Diazotrophic cyanobacterium Trichodesmium spp. in China marginal seas: Comparison with other global seas

Acta Ecologica Sinica 35 (2015) 37–45

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Acta Ecologica Sinica j o u r n a l h o m e p a g e : w w w. e l s e v i e r. c o m / l o c a t e / c h n a e s

Diazotrophic cyanobacterium Trichodesmium spp. in China marginal seas: Comparison with other global seas Zhibing Jiang a,b, Jiangning Zeng b, Jianfang Chen b, Quanzhen Chen b, Dongsheng Zhang b, Xiaojun Yan a,* a b

Key Laboratory of Applied Marine Biotechnology, Ministry of Education, Marine College of Ningbo University, Ningbo 315211, China Key Laboratory of Marine Ecosystem and Biogeochemistry, Second Institute of Oceanography, State Oceanic Administration, Hangzhou 310012, China

A R T I C L E

I N F O

Article history: Received 13 June 2013 Revised 20 January 2015 Accepted 29 January 2015 Available online Keywords: Trichodesmium spp Nitrogen fixation Carbon fixation Biogeochemical cycle Marginal sea

A B S T R A C T

Trichodesmium spp. is probably the most important diazotroph in oligotrophic tropical and subtropical seas. Their N fixation relieves the restraining of N limitation to the marine primary production, and promotes the absorption of CO2. Therefore, their distribution and N fixation are paid more attention on the condition of global warming in recent years. We summarize the studies on Trichodesmium spp. in China marginal seas and other global seas/oceans, including population structure, distribution pattern, sampling methods, restriction factors, N fixation rate, and their contribution to primary/new production and N input, as well as their function in oceanic C and N biogeochemical cycles. Finally, we point out the future research directions in China marginal seas. © 2015 Ecological Society of China. Published by Elsevier B.V. All rights reserved.

Contents 1.

2.

3. 4.

5.

6.

Species composition and population structure ........................................................................................................................................................................................ 1.1. Species composition .............................................................................................................................................................................................................................. 1.2. Filament forms ........................................................................................................................................................................................................................................ 1.3. Size structure (cell number) ............................................................................................................................................................................................................... Abundance distribution ..................................................................................................................................................................................................................................... 2.1. Horizontal distribution in China marginal seas .......................................................................................................................................................................... 2.2. Horizontal distribution in global oceans ....................................................................................................................................................................................... 2.3. Vertical distribution .............................................................................................................................................................................................................................. Sample collection methods .............................................................................................................................................................................................................................. Main limiting factors .......................................................................................................................................................................................................................................... 4.1. Temperature ............................................................................................................................................................................................................................................. 4.2. P ................................................................................................................................................................................................................................................................... 4.3. Iron (Fe) ..................................................................................................................................................................................................................................................... Carbon, nitrogen fixation and function in biogeochemical cycles ..................................................................................................................................................... 5.1. Contribution of the biomass and primary production .............................................................................................................................................................. 5.2. Nitrogen fixation rate and budget ................................................................................................................................................................................................... 5.3. Function in carbon and nitrogen biogeochemical cycles ......................................................................................................................................................... Prospect .................................................................................................................................................................................................................................................................. Acknowledgements ............................................................................................................................................................................................................................................. References ..............................................................................................................................................................................................................................................................

38 38 38 38 38 38 38 40 40 40 40 40 41 41 41 41 42 43 43 43

* Corresponding author. Key Laboratory of Applied Marine Biotechnology, Ministry of Education, Marine College of Ningbo University, Ningbo 315211, China. Tel.: +86 574 87609570; fax: +86 574 87600590. E-mail address: [email protected] (X. Yan). http://dx.doi.org/10.1016/j.chnaes.2015.01.003 1872-2032/© 2015 Ecological Society of China. Published by Elsevier B.V. All rights reserved.

