Growth pattern of Chaetoceros
Chaetoceros CHAN had the greatest cell numbers between day 3 (183×104 cell/mL) and day 6 (192×104 cell/mL) during the culture period (Fig. 1). The growth of Chaetoceros CHAN and Chaetoceros BIM exponentially increased during days 2–3, and Chaetoceros CEMB exponentially increased during days 3–5. At day 6, the numbers of cells of Chaetoceros CEMB and Chaetoceros BIM were 168×104 and 150×104 cell/mL, respectively.
Morphological characterization of Chaetoceros
Chaetoceros from this study were isolated from the Gulf of Thailand: Chaetoceros CEMB and Chaetoceros BIM were isolated from Chonburi province, and Chaetoceros CHAN was isolated from Chanthaburi province. The most abundant phytoplankton in the Central Gulf of Thailand are diatoms and blue-green algae (Kajonwattanakul et al. 2008).
Light microscope images are shown in Figs. 2a–c. The sizes of the three Chaetoceros isolates were small (ca. 5 µM). They were delicate and nearly square or rectangular in girdle view, with the pervalvar axis longer than the apical axis. Traditionally, the identification of diatoms at the species level has been based on morphological features determined with the aid of light microscopy, including the morphology of the colonies, the shape and dimensions of cells, the thickness and direction of setae, the number and shape of chloroplasts and the presence and morphology of resting spores. However, other features that can only be resolved with electron microscopy, such as the fine structure of valves and setae and the location and number of rimoportulae, are now considered important (Sunesen et al. 2008). To minimize possible misidentifications, we used both scanning electron microscopy and light microscopy.
Scanning electron microscopy shows that the cells of Chaetoceros CEMB, CHAN and BIM were usually solitary with flat or slightly convex valves (Figures. 2d–l). The setae are straight and narrow in diameter and arise from the poles of the cells. The surfaces of the frustules or cell walls were smooth and rectangular in girdle view. Chaetoceros CEMB cells were shallow rectangular, whereas those of Chaetoceros CHAN and Chaetoceros BIM were square to rectangular. Setae size (18.37 ± 7.41 µm) and transapical axis (4.66 ± 1.25 µm) were significantly higher in Chaetoceros CEMB than in Chaetoceros CHAN and Chaetoceros BIM (Table 1). Chaetoceros is a centric diatom with lightly silicified frustules. Each frustule possesses four long, thin spines or setae. The setae link the frustules together to form colonies of several cells. Frustules can usually be seen in girdle view. Distinguishing Chaetoceros species is difficult using a light microscope. The form of the chains, the sizes of the apical axis and valve shape are some of the most important morphological characters for recognizing species in this genus (Lee et al. 2014). All Chaetoceros isolates were confirmed to be Chaetoceros based on observation of their morphological features with a scanning electron microscope (Table 2).
Table 1
Morphological characters for differentiating the Chaetoceros isolates in this study
Character
|
Chaetoceros CEMB
|
Chaetoceros CHAN
|
Chaetoceros BIM
|
Cell shape
|
Shallow rectangular
|
Square to rectangular
|
Square to rectangular
|
Setae shape
|
Round
|
Round
|
Round
|
Setae size (µm)
|
11.56–28.93
(18.37 ± 7.41)*
|
10.25–13.58
(12.01 ± 1.50)
|
10.27–11.14
(10.62 ± 0.36)
|
Transapical axis (µm)
|
3.24–6.16
(4.66 ± 1.25)*
|
3.50–4.05
(3.70 ± 0.25)
|
3.27–3.90
(3.53 ± 0.25)
|
Apical axis (µm)
|
4.15–8.26
(6.29 ± 1.50)
|
4.77–6.18
(5.33 ± 0.58)
|
4.42–5.49
(4.90 ± 0.49)
|
* indicates statistically significant differences between isolates (p < 0.05) |
Table 2
Comparison of the morphological features of Chaetoceros CEMB, CHAN and BIM by scanning electron microscopy.
