Advances in sampling strategies and analysis of phytoplankton

Chapter 31

Advances in sampling strategies and analysis of phytoplankton Priya M. D’Costa1, Ravidas K. Naik2 1 Department of Microbiology, Goa University, Taleigao Plateau, Goa, India; 2ESSO-National Centre for Polar and Ocean Research, Vasco, Goa, India

31.1 Introduction The word phytoplankton is derived from the Greek words phyton ¼ plant, and plankton ¼ wanderer. Phytoplankton are microscopic, single-celled organisms found in all aquatic habitats. They also account for up to 50% of the global primary production [1], and are the base of food webs in aqueous environments. They also play crucial roles in influencing the climate through dimethylsulfoniopropionate (DMSP) production and influences on cloud formation [2], and formation of harmful algal blooms (HABs). Phytoplankton is composed of both eukaryotic and prokaryotic species. The dominant group in coastal waters is diatoms; other phytoplankton groups are dinoflagellates, coccolithophores, raphidophytes, flagellates, etc. (Fig. 31.1). Diatoms are silica-shelled protists belonging to phylum Bacillariophyta. They are divided into two major orders: order Centrales (centric diatoms having radial symmetry) and order Pennales (pennate diatoms exhibiting bilateral symmetry). Centric diatoms usually inhabit the water column; only some genera grow attached to substrates during their entire life cycle. A few pennate diatoms possess a raphe (long slit along the length of the frustules) that helps in motility and attachment to substrata. Diatoms reproduce asexually by vegetative division and also through sexual means via spores or resting cells. Though the majority of them are photosynthetic, some diatom species also possess a heterotrophic mode of nutrition [3]. Dinoflagellates, another phytoplankton group, are characterized by the presence of cellulose cell walls and two flagella. They possess a range of nutritive strategies (autotrophic, mixotrophic, heterotrophic) and contribute to HABs. They produce resting stages (termed cysts) that help them to survive unfavorable conditions; long-term cysts are reported to provide a seed bank for HABs [4]. Advances in Biological Science Research. https://doi.org/10.1016/B978-0-12-817497-5.00031-8 Copyright © 2019 Elsevier Inc. All rights reserved.

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FIGURE 31.1 Diatoms e (A) Odontella sp., (B) Coscinodiscus sp.; dinoflagellates e (C) Dinophysis caudata, (D) Tripos furca. (Scale bar in the image ¼ 20 mm).

Picophytoplankton, <3 mm in size [5], are the smallest group of phytoplankton. They have a ubiquitous distribution in both fresh- and marine waters [6], with abundance from 104 cells ml1 upwards [7]. They consist of three groups: Synechococcus and Prochlorococcus (both cyanobacteria) and picoeukaryotes, and contribute a significant proportion of the phytoplankton biomass as well as the primary productivity in various ecosystems [8,9].

31.2 Sampling strategies Since phytoplankton encompasses a wide range of organisms, both prokaryotic and eukaryotic, with different characteristics, nutritional strategies, varying size ranges, and different econiches, varied strategies have been developed for sampling and analysis of different phytoplankton groups.

31.2.1 Choice of research vessel The type of cruise vessel used for sampling in coastal waters (the waters above the continental shelf) depends on the depth of the water column and also on the intended number of sampling stations. For near-shore sampling in areas with water depth below 50 m, trawlers of 10e12 m length are used. The sampling duration is restricted to 12 h and, accordingly, few stations are targeted.

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In coastal sampling where the water column depth varies between 100 and 500 m, coastal research vessels (CRVs) are used, with a sampling duration of 1e15 days. Oceanic waters (above the continental slope) are studied for phytoplankton by conducting cruises aboard ocean research vessels (ORVs). The cruise duration may extend up to 60e70 days depending on the endurance of the ship. In addition to this, there is another type of research vessel that is used specifically in polar oceans. They are called polar research vessels (PRVs) and are characterized by their ability to navigate successfully through ice-covered waters. Also known as “icebreakers,” these ships are the mainstays of research in polar environments. Research vessels (CRV, ORV, and PRV) are equipped with basic scientific instruments like conductivity temperature depth/CTD (portable, on CRV), expendable bathy thermograph, expendable conductivity temperature depth; and grabs and corers to sample water and sediment. CTD is mounted on a large round metal frame called a rosette along with water sampling bottles. Different sensorsdfor chlorophyll, photosynthetic active radiation, turbidity, dissolved oxygen, and nitratedare mounted on a vertical panel so as to enable collection of water samples and determination of the associated physicochemical parameters from various desired depths. In this manner, depth-wise profiles can also be discerned. Surface samples are generally collected by using a grab sampler or a box corer, whereas, deep cores are collected mainly by using gravity corer by geological/paleontological studies. A multicorer is an instrument that can simultaneously collect several subsamples without disturbing surface sediments.

31.2.2 Sampling in coastal waters In shallow coastal waters, water samples are collected using either buckets (for surface waters) or water samplers like HYDROBIOS/NISKIN samplers (for near-bottom waters). Sediment samples are collected using either grabs such as the van Veen grab or manually by divers using corers (in shallow waters). In case of rocky, intertidal areas and beaches, surface water, interstitial water and sediment samples are collected for phytoplankton analysis. Collection of surface waters is done with a bucket. Interstitial water (also called pore water) is collected with a syringe after digging a hole in the sand and allowing the interstitial water to accumulate. However, this method is rather laborious. Sediment, in addition to interstitial water, can be collected from three zones, which are demarcated depending on the tidedlow-tide, midtide, and high-tide levels. Surface sediment (0e2 cm), when exposed, is collected in zip-lock bags, using a spatula. Vertical sediment cores can be collected for analysis of depth-wise profiles of benthic phytoplankton, and especially for diatom migration investigations, using PVC cores of a suitable diameter (2.5 cm). This enables collection of sediment from depths ranging

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from surface to up to even 10 cm. Epibiotic phytoplankton are phytoplankton that grow attached to either living hosts (plant/animal) or nonliving substrates (e.g., rocks); these are sampled by transferring the host organism with forceps into polyethylene zip-lock bags. All the samples should be kept in the shade, in cool conditions, and transferred to the laboratory as soon as possible.