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Z. Jiang et al./Acta Ecologica Sinica 35 (2015) 37–45

Trichodesmium spp. (hereafter, Trichodesmium), the diazotrophic filamentous nonheterocystous cyanobacterium, is living as colonyforming or individual filaments. Trichodesmium are broadly distributed at the photic zone in oligotrophic tropical and subtropical seas, which probably contribute the major primary production and nitrogen (N) fixation [1], as they can fix the free dissolved gaseous N to the combined N by photosynthesis. Such N2 fixation relieves the restraining of N limitation to the marine primary production, produces more biological carbon (C), efficiently enhances the biological pump effects, and brings about more “effective” biological N sinks [2]. Therefore, they promote the absorption of CO2, and thereby regulate the global ocean/atmosphere CO2 balances [3]. With serious global warming problem recently, the distribution, C and N fixation of Trichodesmium are obtaining great interest. We find broad documents about their distribution and primary production around the global open seas/oceans, such as the eastern Caribbean Sea [4], Red Sea [5], Arabian Sea [6], Gulf of Mexico [7], East China Sea (ECS) [8–11], South China Sea (SCS) [12–14] and the upstream Kuroshio [13,15,16], coastal waters of Tanzania [17], Atlantic Ocean [18–20], Pacific Ocean [21,22], and Indian Ocean [23]. This article summarizes the studies on Trichodesmium spp. in China marginal seas and other global seas/oceans, including the population structure, distribution pattern, sampling methods, restriction factors, C and N fixation, and their contribution to primary/new production and N input, as well as their function in oceanic C and N biogeochemical cycles. Besides, we also propose the key scientific problems for future study in China marginal seas.

in the tropical North Atlantic Ocean. And they also pointed that at stations with low Trichodesmium densities, free trichomes often dominated while at stations with appreciable colony densities, the bulk of trichomes were in colonies. The free trichomes (single filaments) of attached cells can arrange themselves either in parallel or radially to form colonies, and also present the ability to fix N2 [20], although they do so with a lower per cell rate than when present in the colony morphology [22]. Individual free trichomes also show the highest growth rates under laboratory conditions [34,35], and their proportion is thus assumed to indicate an active dynamic state in Trichodesmium populations [20]. Previous work conducted at station ALOHA (central, subtropical North Pacific) [21,22] and at the BATS (Bermuda Atlantic Timeseries Study) site [36] has shown that free trichomes could represent an important fraction of their populations, contributing up to 75% of Trichodesmium N2 fixation. Therefore, the abundant individual filaments in the ocean also play an important role in the global N2 fixation. Wu et al. [33] reported the Trichodesmium size structure of bottle sample in SCS usually less than 10 trichomes per colony. However, this finding greatly lower than the values reported in Gulf of Aqaba, Red Sea (range 35–190) [5], subtropical eastern North Pacific Ocean (182 trichomes per colony; range 10–375) [21], subtropical eastern North Atlantic gyre (112 ± 47 for puffs [range 57–156], and 49 ± 16 for tufts [range 31–63]) [20], and tropical North Atlantic Ocean (98 ± 11, range 53–153) [19]. 1.3. Size structure (cell number)

1. Species composition and population structure 1.1. Species composition By far 8 Trichodesmium species are identified both by the methods of traditional morphology and modern molecular biology, consists of T. thiebautii, T. erythraeum, T. hildebrandtii, T. aureum, T. contortum, T. havanum, T. pelagicum, and T. tenue [24]. The first three species mentioned are also recorded in the ECS, especially T. thiebautii is the absolute dominant species [9,25]. This feature is similar to other China marginal seas and coastal waters (e.g., southern Yellow Sea, Fujian costal areas, Taiwan Strait, Sanya Bay, and central SCS) [9,26], Kuroshio areas [8,27], Caribbean and Sargasso Sea [28], and tropical North Atlantic Ocean [19]. However, the harmful algal bloom in ECS is usually caused by the T. erythraeum and T. hildebrandtii [8], and so does the other seas around the world, such as Daya Bay in SCS [29], Arabian Sea [30], Mediterranean Sea [31], and Northwest African Upwelling [32]. Thus, we need further study on this phenomenon. 1.2. Filament forms Most often, studies have considered only two of the macroscopic colonial forms, i.e. “tufts” (fusiform colonies) and “puffs” (spherical ones), although the individual free trichomes usually present and commonly ignored [20]. However, the free form is the absolute dominant population in SCS and southern ECS, while the colonial trichomes were rarely observed [10,33]. Likewise, other studies in the Pacific Ocean have also reported a preponderance of Trichodesmium biomass as free living trichomes rather than colonies [21,34]. This trait is in better agreement with the recent investigation in the subtropical eastern North Atlantic Ocean [20]. Especially at the station near the Madeira, the free trichomes account for 100% in the population with a minimal biomass of <10 3 trichomes/m2, although the proportion of colonial individuals increased steadily with increasing longitude (away from the eastern coast) and abundance from this station. Nevertheless, Carpenter et al. [19] found that the colonial filaments were dominant (≥89%) forms