Characteristic
|
Chaetoceros CEMB
|
Chaetoceros CHAN
|
Chaetoceros BIM
|
Cell
|
Thick and stiff cells that are shallow rectangular
|
Thick and stiff cells that are rectangular
|
Thick and stiff cells that are rectangular
|
Valve face
|
Oval
|
Rectangular to oval
|
Rectangular to oval
|
Cell Length
|
4–8 µm
|
4–6 µm
|
4–6 µm
|
Apical axis
|
3–6 µm
|
3–4 µm
|
3–4 µm
|
Setae
|
4 long and thin intercalary setae with rounded terminal parts
|
4 long and thin intercalary setae with slender terminal parts
|
4 long and thin intercalary setae with slender terminal parts
|
Spines
|
Invisible spine
|
Invisible spine
|
Invisible spine
|
Cells of C. debilis are roughly rectangular in girdle view and connected in spiraling chains. The basal part of the setae is distinct, and setae extend outward from the spiral. Valves are flat or slightly convex (although the spines make it appear concave). Apertures are narrowly oval and sometimes slightly constricted in the middle. Their diameter ranges from 8–40 µm; although they are distributed worldwide, they mainly occur in cold waters (Hasle and Syvertsen 1996). Species similar to C. debilis include C. curvisetus, which forms spirals wider in diameter; in addition, the apertures are larger and widely oval in C. curvisetus (Guiry and Guiry 2012; Tas and Hernández-Becerril 2017). The cells of C. gracilis and C. muelleri have an apical axis that ranges from 6–8 and 8–9 µm and transapical axis that ranges from 3–6 and 6–7 µm, respectively (Olenina et al. 2006). C. calcitrans has a cell diameter that ranges from 2–5 µm (Soliman et al. 2010). C. debilis is larger than C. gracilis and C. muelleri. Light and scanning microscope observations showed that Chaetoceros CEMB may be a different species compared with Chaetoceros CHAN and Chaetoceros BIM. Nevertheless, light microscopy is still faster and more reliable for diatom identification in a mixed sample for trained diatomologists.
C. gracilis and C. calcitrans are extensively used as food sources for rearing prawn larvae (Chu 1989; Soliman et al. 2010). C. gracilis is the phytoplankton species most commonly used in bivalve mollusk and fish hatcheries (Helm et al. 2004). Their effectiveness stems in part from their small size and n-3 HUFA content (Barclay and Zeller 1996).
Nucleotide sequences and 18S rDNA phylogeny of Chaetoceros
The 18S rDNA sequences of Chaetoceros were obtained from gene cloning and unidirectional sequencing. Chaetoceros CEMB contained 18S rDNA sequences that were 1794 bp in length, which were similar to those of C. gracilis (e-value = 0.0, identity = 98%). Chaetoceros CHAN had 18S rDNA sequences that were 1788 bp in length, which were similar to those of C. debilis (e-value = 0.0, identity = 99%). Chaetoceros BIM contained 18S rDNA sequences that were 1789 bp in length, which were similar to those of C. debilis (e-value = 0.0, identity = 99%). The 18S rDNA sequences of the three Chaetoceros showed high similarity (Fig. 3).
The BLAST analysis revealed high similarity between the Chaetoceros sequences obtained in our study and Genbank sequences (Table 3). We characterized nuclear 18S rDNA sequences of three Chaetoceros and compared them with available DNA sequences (12 sequences) obtained from GenBank (www.ncbi.nlm.nih.gov). These included complete and partial 18S rDNA sequences. Chaetoceros CHAN and BIM were clustered in the same clade with C. debilis and C. curvisetus, and Chaetoceros CEMB was distinct from the others. This result was consistent with morphological data suggesting that Chaetoceros CEMB contained significantly larger setae and apical axes than Chaetoceros CHAN and BIM. The lack of complete consistency between molecular and morphological identification may stem from morphological shifts that occur between environmental species and cultured ones. Thus, species identification both before and after culture might be required to ensure the accuracy of identification (Kesici et al. 2013).
Table 3
Sequence similarities between 18S rDNA sequences of Chaetoceros obtained in this study and GenBank sequences.
Isolate
|
Morphological identification
|
18S rDNA similarities
|
Accession number
|
Chaetoceros CEMB
|
Chaetoceros sp.
|
Chaetoceros gracilis,
e-value = 0.0, identity = 98%
|
MW513719.1
|
Chaetoceros CHAN
|
Chaetoceros sp.
|
Chaetoceros debilis,
e-value = 0.0, identity = 99%
|
MW513720.1
|
Chaetoceros BIM
|
Chaetoceros sp.
|
Chaetoceros debilis,
e-value = 0.0, identity = 99%
|
MW513721.1
|
Chaetoceros is a diverse genus of marine diatoms. Although the morphology of many members of the genus has been well described, molecular taxonomic studies of Chaetoceros are scarce. 18S and 28S rDNA phylogenies indicate that these sequences might provide suitable markers for resolving the species-level taxonomy of Chaetoceros (Oh et al. 2010).