31.2.3 Aspects to be considered One crucial aspect that needs to be considered while planning sampling strategies for phytoplankton is tide timings, especially for intertidal sampling. Secondly, information regarding the depth of the water column, chlorophyll concentration and supporting physicochemical parameters (temperature, salinity, pH, nutrients, suspended particulate matter, etc.) must also be collected. Thirdly, if the purpose of the study is to examine the community structure of benthic phytoplankton, microscopic analysis of a defined number of samples, whether fresh or fixed, will be sufficient. In case of fixed samples, the choice of preservative is of paramount importance, since use of incorrect preservatives will lead to the production of artifacts [10], which will interfere with sample analysis. The most preferred and commonly used fixative for microplankton is Lugol’s iodine solution, added to achieve a final concentration ranging from 1% to 10%. Lugol’s iodine has several advantages over that of other preservatives, the main one being that flagellates do not lose their flagella when exposed to Lugol’s iodine [10]. Also, Lugol’s iodine is relatively easy to prepare and is stable for years. However, fixed samples require to be monitored during storage as iodine oxidizes with time.

31.3 Analysis of phytoplankton Since phytoplankton can be studied for different purposes, the analytical methods depend on the aim of the study and are detailed in the following discussion.

31.3.1 Phytoplankton taxonomy This study requires observing minute morphological details of individual phytoplankton species to identify and classify them as per taxonomic nomenclature. Conventional light microscopy has been the mainstay of phytoplankton taxonomy and analysis for several decades. However, since the late 1960s, electron microscopy (scanning electron microscopy, SEM; and transmission electron microscopy, TEM) has proved extremely useful in establishing accurate phytoplankton taxonomy using ultra-structural features such as ornamentation of body scales or architecture of flagellar roots and central nodules.

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SEM has helped substantially in clarifying the taxonomy of phytoplankton. A case in point is that of Skeletonema species. Skeletonema species can be differentiated on the basis of certain morphological peculiarities that can be seen only with SEM. Skeletonema species, which are abundant in the coastal waters around India, were resolved up to species level using SEM [11]. The intricate frustule designs observed in diatoms using SEM led to them being recognized as a source of nanomaterials [12]. TEM has proved useful for the determination of ultrastructural features in phytoplankton, and the detection of intracellular bacteria in Pinnularia [13], a pennate diatom, and in Alexandrium catenella and Protoceratium reticulatum (toxic dinoflagellates) [14].

31.3.2 Analysis of phytoplankton community structure Fixed samples, once transported to the laboratory, are kept undisturbed for settling [15], followed by siphoning of the upper water with a 10 mm meshcovered tube. This is followed by microscopic analysis of aliquots of the concentrated sample, identification based on standard taxonomic keys and quantification. Given that the above procedure is time-consuming, the FlowCAM (Fig. 31.2), an integrated system for detection and analysis of marine

FIGURE 31.2 The FlowCAM instrument used for analysis of phytoplankton.

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FIGURE 31.3 The image library generated during phytoplankton analysis using FlowCAM.

plankton and particles in a continuous fluid flow, has revolutionized phytoplankton analysis. It was originally developed for investigations of organisms and particles in seawater [16]. It combines the selective capabilities of flow cytometry, microscopy, and fluorescence detection. The main strength of the FlowCAM is the availability of individual images of each particle counted in a library (Fig. 31.3), along with information about its optical properties. By acquiring and storing a digital image of each particle detected, postanalysis identification, differentiation and quantification is made possible. Also, since it acquires high-resolution microscopic images at a very rapid rate, typically up to 10,000 images/minute, rapid estimation of plankton species and biovolume is possible, in addition to information about size, shape, fluorescence, and concentration statistics and, that too, in a fraction of the time required by traditional microscopy. Thus, FlowCAM analysis provides almost real-time information about the dynamics of the plankton community being studied.

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Another instrument called the continuous plankton recorder (CPR) has been in use since 1931 for the real-time analysis of phytoplankton in the field [17]. This instrument is towed through the water by a research vessel. In fact, there is a major CPR survey program conducted by the Marine Biology Laboratory, Plymouth, UK. Such surveys have been carried out in various parts of the world’s oceans. However, due to the higher mesh size used (>200 mm), it is preferred for zooplankton analysis [17].

31.3.3 Analysis of benthic diatoms Benthic diatoms (those occurring in sediment) can be studied using various techniques, depending on the aim of the study, as discussed above. It is problematic to discern diatoms microscopically due to the presence of sediment grains, which may hide from view, diatoms attached to sediment grains, on their side away from the light. The extinction-dilution method [18] has been widely employed for study of benthic diatoms (vegetative forms and resting stages). This method involves incubation of the sediment sample in appropriate seawater-based media, after which the diatom growth is monitored microscopically, identified based on standard taxonomic keys, and enumerated. Since it is a culture method, it enables the detection of vegetative cells as well as algal resting stages that tend to be obscured in soil aggregates [19]. The most probable number is then generated based on a statistical table, and then multiplied with the specific gravity of the sediment sample, which is calculated separately. However, this method does not give the exact number of diatom cells but rather enumerates the relative abundance of different taxa. This is because it is based on the presence and/or absence of diatoms and involves a statistical table [18] and probability theory as part of its calculations [19].

31.3.3.1 Modifications of the extinctionedilution method The extinctionedilution method can be modified to suit the needs of the investigator. For example, artificial seawater can also be used as the basal diluent instead of natural seawater, especially in cases where comparison across seasons or study areas is essential. This will avoid differences in seawater composition from affecting the diatom community under study. Another modification is the use of UV to detect autofluorescence of chlorophyll-containing cells. This helps to detect live diatoms/resting stages even if they are obscured in detritus. The extinctionedilution method, modified by the incorporation of penicillin (an antibiotic that is effective mainly against actively growing cells), has been used to detect the influence of bacteria on benthic diatom community structure [20]. It has also been used to study the effect of different classes of

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antibiotics (aminoglycosides e streptomycin, chloramphenicol) on benthic diatom community structure [21].