Every filament of Trichodesmium in different seas is different in cell numbers. The mean trichome length was 107 cells in the southern ECS [10]. This value matched well with the typical average cells per trichome (~100) [1,21], but much higher than the average number of cells per filament (70) in the Atlantic Ocean [18]. Besides, a specific difference was found in the average cells per trichome. For example, in the open waters of the Gulf of Aqaba, Red Sea, where samples collected by 100 μm mesh plankton net showed Trichodesmium sp. (110–270) ≈ T. tenue (80–310) > T. thiebautii (55–180) > T. erythraeum (40–130) [5]. And their colony shapes presented as spherical (puffs), bow-tie, parallel-straight, and paralleltwisted, in turn. 2. Abundance distribution 2.1. Horizontal distribution in China marginal seas It is seems that Trichodesmium has latitudinal distribution patterns that the abundance increases from high latitude to low latitude, though the samples were collected using different methods (Table 1): the net-collected data of China seas indicated that the abundances of Trichodesmium in ECS (e.g., total area, Taiwan Strait, and southern ECS) were similar, but significantly higher than SYS; the depth integrated abundance shows that Upstream Kuroshio > Kuroshio in southern ECS > SCS > Upwelling in southern ECS. The abundance in China marginal seas had significant seasonal differences, and the abundances in summer and autumn were much higher than those in winter and spring (Table 1). However, Trichodesmium in the Kuroshio areas bloomed much earlier than in the ECS and SCS, and declined quickly in autumn. This indicated that Trichodesmium in China marginal seas might have been introduced by the Kuroshio. 2.2. Horizontal distribution in global oceans Trichodesmium overall distribution represents strong patches and strictly geographical limitation, mainly can be found in tropical and subtropical oligotrophic oceans/seas. They usually encountered in

Z. Jiang et al./Acta Ecologica Sinica 35 (2015) 37–45

39

Table 1 Trichodesmium density in the southern Yellow Sea (SYS), East China Sea (ECS), and South China Sea (SCS). Study area

Year

Density Spring

SYS (32°–34°N, 123.5°–124.5°E) ECS (28°–32°N, 122°–127°E) Coastal water of Fujian (23.4°–27.4°N, 117.0°–120.4°E) Taiwan Strait (depth >20 m) East of Taiwan (18.2°–25.8°N, 120.8°–129.2°E) Northern ECS (~30°–32.5°N, 122°–127°E) Southern ECS

Central SCS (12°–20°N, 111°–118°E) Northern SCS Mesoscale cyclonic eddy, SCS Sanya Bay, SCS (18°11′–18°18′N, 109°20′–109°30′E) Northern SCS (18°–22.5°N, 111.5°–123°E) Upstream Kuroshio (19°–24°N,121°–124°E) Northern SCS basin (18°–22°N) Kuroshio in southern ECS Upwelling in southern ECS

1977–1978

0.60 0.93 0.01 6.07 4.46 0.96

1984–1985 1996–1997 2006–2007 1994 1995 1994 1995 1983–1984 2004–2007 2007 2004 2000–2002 2002–2006 2004–2007 1995–1996 1994–1995

0.69 5.8 23.77 435 2510 850 6050 55

Sampling method Summer 1.13 17.93 13.68 22.16 15.27 35.70 19.7 8.8 14.4a 58.6a 2.14 15.5 330–850b 2755.60 5267 34610 4870 6000

Autumn

Winter

4.40 12.07 11.96 32.02

0.37 3.27 0.05 2.38

16.26

3.17

Reference

Net (64 μm)

[9]

Net (77 μm) Net (77 μm) Net (20 μm)

[32] [27] [37]

2.5-L bottle (1 L) 4.01 31.3

0.30 1.9

248.07 5345

165.70

3550 610 380

490 150 20

Net (64 μm) 20-L bottle (1.2/2.4 L) Bottle (10 L) Net (20 μm) 20 L bottle (1 L) 20 L bottle (1.2/2.4 L) 20 L bottle (1.2 or 2.4L) 2.5 L bottle (1 L)

[9] [16] [14] [26] [12] [15] [10]