RAPD profiles of Chaetoceros
Amplified fragments 300–2000 bp in size were obtained using RAPD-PCR analysis with UBC10 and OPB01 primers (Fig. 5). A dendrogram based on the RAPD-PCR band was created. The RAPD dendrogram was consistent with the 18S rDNA phylogeny shown in Fig. 4.
DNA barcoding requires molecular loci that are variable enough to discriminate species and a molecular reference database for comparison. The similarity or divergence of the molecular sequence of an unknown organism to a vouchered reference sequence in the database is used for species identification. DNA barcoding of environmental samples involves the extraction of DNA from a pooled sample, PCR amplification of a target locus, cloning of the resulting PCR products, sequencing and analysis. With DNA barcoding techniques, even morphologically similar strains can be identified at the species level. These molecular phylogenetic analyses also enable the rapid, convenient and accurate classification of diatoms and have thus contributed considerably to studies of diatom diversity.
RAPD-PCR has been used for the molecular characterization and identification of 17 samples of Sargassum spp. (Ho et al. 1995). A 450-bp fragment generated using OPA13 was detected in 12 of 17 samples of Sargassum. This fragment was present in profiles from Turbinaria (Sargassaceae). This study showed that RAPD-PCR is useful for discriminating Sargassum samples and developing fingerprints for them. PCR-RFLP analysis has been used to resolve the species-level differences of 18 isolates of Chaetoceros Ehrenberg (Bacillariophyceae) by targeting the rbcL region of chloroplast DNA, which encodes the Rubisco large subunit (Toyoda et al. 2013). RAPD patterns for the species-level differences of Chaetoceros have not been reported to date.
Molecular identification appears to be relatively effective for diatom identification given the similar efficacies of molecular and morphological identification in this study. However, more work is needed to optimize morphological and molecular approaches for diatom identification.
In the study of the interpopulational variability of the three Chaetoceros culture populations, the selection of the RAPD primers was based on the quantity, intensity and repetition of the amplified fragments. These amplified fragments ranged in size from 50 to 2200 bp. A total of 80 fragments were identified, and 113 of these fragments (42.5%) were polymorphic. The average number of fragments per primer was relatively high. The percentage of polymorphic bands was 33.33%, 60.00% and 30.43% for Chaetoceros CHAN, Chaetoceros CEMB and Chaetoceros BIM, respectively (Table 4).
Table 4
Number of RAPD fragments and polymorphic products obtained in the analysis of three Chaetoceros populations.
Primer
|
Total number of fragments
|
Number of amplified fragments
|
Number of polymorphic fragments
|
|
|
Chaetoceros CHAN
|
Chaetoceros CEMB
|
Chaetoceros BIM
|
Chaetoceros CHAN
|
Chaetoceros CEMB
|
Chaetoceros BIM
|
UBC10
|
39
|
14
|
14
|
11
|
5
|
6
|
2
|
OPB01
|
41
|
13
|
16
|
12
|
4
|
12
|
5
|
Total
|
80
|
27
|
30
|
23
|
9
|
18
|
7
|
Polymorphism
|
|
|
|
|
33.33%
|
60.00%
|
30.43%
|
Identification of metabolites extracted from Chaetoceros by NMR spectroscopy
The 1H-NMR spectra of methanol extract from all Chaetoceros isolates showed similar characteristic chemical shift peaks. There was a total of 27 metabolites that were clearly identified based on comparison with previous research, a free NMR database (The Human Metabolome Database, HMDB) and a commercial NMR database (Bruker AssureNMR). The 1H-NMR spectra shown in Fig. 6 contain different groups of metabolites, including amino acids, sugars, carboxylic acids, fatty acids, vitamins and carotenoids. The peaks corresponding to the structures of each metabolite are summarized in Table 5.
Table 5
The metabolites from Chaetoceros methanol crude extract identified by NMR spectroscopy.