31.3.4 Analysis of dinoflagellate cysts Dinoflagellate cysts can be analyzed in several ways depending on the requirements of the study. They are reported to have a long life span; they have been successfully germinated from 32 to 63 cm depth in the sediment, corresponding to a time span of 100 years (based on dating techniques) [22e24]. To check the germination potential of dinoflagellate cysts, sediment suspensions are processed to dislodge sediment particles, and then the concentrated cyst fractions are viewed microscopically to differentiate between entire and ruptured cysts. The entire cysts are picked up and transferred to autoclaved seawater in multiwells (with salinity adjusted to an optimum level, usually 32 psu) and then incubated in cool, bright light (12 h:12 h light: dark conditions) for up to several weeks. Intermittent microscopic observations are required to check whether the cysts have germinated. In this regard, it is important to note that cysts have a mandatory dormancy period during which they will not germinate even when provided with favorable conditions. They will germinate subsequently, only when a “germination window” (favorable conditions) opens up. Dinoflagellate cysts can also be studied in recent sediments or dated sections from deeper cores by a paleontological method, involving treatment of sediment samples with hydrochloric and hydrofluoric acid, respectively [25]. Hydrochloric and hydrofluoric acids dissolve calcium carbonate and silica of coccolithophores and diatoms, respectively. Dinoflagellate cysts, which have a sporopollenin layer, are resistant to the action of acids. The sodium polytungstate density gradation method, which separates living dinoflagellate cysts from inorganic particles and organic detritus [26], has also been used to study dinoflagellate cyst assemblages [27]. Cysts recovered in this manner can be used for germination experiments [26]. Identification of dinoflagellate cysts can be done based on two approaches: (1) based on the names of the vegetative dinoflagellate forms, and (2) based on paleontological nomenclature. This again depends on the nature of the investigation.

31.3.5 Study of fouling diatoms/biofilms If fouling diatoms are to be examined, a substrate of choice (glass/fiberglass/ polystyrene/stainless steel) is suspended in the water column or anchored in the soil (for benthic diatoms) at the study area for a suitable incubation period. The duration of the incubation period will vary depending on the season, the potential for macrofouling, etc. In a study on the fouling diatom community structure from a monsoon-influenced tropical estuary in Goa, west coast of India, macrofouling interference in biofilm development was observed after

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4 days of incubation [28]. Thus, these crucial points must be considered while deliberating on the duration of the incubation period for development of natural marine biofilms. Biofilms can also be developed on panels suspended in aquaria in the laboratory containing fresh natural seawater that has been transported from the sampling area [29,30]. However, biofilms developed in this manner will differ from those developed in natural environments, mainly in the degree of predation they face, the nutrient pulses and turbulent conditions that they are exposed to, and so on. Once biofilms are developed, whether in the natural environment or artificially in the laboratory, they can be studied for fouling diatoms either directly, by viewing the substrate microscopically, or by scraping of the biofilm material, fixing it, and then viewing it microscopically to identify and quantify the fouling forms. Brushing and scraping are the conventional means of biofilm removal from solid substrates, especially for fouling diatoms. Since this pivotal step will influence the results of the analysis, it is extremely important to select the right method to dislodge fouling diatoms from the substrate. Patil and Anil [31] compared two methods of biofilm removal (nylon brush and ceramic scraper), with emphasis on diatoms. They reported that the nylon brush showed a higher efficiency in recovering diatoms compared to the ceramic scraper.

31.3.6 Analysis of epibiotic phytoplankton Epibiotic phytoplankton are usually scraped off the surface of the basibiont host, with a nylon brush into a predetermined amount of filtered seawater. This can be done in case of a hard surface like the chitinous exoskeleton of horseshoe crabs [32]. For epiphytic phytoplankton attached to the surface of seaweeds, they are detached by suspending the seaweeds in a known volume of filtered seawater, mixed manually [33] or by incubating on a shaker (250 rpm) for approximately 30 min. In both the cases mentioned above, the scraped material/seaweed suspension is fixed with Lugol’s iodine, concentrated, and enumerated microscopically.

31.3.7 Study of picophytoplankton Picophytoplankton, due to their small size, are difficult to analyze with traditional methods like epifluorescence microscopy. However, flow cytometry, initially developed for use in the biomedical field, has been successfully applied to picophytoplankton studies since the 1980s. In fact, flow cytometry played a key role in the discovery of picoplankton [34]. Basically, it is a technique for making measurements (-metry) on cells (cyto-), using microscopic methods, performed while the cells flow in a stream past an array of optical detectors. It offers information regarding the pigment content (based on

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fluorescing pigmentsdchlorophyll, phycoerythrin, phycocyanin), cell size, scatter properties, and abundance of the major photosynthetic picophytoplankton. It does so at a rapid rate (several thousand cells per minute). It can also be applied for study of larger phytoplankton (100e200 mm and above), with certain modifications incorporated in the instrument [35]. Flow cytometry has been used to study various aspects of picophytoplankton, even in the field. For example, community structure of picophytoplankton by discriminating between phycoerythrin-rich Synechococcus, phycocyanin-rich Synechococcus, Prochlorococcus, and picoeukaryotes. It has also been used to study viral infection of algal cells [36]. Another interesting aspect is that some flow cytometry instruments (sorters) have the ability to physically sort cells into fractions. This has been of immense help in cell cycle studies and discrimination of several morphologically and ultrastructurally variant cells within a single clonal phytoplankton population [37]. For picoplankton analysis, water/biofilm samples (approx. 2 mL) should be collected in cryovials, kept away from sunlight, stored at 4 C without any fixative, and analyzed within 12 h of collection [36]. This can be done using benchtop flow cytometry instruments on board the ship. This is a huge advantage in that samples, in their live state, can be analyzed immediately, and also, cell loss due to the disruptive effects of preservatives is not an issue [38]. If immediate analysis is not possible, samples can be preserved in paraformaldehyde or glutaraldehyde and stored frozen, either at 80 C or in liquid nitrogen, until analysis [39].