Abundance is adopted by ×103 trichomes/m3. Boldfaced number indicated the depth integrated abundance (×103 trichomes/m2). a Sample at 10 m depth. b Sample at top 200 m.

high biomass in western boundary currents (e.g., Gulf Stream and Kuroshio), tropical central gyres, and several margin seas (Tables 1 and 2). These water regiments are generally characterized by low N level, very clear waters, and deep light penetration [1]. Generally, it seemed that Trichodesmium abundances in Atlantic Ocean are much higher than that in Pacific Ocean (Table 2). These findings

consisted with the supposition by previous researchers [44,45], who suggest a greater new N production, presumptively through biological N2 fixation, in North Atlantic gyre, compared to North Pacific. Besides, Table 2 also shows that their densities are much greater in the tropical North Atlantic, compared to the sub-tropical zone, although this trend is not observed in North Pacific.

Table 2 Trichodesmium density in global open seas/oceans. Study area

Time

Sample method

Abundance

Reference

N. Pacific (0°–44°N, 155°E) Central Pacific (7°–10°N, 155°–159°E) Eastern Pacific (3°–7°N, 84°–143°E) Western N. Pacific (20°–21°N, 121°–156°E) Southeast Asian sea (11°–9°S, 100°–112°E) South Indian Ocean (10°–37°S, 52°–97°E) South and east of Madagascar, Indian Ocean (21°–28°S, 37°–56°E) Arabian Sea (10°–22°N, 65°–74°E) N. Atlantic gyre (15°–30°N, 30°W) Equatorial Atlantic (5°S–15°N, 30°W) S. Atlantic gyre (5°–30°S, 30°W) Subtropical N. Atlantic (26°N, 17°–38°W) Subtropical N. Atlantic (29°N, 15°–29°W) Subtropical N. Atlantic (24°N, ~15.5°–80°W) Northwestern Gulf of Mexico Tropical N. Atlantic (0°−25°N, 27°−80°W)

Feb–Mar 2007 Jul 2008 Aug 2008 May 2008 Mar 2009 Feb 2009 Feb 2005 2000–2004 2007–2008

Bottle Bottle (4 L)

1.73b 1.76b 0.87b 25.2b 33.3b 1.52b 312b 24138 19.5b 222b 0.75b 6.6b 0.8b 3.88 1500b 2250b 292b 222b 300b 52.3 7200 5600 91,000 21,000 8600 180–1400a ~8500 ~10–100 5338

[38] [23]

Tropical N. Atlantic (0°−15°N) Station ALOHA (22°45′N, 158°W) N. Pacific (19°–25°N, 154°–160°W) N. Pacific (18°–28°N, 170°E–154°W) Tropical N. Atlantic (0°−25°N, 27°−80°W)

N. Atlantic (25.4°–34.1°N, 13.9°–36.7°W) Tropical N. Atlantic (0°−15°N) Gulf of Aqaba, Red Sea (27.5°–29.5°N, 34.5°–35°E) Arabian Sea (10°–22°N, 65°–74°E)

Autumn 2007 Spring 2008 Jan–Mar 2011 Jul 2000 May–Jun 1994 Apr 1996 Oct 1996 1995–1999 1989–1992 Autumn 2002 Summer 2003 May–Jun 1994 Apr 1996 Oct 1996 Oct 2006 1995–1999 Summer 1996 2000–2004

Abundance is adopted by ×103 trichomes/m3. Boldfaced number indicated the depth integrated abundance (×103 trichomes/m2). a The unit was ×103 free trichomes/m2. b Surface sample.

Bottle (10 L), filtered by 50 μm mesh Bottle Pump (50–130 L), filtered by 40 μm mesh

Net (40 μm) Bottle 10-L bottle

30-L bottle (30 L) Net and bottle 10-L bottle (10 L) Bottle

Net (53 μm) Net (65 μm) Net (100 μm) Bottle

[39] [6] [40]

[41] [42] [7] [19]

[18] [21] [43] [19]

[20] [18] [5] [6]