Number
|
Metabolite
|
Functional Group
|
Chemical Shift (ppm)
|
1
|
Glutamic acid
|
-CH2-CH2-COOH
-CH2-CH2COOH
-CH2-COOH
|
2.34 (m)
2.13 (m)
2.06 (m)
|
2
|
Proline
|
-(CH2)2-CH-
-CH2-
-CH2-
-CH2-
|
4.10 (dd)
2.32 (m)
2.05 (m)
1.98 (m)
|
3
|
Alanine
|
(COOH)-CH(CH3)-NH2
|
1.48 (d)
|
4
|
Isoleucine
|
CH3-CH(CH)CH2-
CH3-CH2-
|
0.99 (d)
0.96 (t)
|
5
|
Methionine
|
-CH2-CH(NH2)-
|
2.14 (m)
|
6
|
Choline
|
(CH3)3N-CH2-CH2-OH
|
4.05 (ddd)
|
7
|
Glycine
|
NH2-CH2-COOH
|
3.54 (s)
|
8
|
Cholesterol
|
-CH3
-CH(CH3)2
-CHCH3
-CHCH3
|
0.69 (s)
0.82 (d)
0.87 (d)
0.92 (s)
|
9
|
Palmitic acid
|
-CH2-COOH
-CH2-CH2-CH2-
-CH2-(CH2)14-CH3
|
2.31 (m)
1.66 (m)
1.29 (m)
|
10
|
Oleic acid
|
-CH2-CH = CH-CH2-
-CH2-COOH
-(CH2)n -CH2CH3
-(CH2)n-CH2COOH
-CH2-CH3
|
5.40 (m)
2.30 (t)
1.93 (m)
1.33 (m)
0.88 (t)
|
11
|
Linolenic acid
|
-CH2-CH = CH-CH2-
-CH2-COOH
-(CH2)n-CH2COOH
-CH2-CH3
|
5.38 (m)
2.32 (t)
1.34 (m)
0.90 (t)
|
12
|
α-linolenic acid
|
-CH = CH-
-CH2-
-CH2-COOH
-CH2-
-CH2-
-CH2-CH3
|
5.36 (m)
2.79 (m)
2.34 (t)
2.04 (m)
1.31 (m)
0.97 (t)
|
13
|
Arachidic acid
|
-CH2-COOH
-CH2-CH2COOH
-CH2-CH3
-CH2-CH3
|
2.32 (t)
1.64 (m)
1.29 (m)
0.89 (t)
|
14
|
Glucose
|
-CH-
-CH-
-CH2-
|
5.20 (d)
3.82 (m)
3.54 (d)
|
15
|
Sucrose
|
-CH-
-CH-
-CH2-
-CH2-
-CH-
|
5.39 (d)
4.19 (d)
3.82 (m)
3.67 (s)
3.46 (t)
|
16
|
Myo-inositol
|
-CH-
|
4.06 (t)
|
17
|
Fucoxanthin
|
Olefinic-H
Olefinic-H
Olefinic-H
Olefinic-H
Olefinic-H
Olefinic-H
-CH-OH
-CH2-
-CH2-
-CH3
|
6.81 (t)
6.74 (dd)
6.45 (dd)
6.50 (d)
6.41 (m)
6.10 (m)
3.65 (m)
1.52 (dd)
1.38 (dd)
0.97 (s)
|
18
|
Astaxanthin
|
-CH-
Olefinic-H
-CH2-
-CH2-
-CH3
-CH3
-CH3
-CH3
-CH3
-CH3
|
7.01 (d)
6.20–6.85 (m)
4.36 (dd)
3.66 (s)
2.01 (s)
1.98 (s)
1.93 (s)
1.86 (m)
1.33 (m)
1.26 (s)
|
19
|
Lutein
|
Olefinic-H
Olefinic-H
Olefinic-H
Olefinic-H
-CH- (eq)
-CH3
|
6.67–6.57 (m)
6.38 (d)
6.36(d)
5.39 (m)
2.35 (m)
1.19–1.33 (m)
|
20
|
Zeaxanthin
|
-CH2-
-CH3
-CH3
-CH3
|
2.02 (s)
1.99 (s)
1.68 (s)
1.06 (s)
|
21
|
Violaxanthin
|
-CH2-
-CH3
-CH3
-CH3
|
1.93 (s)
1.86 (s)
1.60 (s)
0.92 (s)
|
22
|
Chlorophyll c1
|
-NH-
-NH-
-NH-
-NH-
|
9.95 (s)
9.86 (s)
9.77 (s)
8.20 (s)
|
23
|
Chlorophyll a.