31.3.8 Phytoplankton pigment analysis This study requires filtration of water samples through glass fiber filters, which are then stored at low temperature (20 C, 40 C, 80 C or liquid nitrogen) until analysis to estimate phytoplankton biomass (chlorophyll a) or phytoplankton community structure (pigment indices). Chlorophyll a (chl a) can be estimated using different methods including spectrophotometry (samples scanned between 750 and 350 nm) and fluorometry (excitation CS-5-60 and emission CS-2-64). Spectrophotometric analysis of pigment extracts can accurately measure chl a, b, and c (not carotenoids) through the use of trichloromatic equations [40,41]. However, the fluorometric method, with a higher sensitivity (50 times more sensitive for chl a) has been widely used in present-day research. In vivo fluorometry is commonly used to detect phytoplankton chlorophyll fluorescence in seawater [42]. It is semiquantitative; the fluorescence of chlorophyll depends on the species present, the time of day, accessory pigments, and the physiological condition of the cells [43]. In extracted fluorometry, the chlorophyll is extracted into methanol or acetone before measurement [44]. The response is quantitative only if chlorophyll a is the dominant chlorophyll and there are no chlorophyll b, pheophytins, pheophorbides, or chlorophyllides present. Another advantage of fluorometry is

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that some problems of interference from other chlorophylls and degradation products are reduced [45], since the excitation and emission wavelengths are varied to detect chlorophyll a, b, and c and their degradation products separately. These methods are now superseded by pigment discrimination and quantification by high performance liquid chromatography (HPLC) directly from seawater samples. HPLC systems were developed for higher plant pigments in the late 1970s. Subsequently, HPLC systems of increasing complexity were developed for chlorophylls and carotenoids from microalgae and natural phytoplankton populations [46e49]. HPLC profiles provide estimates of the oceanic distribution of specific algal classes, based on diagnostic pigments such as fucoxanthin (specific for diatoms), peridinin (dinoflagellates), zeaxanthin (cyanobacteria), and prasinoxanthin (Prasinophyceae) [50,51]. The linear relationship between diagnostic pigment and total chlorophyll a is used to calculate the percentage contribution of phytoplankton functional groups [52,53]. It is one of the most prolific separation techniques in use and compares well with other techniques used in phytoplankton analysis (Table 31.1). A very relevant UNESCO monograph Phytoplankton Pigments in Oceanography, published in 1997, covered most of the aspects of HPLC analysis of planktonic pigments, reviewing the application of existing methods to oceanography, and proposing new isocratic and gradient HPLC methods [54]. Generally, reversed-phase HPLC systems are employed rather than normal-phase HPLC systems. C8 columns are preferred over C18 columns due to the ability of C8 phases to separate isomeric pairs of pigments with slight differences in polarity, such as mono- and divinyl chlorophylls or lutein and zeaxanthin. Isocratic HPLC can rapidly analyze chlorophylls and derivatives from relatively small volumes of seawater. It is especially sensitive when coupled with fluorescence detection. Its limited resolution, however, usually prevents its use for full pigment analysis. Gradient HPLC is the method of choice for full analysis of chlorophylls and carotenoids in oceanography. The equipment is, however, expensive and the analyst requires more time and technical expertise than for nonchromatographic techniques.

31.3.9 Analysis of viability and photosynthetic parameters of phytoplankton populations Quantification of photosynthetic pigments (mainly chlorophyll) through spectrophotometry, fluorometry, and HPLC does not always accurately reflect the status of the resident phytoplankton community. This is because, firstly, chlorophyll molecules stay intact within a dead cell for up to 2 weeks [55]. Secondly, it is tricky to quantify the number of living cells per volume of water from a biomass index, mainly so when the species and physiological states are

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TABLE 31.1 Comparison of different techniques involved in phytoplankton analysis. S. No.

Property

LM

SEM

FlowCAM

HPLC

1.

Specimen status

Live, preserved

Preserved

Live, preserved

Preserved

2.

Level of preparation of sample

Simple

Expert

Simple

Expert

3.

Magnification extent

Low

High

Low

e

4.

Resolution

Low

High

Low

Resolution in terms of diagnostic pigment indices

5.

Type of output

Colored 2D images

Blackand-white 3D images

Colored 2D images

Chromatogram

6.

Requirement for specific conditions

No vacuum required

Vacuum required

No vacuum required

Solvents required

7.

Cost

Cheap

Expensive

Expensive

Expensive

unknown [56]. Thus, additional techniques that focus on the viability and physiological status of the phytoplankton population are necessary. The viability of phytoplankton cells can be determined using vital fluorescent stains such as fluorescein diacetate (FDA), 2-(4-pyridyl)-5-{[4dimethylaminoethyl-aminocarbomoyl-methoxy]phenyl}oxazole (PDMPO) and 5-chloromethylfluorescein diacetate (CMFDA) [56,57]. FDA and CMFDA are membrane-permeable vital stains, which are acted upon by nonspecific esterases, present inside viable, intact cells. The end result is nonpermeable, green fluorescent products, which indicates viable cells [56]. PDMPO has a different mechanism of action; it gets incorporated along with silica precipitation during frustules synthesis in viable, growing cells. Thus, the intensity of PDMPO fluorescence is directly related to the quantity of silica precipitated [57,58]. Another interesting parameter that can be measured is the photosynthetic efficiency of the sample, indicative of the physiological status of the phytoplankton cells. It analyzes the amount and organization of protein molecules

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around the light-harvesting complex, PSII. It is measured using the fluorescence induction and relaxation (FIRe) technique [59]. The instrument involved is a FIRe fluorometer, fitted with a fiber-optic probe. These measurements are taken from fresh samples, after allowing an adaptation period of 30 min in the darkness. This method is quick and nondisruptive [59].

31.3.10 Toxin analysis Several phytoplankton species are known to produce toxins, especially those phytoplankton that cause HABs. The presence of toxins is a growing global concern, due to its ill effects on human health and also to the national economy. There are several toxinsddomoic acid, saxitoxin, okadaic acid, gymnodimine, nodularin, yessotoxin, brevetoxins, etc.dthat are genus/species specific. The reasons for toxin production by phytoplankton could vary from species to species; it could be a part of the defense mechanisms, metabolism, allelopathy, resource competition, and also a result of nutrient stress. Since the mechanisms involved are complex, one needs to use specific experimental approaches with targeted species. However, the most pressing issue that needs to be addressed is the detection and monitoring of coastal waters for toxins from all aquatic bodies and especially their filter-feeding inhabitants. There are different methods by which toxins can be collected, extracted, and analyzed. Toxins can be extracted from muscle tissue of organisms affected during algal blooms or directly from the cell extract of bloom-forming species. However, these are the possibilities only if there is a bloom, whereas, during nonbloom conditions, any given aquatic area can be surveyed for toxins by using grab sediment samples followed by laboratory cleanup and extraction. Such sampling still remains one of the common approaches of researchers. However, a new technique has been introduced by MacKenzie et al. [60] wherein passive sampling of toxin can be done using solid-phase adsorption toxin tracking (SPATT). The SPATT device is a simple instrument made up of a polyester mesh bag containing activated resins of polystyrenedivinylbenzene, which can adsorb lipophilic toxins dissolved in water. The SPATT bags are deployed and retrieved from the study location to get a timeaveraged toxin concentration. This technique provides a clean sample matrix, which simplifies the subsequent extraction and analysis using enzyme-linked immunosorbent assay or liquid chromatography-tandem mass spectrometry. Marine phytoplankton toxins also have medical and commercial significance; dinoflagellate toxins, particularly, are receiving increasing interest [61]. For example, pectenotoxin produced by Dinophysis species displays cytotoxic activity against various human cancer cells [62]; goniodomin-A, produced by Goniodoma pseudogonyaulax, and gambieric acid, produced by Gambierdiscus toxicus, have antifungal properties; zooxanthella toxins produced by Symbiodinium sp. exhibit potent vasoconstrictive activity. Similarly, the