40

Z. Jiang et al./Acta Ecologica Sinica 35 (2015) 37–45

2.3. Vertical distribution Maximum Trichodesmium density is found at a depth of 10–40 m in the upper water column [1]. The most biomass usually occurred at a depth of 20–30 m both in southern ECS [10] and northern SCS [12]. Sometimes, a second maximum appeared near the surface. It is observed that their abundance decreased rapidly when depth deeper than 50 m. The maximum abundance in south-west of Atlantic Ocean is found at the depth of 15 m [46]. Investigation in the tropical North Atlantic showed that peak abundance was generally in the upper water column, with an average biomass maximum at 12 m in May–June 1994 and October 1996 cruises and at 40 m in April 1996 [19]. Therefore, the vertical distributions are different among different seas and seasons. The light intensity difference is the main reason, as Trichodesmium has high dark respiration rate, high light compensation point, and high light saturation value. So it prefers the environment with high light intensity [46], and the water depths mentioned earlier are just fit for their light saturation. 3. Sample collection methods As presented in Tables 1 and 2, the collection methods of Trichodesmium are mainly consisted of various mesh-size (20– 100 μm) plankton nets and different volume bottles (sample water volumes). Larger mesh sizes of the tow nets were also used to capture them, such as 202 and 335 μm plankton nets were performed in the eastern Caribbean Sea and at the BATS, respectively. And Trichodesmium sample can also be pumped through a pump, and the obtained large (50–130 L) bulk seawater then filtered by gravity through a small (40 μm) nylon mesh. Recently, the in situ colony abundance was underwater semi-automatic measured successfully by a Visual Plankton Recorder [47], although the free trichome was arbitrarily neglected currently. Water samples with a volume from 250 mL to 30 L are routinely used for such purposes (Tables 1 and 2), and a process of sedimentation or filtration is required to concentrate trichomes before the actual counting [8,37]. Thus, sampling issues appeared as one key determinant of Trichodesmium studies, and it should be noted that the sampling methods employed in all these studies differed greatly from those used around the global seas/oceans. The discrepancies observed may come from the fact that the two methods operate on very different spatial scales [37]. Because a bottle sample represents trichome abundance in the vicinity of a rosette sampler, while a net tow represents the mean abundance across several ten or hundred meters. For the water samples, a drawback of this approach is that large, but sparse, colonies are easily missed due to small sample volumes [1]. In addition, unless the intracellular gas vesicles are destroyed by acid treatment, the positively buoyant Trichodesmium trichomes may float to the surface during the sedimentation procedure [1]. Another consideration of bottlecollected samples is that a sample with very limited volume may not be representative enough to reflect their abundance at a sampling station [37]. Indeed, these species present positive buoyancy, great size and low concentrations in the water column with respect to other phytoplankton groups. In this sense, bottle sampling has been criticized because the low volume of water sampled can introduce a bias in colony abundance estimates [20,37]. To overcome this difficulty, the general recommendation is to use a mesh size of less than 100 μm and to tow the net at a low speed [1] for the estimation of Trichodesmium abundance. Nets were nevertheless preferred over bottle samples [20,37], it could promote colony fragmentation and allow the escape of free trichomes (their cells are generally 5–15 μm in diameter) through the mesh and on occasion the volume filtered can be overestimated because of clogging. Therefore, the net-collected methods are prone to underestimate the free trichomes density.

Different sampling methods have the merits and demerits. Bottle sample is more appropriate (not definitely) when the areas are dominated by the free forms. The sample volume is determined by the abundance. When the abundance is high or even the sampling areas bloom, the sample volume can be less; and when the abundance is low, the sample volume can be more. The vertical tow sample is more suitable when the areas are dominated by the colony population. However, along the short transect in the southern ECS, these two methods generated approximately the same distribution pattern for Trichodesmium [37].

4. Main limiting factors 4.1. Temperature It is generally considered that seawater temperature sets a physiological constraint to the geographic distribution of Trichodesmium, and the 20 °C is the common lowest temperature for their normal growth [1]. Although they are also found in waters colder than 20 °C, growth and activity are usually restricted. Laboratory experiment of Breitbarth et al. [48] also demonstrated that the strain IMS-101 of Trichodesmium was adapted to optimal growth at temperatures between 24 °C and 30 °C and can tolerate water temperatures from 20 °C to 34 °C. Recently, Fu et al. [49] showed thermal limits of Trichodesmium ranged from 18 °C to 32 °C, and optimum growth temperatures were ~26 °C. A significant positive correlation between their abundance and temperature was observed in other field studies [1,12,17,40]. Also, the abundance distribution of Trichodesmium is greatly controlled by the temperature besides the effect of Kuroshio (Table 1). However, this correlation is generally attributed to oceanographic features associated with warm waters, such as shallow mixed layer depth, high light intensity, and oligotrophic conditions rather than a direct physiological response to temperature itself [50]. For example, some researchers did not find such relation [18,19,23].