|
-NH-
|
9.56 (s)
|
24
|
Glutamine
|
-CH2-CONH2
-CH2-
|
2.43 (m)
2.12 (m)
|
25
|
Valine
|
(CH3)2-CH-
-CH3
-CH3
|
2.29 (m)
1.03 (d)
0.98 (d)
|
26
|
Leucine
|
-CH3
-CH3
|
1.68 (m)
0.94 (d)
|
27
|
Stearic acid
|
-CH2
-CH2
-CH3
|
1.73 (t)
1.47 (t)
1.01 (s)
|
The characteristic chemical shifts of eight amino acids and sugars were observed around the region 4.10–1.98 ppm, which correspond to the -CH2- protons of amino acids, and 1.48–0.96 ppm, which correspond to the -CH- and -CH3 protons of amino acids (Azizan et al. 2018; Ma et al. 2019; Iglesias et al. 2019). The peaks around 5.20–3.82 ppm correspond to the -CH- protons of glucose and sucrose, and the peaks around 3.82–3.67 ppm correspond to the -CH2- protons of glucose and sucrose (Richter and Berger 2013). The representative proton signals of fucoxanthin (olefinic-H), astaxanthin, lutein and zeaxanthin were observed at a chemical shift around 7.01–6.10 ppm (Zailanie and Purnomo 2017; Shumilina et al. 2020; Otaka et al. 2016; Iwai et al. 2008). The identifications of these carotenoids have been confirmed by 2D-NMR (HMBC and JRES); the JRES spectrum showed the singlet signals of fucoxanthin and astaxanthin at 2.01 and 1.98 ppm, respectively, which correspond to the methyl groups (Fig. 7) (Subramanian et al. 2015). The correlation between the proton and carbon signals in the HMBC spectrum is consistent with the results of previous studies (Azizan et al. 2018) (Figs. 8–9). The signals of chlorophyll a and chlorophyll c1 observed around 9.77–8.20 ppm correspond to the -NH- protons of chlorophyll structures.
1H-NMR and 2D-NMR spectra of the crude extract revealed signals for six fatty acids, including palmitic acid, oleic acid, linoleic acid, α-linolenic acid, arachidic acid and stearic acid (Figs. 10). The characteristic peaks were similar to the results of previous studies (Roswanda et al. 2017; Otto et al. 2014; Singer et al. 1996). The correlation between proton and carbon signals observed around 1.73–1.29 ppm corresponded to arachidic acid, palmitic acid and stearic acid. Other valuable metabolites, such as myo-inositol, cholesterol and choline, were detected by 1D and 2D NMR spectroscopy. The results indicated that further purification was not required for the identification of some major and minor small metabolites by NMR spectroscopy. Gas chromatography coupled with mass spectrometry (GC − MS) is most widely employed for its separation power, choice of detectors, and the relatively inexpensive cost of the instrumentation. Multiple extraction methods have been developed based on GC − MS techniques according to it allowed the choices of reagent and extraction processes (Zheng et al. 2013). Consequently, NMR spectroscopy is commonly used in metabolomic studies due to its high reproducibility and simple preparation process (Savorani et al. 2013).
Lipids play an important role in larval growth and survival. Eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA) are considered essential fatty acids because they are integral components of the plasma membrane and marine fish larvae cannot synthesize them from linoleic acid 18:3 (n-3). M. rosenbergii also lacks the ability to synthesize linolenic acid and linoleic acid (D’Abramo and Sheen 1993) and has a limited ability to elongate and desaturate short-chain n-3 and n-6 polyunsaturated fatty acids (e.g., C18) to long-chain polyunsaturated fatty acids (e.g., C20) (Reigh and Stickney 1989). Schizochytrium spp. contains lipids in amounts as high as 55% of cell weight, of which DHA 22:6 (n-3), EPA 20:5 (n-3) and arachidonic acid represent 35%, 7% and 5%, respectively (Nakahara et al. 1996; Barclay 1997; FAO 2012).
Thus, marine fish larvae must acquire HUFAs through their diet of zooplankton (e.g., rotifers and crustaceans), which are enriched in these nutrients. Increasing the HUFA content of zooplankton before feeding larval fish and shrimp is a regular practice in the aquaculture industry (Apt and Behrens 1999).