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marine biotoxin okadaic acid is vital in medical research for understanding several cellular processes [63]. Mass culturing of such potent species on a commercial scale in the future could benefit medical research. However, even though toxins have high medical and commercial importance, their inaccessibility, especially in case of dinoflagellates, is the major hindrance in fully exploring their research potential.

31.4 Primary productivity Primary productivity is the rate of carbon fixation with respect to space and time. There are various methods to estimate the primary production such as oxygen evolution in dark and light incubations, 14C, 13C, 18O, and the fast repetition rate fluorometry (FRRF) method. Comparison of these methods revealed differences of varying magnitude [64,65]. Among these methods, oxygen evolution in dark and light incubation and 14C methods are classical methods that have been in practice for the last 8 decades [66,67]. The comparatively new methods involve the use of 13C, 18O that are stable isotope tracers [68,69]. In the 14C method [70], bicarbonate ion is labeled with a radioactive isotope of carbon. This is done by adding a known quantity of radioactive bicarbonate (HCO-3) to two bottles of marine samples containing phytoplankton. Further, these bottles are incubated in two tanks, one of which is covered with a black film to avoid light and the other is without the film. In the bottle with film, only respiration will take place. In the other bottle, which is exposed to light, both photosynthesis and respiration will occur. After the incubation period is over, the samples from bottles are filtered out. The amount of radioactive carbon taken up per unit time is measured using a scintillation counter and the primary productivity calculated (mg C m3 h1). In the 13C method [68], bicarbonate ion is labeled with natural stable isotope 13C instead of radioactive isotopes, and hence there is no risk factor involved during analysis. In the 18O method [69], samples are enriched with stable isotope 18O as a tracer to measure the gross primary production. This method measures direct gross photosynthetic O2 production compared to the 14C method. The error in gross primary production estimates from this method is less than 2% [69]. In addition to this, active fluorescence and incubation-free techniques such as FRRF [71] measure the instantaneous depth and time-dependent productivity from active chlorophyll fluorescence at spatial (<1 m) and temporal (1 s) resolutions [72]. This method has better signal-to-noise ratio, which makes possible robust measurements even in oligotrophic regions. This method has the potential to measure physiological parameters instantaneously and also to note the rapid changes that occur in productivity [73]. Yet, this method is also fraught with uncertainties. High concentration of colored-dissolved organic

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matter (C-DOM) affects the spectral absorption of the FRRF, resulting in errors in estimation [74].

31.4.1 Estimation of primary productivity using remote sensing Due to the limitation of in situ observations on a regular basis and nonaccessibility of certain study areas, it is difficult to estimate primary productivity on a large scale. These limitations can be overcome by using the remote-sensing approach, wherein satellite-derived chlorophyll can be used to compute the primary productivity of a given area [75]. However, such remotesensing algorithms need to be calibrated; in a majority of the cases, such calibration has been carried out using the 14C method [76]. The limitations of this method include nonavailability of data during cloud coverage and substantial errors in coastal waters due to C-DOM.

31.4.2 Monitoring of HABs using remote sensing Generally, HABs are monitored using a two-pronged approach: (1) Physical survey at bloom locations during the potential bloom period, and (2) monitoring of blooms using remote sensing as a tool. The former approach gives more details about the bloom dynamics and species responsible for the bloom. However, practically, it is not possible to be present at every bloom location or to conduct frequent surveys to capture initiation of bloom, its decline, and consequences. In such cases, remote sensing is useful to monitor a large area just by using remote-sensing images obtained from ocean color satellites at daily intervals. By looking at chlorophyll concentration from images, it is easy to identify bloom locations. However, due to the lack of species-specific algorithms, it is difficult to identify the bloom-causing genera/species. Hence, the combined approach can be very useful to conduct a bloom survey at the targeted location.

31.5 Future perspectives Considering the diversity of phytoplankton parameters and the resulting varied approaches to phytoplankton studies, it is not surprising that several new techniques, individually and in conjunction, are being applied to phytoplankton analysis. Phytoplankton taxonomy has seen several advancements. In recent years, phylogenetic analysis of the 18 and 28S rDNA genes has been increasingly applied to confirm the validity of subgroups that have been defined based on morphological characters. In many instances, new species have been discovered and the number and circumscription of subgroups has been refined, for e.g., Chaetoceros [77,78] and Skeletonema [79]. This approach has been aided by the incorporation of advanced TEM techniques, which allow far higher resolution compared to earlier techniques.

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Phytoplankton diversity at the genomic level has been studied using various metagenomic techniques, which can be divided into two main categories of analysis based on (1) probes and (2) sequencing [80]. In the first category, probes specific for particular phytoplankton genes are designed and fixed onto a suitable array. Subsequently, samples are analyzed using these array-based probes. This technique allows the analysis of expression patterns of particular genes in a mixed phytoplankton community. Several array-based metagenomic techniques have been devised and employed for marine picophytoplankton communities [81,82]. The second metagenomic approach, based on sequencing, has been successfully used to determine phytoplankton diversity [80]. This approach is advantageous since it offers a comparatively unbiased determination of phytoplankton diversity and has, therefore, led to the discovery of several unknown phytoplankton lineages. DNA metabarcoding is another metagenomic technique that has been applied to unravel diversity within morphologically similar phytoplankton species [83]. It involves combining high-throughput sequencing approaches with sequencing of environmental DNA that has been PCR amplified using primers specific for particular phytoplankton groups. This metagenomic method not only enables the determination of spatial and seasonal distribution patterns without cultivation but also allows the discovery of new taxa linked with new sequences, not assigned to a known organism. Additionally, the use of next-generation sequencing methods has enhanced the number of sequences recovered from individual samples, and overcome the bias against rare taxa [83]. In the context of primary productivity, it is relevant to arrive at a global carbon budget with less error. Thus, it is imperative to set a common global protocol to estimate primary productivity. This can be achieved either by following a common method worldwide or by applying a suitable correction factor to nullify the bias that occurs between different methods [84].