4.2. P If phytoplankton assimilate N and P at a fixed ratio (16N:1P), excess N input from N2 fixation increases biological P uptake which depletes surface P and alters the dissolved N/P. However, the bottom P input is not enough to satisfy their growth, thus P become the limited factor. The stoichiometry of 600C:101N:1P in Trichodesmium collected from the Sargasso Sea reflects P-limiting conditions [51]. N2 fixation in the Fe-replete subtropical North Atlantic is thought to be limited by available P [52,53]. To overcome P limitations in oligotrophic waters, they migrate vertically in the water column to scavenge P using a buoyancy-regulating mechanism [54,55]. Carbohydrate:protein ratio was the best predictor of buoyancy and fit a cosine curve with increasing values during the day and decreasing values at night in cycles that paralleled observed diel buoyancy patterns. In the western Gulf of Mexico, the N/P in sinking (87.0) and ascending (43.5) colonies provide the best direct evidence to date of vertical migration for P acquisition, although no significant differences between them were found in the waters of northern Australian coast and northern Hawaii [54]. Experimental studies showed that Trichodesmium is a poor competitor for dissolved inorganic (DIP) relative to bulk phytoplankton and might meet a majority of its P demand by taking up dissolved organic phosphorus (DOP, e.g., phosphonate and monophosphate esters) [56,57]. Thus, their P uptake enhanced via the uptake of both DIP and DOP under P-limiting conditions [56–58]. Interestingly, experimental results of Spungin et al. [59] show that Trichodesmium can adjust a series of cellular pathways to compensate for low P availability.

Z. Jiang et al./Acta Ecologica Sinica 35 (2015) 37–45

4.3. Iron (Fe) Fe is the important composition of nitrogenase, and its absorption and utilization directly affect the synthesis and expression of nitrogenase [60], thus influencing Trichodesmium growth and distribution [61,62]. Berman-Frank et al. [60] found that their N2 fixation rate is closely related to the biological absorption of Fe and Fe/C ratio in cell. Adam [63] proved that the N2 fixation rate positively correlated with Fe/C ratio in cell during the Fe limitation range. So their growth and N fixation are both affected by Fe biological absorption. Organisms using N2 as their only N source require one order of magnitude higher Fe than cells using fixed N (see the references in Refs. 33 and 64). Therefore, Fe supply has been considered as the most limiting factor for the distribution, growth rate, and metabolic activity of this genus and, by extension, N2 fixation in the ocean [3,61,62,65]. Especially in the rich-nutrients and low-chlorophyll areas, Fe is the main limitation factor for the primary production [41]. Both the field fertilization tests [66] and laboratory cultural experiments [60,61] suggest that Fe is a key factor limiting their N2 fixation and growth rates, as well as the novel molecular method monitoring the expression of an Fe limitation-induced gene [67]. Interestingly, Trichodesmium can absorb and save a lot of molecular states Fe [51] to meet the demand of maximum biomass, and it may be an adaption mechanism for their low Fe tolerance in the surface water [63]. Fe exists in seawater as particulate (>0.4 μm) and dissolved (<0.4 μm) forms [33]. Particulate Fe cannot diffuse easily in water, and thus most of particulate Fe is not considered to be directly available to marine microorganisms [33]. Dissolved Fe has long been considered to exist in seawater predominantly as soluble Feorganic complexes that are available for assimilation [67,68], although recent evidence has shown that only a small portion of the “dissolved” Fe exists as soluble Fe species whereas much of the “dissolved” Fe is present as less available colloidal particles of 0.02– 0.4 μm diameter [65]. Fe enters the euphotic water by the atmospheric subsidence. Based on the present deposition, Fe availability limits N fixation by this organism in 75% of the global ocean [60]. Recent studies [41,53,69] demonstrated a close association between dissolved Fe concentration, in turn related to increased atmospheric deposition of Saharan dust, and N2-fixation rates in the Atlantic Ocean. Fernández et al. [41] suggested that Fe supply through atmospheric deposition is a major determinant of planktonic N2 fixation in the Atlantic Ocean, as well as in the North Pacific [38]. The levels of soluble reactive P in the surface waters of the subtropical North Pacific are one order of magnitude higher than those in the western subtropical North Atlantic [65]. Meanwhile, 10-fold higher concentrations of “dissolved” Fe in the North Atlantic compared to the North Pacific [65]. The abundance of Trichodesmium in North Pacific is much lower than those in North Atlantic (Table 2). Carpenter et al. [19] believed it might relate to Fe, as aeolian dust is a major source of iron to the upper ocean, and this flux is greatest in the North Atlantic [70]. However, Wu et al. [33] suggested that N fixation can be limited by available Fe even in regions with a high rate of dust deposition such as the SCS. 5. Carbon, nitrogen fixation and function in biogeochemical cycles 5.1. Contribution of the biomass and primary production The contribution of Trichodesmium to the biomass (total Chla) presents spatio-temporal difference due to their dominance variation in phytoplankton biomass. For example, they accounted for 62%, 13% and 11% of depth integrated total chlorophyll for May– June 1994 and April and October 1996 [19]. Carpenter and Price [28] found Trichodesmium to account for, on average, about 61% of total

