine angiosperms: abundance and functions. Natural Products and Bioprospecting,3(4):129– 136.
64. Sunita D., Sudhir K.K., & Sastry KV. and Rana S. (2010). Phytochemical Screening of Methanolic
Extract and Antibacterial Activity of Active Principles of Hepatoprotective Herb, Eclipta alba.
Ethnobot. Leaflets.
65. Thangaradjou T, Sridhar R, Senthilkumar, S & Kannan L, (2008). Seagrass resources assessmentin the Mandapam coast of the Gulf of Mannar Biosphere Reserve, India, Appl. Ecol. Environ. Res.,
6;139-146.
66. Upton, R., Graff, A., Jolliffe, G., Länger, R., Williamson, E., (2011). American Herbal
Pharmacopoeia: Botanical Pharmacognosy – Microscopic Characterization of Botanical Medicines.
CRC Press, Boca Raton.
67. Valchos, N., Akopelitia, Y., Psaroudaki, M., Konstantinidou, V., Chatzilazarou, A., Tegou, E. (2006).
Applications of Fourier transforminfrared (FTIR) spectroscopy to edible oils, Anal. Chem. Acta,
573/574, 459–465.
68. Waycott, M., Duarte, C.M., Carruthers, T.J.B., Orth, R.J., Dennison, W.C., Olyarnik, S., Calladine,A., Fourqurean, J.W., Heck, K.L., Hughes, A.R., Kendrick, G.A., Kenworthy, W.J., Short, F.T.,
Williams, S.L., (2009). Accelerating loss of seagrasses across the globe threatens coastal
ecosystems. Proc. Natl. Acad. Sci. 106, 12377–12381.
69. Yves-Alain B, Janat A, Mamyrbekova B, Boua B, Fézan H. Tra BI, (2007). Étude ethnobotanique etscreening phytochimique de Caesalpinia benthamiana (Baill.) Herend. et Zarucchi(Caesalpiniaceae). Sciences & Nature 4:217-225.

Journal of Global Resources, July 2022, Volume 08 (02) J. D. Jaisankar and A. Ramasubramanian P a g e | 61