Acknowledgments PD acknowledges the head of the Department of Microbiology, Goa University, for providing necessary facilities. RN is grateful to the director of NCPOR, MoES, for his encouragement. This is NCPOR contribution no. 59/2018.

References [1] Falkowski P. The power of plankton. Nature 2012;483(Suppl. 7387):S17. [2] Laroche D, Vezina AF, Levasseur M, Gosselin M, Stefels J, Keller MD, et al. DMSP synthesis and exudation in phytoplankton: a modeling approach. Mar Ecol Prog Ser 1999;180:37e49. [3] Admiraal W, Peletier H. Influence of organic compounds and light limitation on the growth rate of estuarine benthic diatoms. Br Phycol J 1979;14(3):197e206.

Advances in sampling strategies and analysis of phytoplankton Chapter | 31 [4] [5]

[6] [7] [8] [9] [10]

[11] [12] [13]

[14] [15] [16] [17]

[18] [19]

[20] [21] [22]

[23]

517

Smayda TJ, Trainer VL. Dinoflagellate blooms in upwelling systems: seeding, variability, and contrasts with diatom bloom behaviour. Prog Oceanogr 2010;85(1e2):92e107. Sieburth JM, Smetacek V, Lenz J. Pelagic ecosystem structure: heterotrophic compartments of the plankton and their relationship to plankton size fractions 1. Limnol Oceanogr 1978;23(6):1256e63. Stockner JG, Antia NJ. Algal picoplankton from marine and freshwater ecosystems: a multidisciplinary perspective. Can J Fisheries Aquat Sci 1986;43(12):2472e503. Contant J, Pick FR. Picophytoplankton during the ice-free season in five temperate-zone rivers. J Plankton Res 2013;35(3):553e65. Paerl HW. Ultraphytoplankton biomass and production in some New Zealand lakes. NZ J Mar Freshw Res 1977;11:297e305. Platt T, Rao DS, Irwin B. Photosynthesis of picoplankton in the oligotrophic ocean. Nature 1983;301:702e4. Naik RK, Chitari RR, Anil AC. Karlodinium veneficum in India: effect of fixatives on morphology and allelopathy in relation to Skeletonema costatum. Curr Sci 2010;25:1112e6. Naik RK, Sarno D, Kooistra WHCF, D’Costa PM, Anil AC. Skeletonema (Bacillariophyceae) in Indian waters: a reappraisal. Indian J Geo-Mar Sci 2010b;39(2):290e3. Mishra M, Arukha AP, Bashir T, Yadav D, Prasad GB. All new faces of diatoms: potential source of nanomaterials and beyond. Front Microbiol 2017;8:1239. Schmid AM. Endobacteria in the diatom Pinnularia (Bacillariophyceae). I.“Scattered ctnucleoids” explained: DAPIeDNA complexes stem from exoplastidial bacteria boring into the chloroplasts 1, 2. J Phycol 2003;39(1):122e38. Co´rdova JL, Escudero C, Bustamante J. Bloom inside the bloom: intracellular bacteria multiplication within toxic dinoflagellates. Rev Biol Mar Oceanogr 2003;38(2):57e67. Hasle GR. Settling, the inverted microscope method. In: Sournia A, editor. Phytoplankton manual. Paris: Museum National d’Histoire Naturelle; 1978. p. 88e96. Sieracki CK, Sieracki ME, Yentsch CS. An imaging-in-flow system for automated analysis of marine microplankton. Mar Ecol Prog Ser 1998;168:285e96. Reid PC, Colebrook JM, Matthews JBL, Aiken J, et al. The continuous plankton recorder: concepts and history, from plankton indicator to undulating recorders. Prog Oceanogr 2003;58(2e4):117e75. Throndsen J. The dilution-culture method. In: Sournia A, editor. Phytoplankton manual. vol. 6. Paris: UNESCO Monographs on oceanographic methodology; 1978. p. 218e24. Harris AS, Jones KJ, Lewis J. An assessment of the accuracy and reproducibility of the most probable number (MPN) technique in estimating numbers of nutrient stressed diatoms in sediment samples. J Exp Mar Biol Ecol 1998;231(1):21e30. D’Costa PM, Anil AC. The effect of bacteria on diatom community structureethe ‘antibiotics’ approach. Res Microbiol 2011;162(3):292e301. D’Costa PM, Anil AC. The effect of antibiotics on diatom communities. Curr Sci 2012;102(11):1552e9. Lundholm N, Ribeiro S, Andersen TJ, Koch T, Godhe A, Ekelund F, et al. Buried alivee germination of up to a century-old marine protist resting stages. Phycologia 2011;50(6):629e40. Miyazono A, Nagai S, Kudo I, Tanizawa K. Viability of Alexandrium tamarense cysts in the sediment of Funka Bay, Hokkaido, Japan: over a hundred year survival times for cysts. Harmful Algae 2012;16:81e8.

518 Advances in Biological Science Research [24] Feifel KM, Fletcher SJ, Watson LR, Moore SK, Lessard EJ. Alexandrium and Scrippsiella cyst viability and cytoplasmic fullness in a 60-cm sediment core from Sequim Bay, WA. Harmful Algae 2015;47:56e65. [25] Matsuoka K, Fukuyo Y. Technical guide for modern dinoflagellate cyst study. Japan: WESTPAC-HAB/WESTPAC/IOC, Japan Society for the Promotion of Science; 2000. p. 1e29. [26] Bolch CJ. The use of sodium polytungstate for the separation and concentration of living dinoflagellate cysts from marine sediments. Phycologia 1997;36(6):472e8. [27] Narale DD, Anil AC. Spatial distribution of dinoflagellates from the tropical coastal waters of the South Andaman, India: implications for coastal pollution monitoring. Mar Pollut Bull 2017;115(1e2):498e506. [28] Mitbavkar S, Anil AC. Seasonal variations in the fouling diatom community structure from a monsoon influenced tropical estuary. Biofouling 2008;24(6):415e26. [29] Mitbavkar S, Raghu C, Rajaneesh KM, Pavan D. Picophytoplankton community from tropical marine biofilms. J Exp Mar Biol Ecol 2012;426:88e96. [30] Khandeparker L, D’Costa PM, Anil AC, Sawant SS. Interactions of bacteria with diatoms: influence on natural marine biofilms. Mar Ecol 2014;35(2):233e48. [31] Patil JS, Anil AC. Quantification of diatoms in biofilms: standardisation of methods. Biofouling 2005;21(3e4):181e8. [32] Patil JS, Anil AC. Epibiotic community of the horseshoe crab Tachypleus gigas. Mar Biol 2000;136(4):699e713. [33] Al-Harbi SM. Epiphytic microalgal dynamics and species composition on brown seaweeds (phaeophyceae) on the northern Coast of Jeddah, Saudi Arabia. J Oceanogr Mar Res 2017;5:153. https://doi.org/10.4172/2572-3103.1000153. [34] Chisholm SW, Olson RJ, Zettler ER, Goericke R, Waterbury JB, Welschmeyer NA. A novel free-living prochlorophyte abundant in the oceanic euphotic zone. Nature 1988;334(6180):340. [35] Dubelaar GB, Groenewegen AC, Stokdijk W, van Den Engh GJ, Visser JW. Optical plankton analyser: a flow cytometer for plankton analysis, II: specifications. Cytometry. The J Int Soc Anal Cytol 1989;10(5):529e39. [36] Marie D, Simon N, Vaulot D. Phytoplankton cell counting by flow cytometry. Algal culturing techniques, vol. 1. Academic Press; 2005. p. 253e67. [37] Arora M, Anil AC, Burgess K, Delany J, Mesbahi E. Asymmetric cell division and its role in cell fate determination in the green alga Tetraselmis indica. J Biosci (Bangalore) 2015;40(5):921e7. [38] Vaulot D, Courties C, Partensky F. A simple method to preserve oceanic phytoplankton for flow cytometric analyses. Cytometry 1989;10:629e35. [39] Rajaneesh KM, Mitbavkar S. Factors controlling the temporal and spatial variations in Synechococcus abundance in a monsoonal estuary. Mar Environ Res 2013;92:133e43. [40] Jeffrey ST, Humphrey GF. New spectrophotometric equations for determining chlorophylls a, b, c1 and c2 in higher plants, algae and natural phytoplankton. Biochem Physiol Pflanz (BPP) 1975;167(2):191e4. [41] Lorenzen CJ, Jeffrey SW. Determination of chlorophyll in seawater. UNESCO Tech Pap Mar Sci 1980;35(1):1e20. [42] Lorenzen CJ. A method for the continuous measurement of in vivo chlorophyll concentration. Deep Sea Res Oceanogr Abstr 1966;13(2):223e7. [43] Falkowski P, Kiefer DA. Chlorophyll a fluorescence in phytoplankton: relationship to photosynthesis and biomass. J Plankton Res 1985;7(5):715e31.

Advances in sampling strategies and analysis of phytoplankton Chapter | 31 [44]

519

Holm-Hansen O, Lorenzen CJ, Holmes RW, Strickland JD. Fluorometric determination of chlorophyll. ICES J Mar Sci 1965;30(1):3e15. [45] Neveux J, Lantoine F. Spectrofluorometric assay of chlorophylls and phaeopigments using the least squares approximation technique. Deep-Sea Res Part I Oceanogr Res Pap 1993;40(9):1747e65. [46] Abaychi JK, Riley JP. The determination of phytoplankton pigments by high-performance liquid chromatography. Anal Chim Acta 1979;107:1e11. [47] Mantoura RF, Llewellyn CA. The rapid determination of algal chlorophyll and carotenoid pigments and their breakdown products in natural waters by reverse-phase highperformance liquid chromatography. Anal Chim Acta 1983;151:297e314. [48] Wright SW, Shearer JD. Rapid extraction and high-performance liquid chromatography of chlorophylls and carotenoids from marine phytoplankton. J Chromatogr A 1984;294:281e95. [49] Gieskes WW, Kraay GW. Analysis of phytoplankton pigments by HPLC before, during and after mass occurrence of the microflagellate Corymbellus aureus during the spring bloom in the open northern North Sea in 1983. Mar Biol 1986;92(1):45e52. [50] Jeffrey SW, Vesk M, Jeffrey SW. Introduction to marine phytoplankton and their pigment signatures. In: Mantoura RFC, Wright SW, editors. Phytoplankton pigments in oceanography. Paris: UNESCO Publishing; 1997. p. 37e84. [51] Barlow R, Stuart V, Lutz V, Sessions H, Sathyendranath S, Platt T, et al. Seasonal pigment patterns of surface phytoplankton in the subtropical southern hemisphere. Deep Sea Res I 2007;54:1687e703. [52] Roy R, Pratihary A, Mangesh G, Naqvi SW. Spatial variation of phytoplankton pigments along the southwest coast of India. Estuar Coast Shelf Sci 2006;69(1e2):189e95. [53] Naik RK, Anil AC, Narale DD, Chitari RR, Kulkarni VV. Primary description of surface water phytoplankton pigment patterns in the Bay of Bengal. J Sea Res 2011;65(4):435e41. [54] Jeffrey SW. Application of pigment methods to oceanography. In: Jeffrey SW, Mantoura RFC, Wright SW, editors. Phytoplankton pigments in oceanography: guidelines to modern methods. Paris: UNESCO; 1997. p. 127. [55] Garvey M, Moriceau B, Passow U. Applicability of the FDA assay to determine the viability of marine phytoplankton under different environmental conditions. Mar Ecol Prog Ser 2007;352:17e26. [56] Steinberg MK, Lemieux EJ, Drake LA. Determining the viability of marine protists using a combination of vital, fluorescent stains. Mar Biol 2011;158(6):1431e7. [57] Durkin CA, Marchetti A, Bender SJ, Truong T, Morales R, Mock T, et al. Frustule-related gene transcription and the influence of diatom community composition on silica precipitation in an iron-limited environment. Limnol Oceanogr 2012;57(6):1619e33. [58] Leblanc K, Hutchins DA. New applications of a biogenic silica deposition fluorophore in the study of oceanic diatoms. Limnol Oceanogr Methods 2005;3(10):462e76. [59] Patil JS, Jagadeesan V. Effect of chlorination on the development of marine biofilms dominated by diatoms. Biofouling 2011;27(3):241e54. [60] MacKenzie L, Beuzenberg V, Holland P, McNabb P, Selwood A. Solid phase adsorption toxin tracking (SPATT): a new monitoring tool that simulates the biotoxin contamination of filter feeding bivalves. Toxicon 2004;44:901e18. [61] Camacho FG, Rodrı´guez JG, Miro´n AS, Garcı´a MC, Belarbi EH, Chisti Y, Grima EM. Biotechnological significance of toxic marine dinoflagellates. Biotechnol Adv 2007;25:176e94.

520 Advances in Biological Science Research [62] Zhou ZH, Komiyama M, Terao K, Shimada Y. Effects of pectenotoxin-1 on liver cells in vitro. Nat Toxins 1994;2:132e5. [63] Bujalance I, Feijoo M, Lopez PM, Munoz MD, Munoz-Castaneda JR, Tunez I, et al. Protective melatonin effect on oxidative stress induced by okadaic acid into rat brain. J Pineal Res 2003;34:265e8. [64] Quay PD, Peacock C, Bjo¨rkman K, Karl DM. Measuring primary production rates in the ocean: enigmatic results between incubation and non-incubation methods at Station ALOHA. Glob Biogeochem Cycles 2010;24:GB3014. https://doi.org/10.1029/ 2009GB003665. [65] Marra J. Comment on “Measuring primary production rates in the ocean: enigmatic results between incubation and non-incubation methods at Station ALOHA” by Quay PD, et al. Glob Biogeochem Cycles 2012;26:GB2031. https://doi.org/10.1029/2011GB004087. [66] Gaarder T, Gran HH. Investigations of the production of plankton in the oslo fjord. Rapp Pv Reun Cons Perm Int Explor Mer 1927;42:1e48. [67] Carpenter JH. The accuracy of the Winkler method for dissolved oxygen analysis. Limnol Oceanogr 1965;10:135e40. [68] Slawyk G, Collos Y, Auclair JC. The use of the 13C and 15N isotopes for the simultaneous measurement of carbon and nitrogen turnover rates in marine phytoplankton. Limnol Oceanogr 1977;22:925e32. [69] Bender ML, Grande K, Johnson K, Marra J, Williams PLB, Sieburth J, et al. A comparison of four methods for determining planktonic community production. Limnol Oceanogr 1987;32:1085e98. [70] Steeman-Nielsen E. The use of radioactive carbon (14C) for measuring production in the sea. J Cons Perm Int Explor Mer 1952;18:117e40. [71] Kolber Z, Falkowski PG. Use of active fluorescence to estimate phytoplankton photosynthesis in situ. Limnol Oceanogr 1993;38:1646e65. https://doi.org/10.4319/ lo.1993.38.8.1646. [72] Robinson C, Williams PLB. Respiration and its measurement in surface marine waters. In: del Giorgio PA, Williams PLB, editors. Respiration in aquatic ecosystems. UK: Oxford University Press; 2005. p. 147e81. [73] Sakshaug E, Bricaud A, Dandonneau Y, Falkowski PG, Kiefer DA, Legendre L, et al. Parameters of photosynthesis: definitions, theory and interpretation of results. J Plankton Res 1997;19:1637e70. https://doi.org/10.1093/plankt/19.11.1637. [74] Laney SR. Assessing the error in photosynthetic properties determined by fast repetition rate fluorometry. Limnol Oceanogr 2003;48:2234e42. https://doi.org/10.4319/ lo.2003.48.6.2234. [75] Kyewalyanga MS, Naik R, Hegde S, Raman M, Barlow R, Roberts M. Phytoplankton biomass and primary production in Delagoa Bight Mozambique: application of remote sensing. Estuar Coast Shelf Sci 2007;74(3):429e36. [76] Behrenfeld MJ, Falkowski PG. A consumer’s guide to phytoplankton primary productivity models. Limnol Oceanogr 1997;42:1479e91. [77] Gaonkar CC, Kooistra WH, Lange CB, Montresor M, Sarno D. Two new species in the Chaetoceros socialis complex (Bacillariophyta): C. sporotruncatus and C. dichatoensis, and characterization of its relatives, C. radicans and C. cinctus. J Phycol 2017;53(4):889e907. [78] Li Y, Boonprakob A, Gaonkar CC, Kooistra WH, Lange CB, Herna´ndez-Becerril D, et al. Diversity in the globally distributed diatom genus Chaetoceros (Bacillariophyceae): three new species from warm-temperate waters. PLoS One 2017;12(1):e0168887.

Advances in sampling strategies and analysis of phytoplankton Chapter | 31 [79] [80] [81] [82]

[83]

[84]

521

Kooistra WH, Sarno D, Balzano S, Gu H, Andersen RA, Zingone A. Global diversity and biogeography of Skeletonema species (Bacillariophyta). Protist 2008;159(2):177e93. Johnson ZI, Martiny AC. Techniques for quantifying phytoplankton biodiversity. Ann Rev Mar Sci 2015;7:299e324. Rich VI, Konstantinidis K, DeLong EF. Design and testing of ‘genome-proxy’microarrays to profile marine microbial communities. Environ Microbiol 2008;10(2):506e21. Shilova IN, Robidart JC, Tripp HJ, Turk-Kubo K, Wawrik B, Post AF, Thompson AW, Ward B, Hollibaugh JT, Millard A, Ostrowski M. A microarray for assessing transcription from pelagic marine microbial taxa. ISME J 2014;8(7):1476e91. Nanjappa D, Audic S, Romac S, Kooistra WH, Zingone A. Assessment of species Ddiversity and distribution of an ancient diatom lineage using a DNA metabarcoding approach. PLoS One 2014;9(8):e103810. https://doi.org/10.1371/journal.pone.0103810. Regaudie-de-Gioux A, Lasternas S, Agustı´ S, Duarte CM. Comparing marine primary production estimates through different methods and development of conversion equations. Front Mar Sci 2014;1:19.