41

Chla in the Caribbean Sea and about 5% in the Sargasso Sea. Letelier and Karl [21] investigated their chlorophyll to account for about 18% of total Chla at Station ALOHA [71]. In contrast, Zhang et al. [23] reported the surface Trichodesmium chlorophyll a to account for 7.79%, 0.35%, 0.06%, 0.37%, and 3.92%, in subtropical west, tropical central, tropical eastern North Pacific, south Indian, and tropical southeastern Asian, in turn. The Trichodesmium contribution to the primary production has temporal and spatial differences. At the station ALOHA, they accounted for 4% of the total C assimilation between 1989 and 1992 [21,72]. Chang et al. [10] estimated their contribution to primary production in Kuroshio near Taiwan to be from 0.2% to 2.3% of the total. Parab and Matondkar [6] estimated Trichodesmium contributed 0.11–60% of total primary production in the Arabian Sea. During investigation of Carpenter et al. [19], they accounted for an average of 47%, 7.9%, and 11%, respectively, of total primary production for each cruise, indicating a greater importance for them in terms of C and N assimilation in the tropical North Atlantic relative to the North Pacific. The Trichodesmium contribution to phytoplankton standing crop and primary production in the subtropical and tropical waters, they may also contribute to the enhancement of overall primary and export production through N2 fixation. N2 fixation by Trichodesmium introduces new N into the euphotic zone and, via recycling and excretion of N, can promote the growth of other phytoplankton [19]. Despite a low contribution to primary production, Letelier et al. [72] concluded that 27% of new production could be supported by their N2 fixation. 5.2. Nitrogen fixation rate and budget The two commonly applied methods to assess N2-fixation rates are the 15N2-tracer addition and the acetylene (C2H2) reduction assay. With the rapid development of isotope mass spectrometry, 15 N 2 -tracer addition can transform 15 N 2 directly to PON for Trichodesmium N fixation rate determination [73]. This method is much more sensitive, precise and simple than the acetylene reduction assay [43]. However, Mohr et al. [74] demonstrated that the 15 N2-tracer addition method underestimates N2-fixation rates significantly when the 15N2 tracer is introduced as a gas bubble. They proposed and tested a modified 15N2 tracer method based on the addition of 15N2-enriched seawater that provides an instantaneous, constant enrichment and allows more accurate calculation of N2-fixation rates for both field and laboratory studies. We summarized the direct areal estimates of Trichodesmium N2fixation rates in the global oceans and main margin seas. Table 3 suggests prominent differences (0.1–1108.15 μmol N/m2·d) of average N2-fixation among the regions and times. Especially in the subtropical waters such as the ECS and Sargasso Sea, previous studies have detected appreciable population only for limited periods during warm seasons (Tables 1 and 2). Subtropical studies found that even during the summer (2.1–59.4 μmol N/m2·d), when Trichodesmium was relatively abundant and active, areal rates of N2 fixation were generally low compared with tropical waters (35–278 μmol N/m2·d) [1]. Likewise, Fernández et al. [41] measured the mean N2-fixation rates in the Equatorial and North gyre regions of Atlantic Ocean (55–66 and 11–25 μmol N/m2·d, respectively), then estimated an annual N2 fixation of ~6 Tg N in the North Atlantic (0–40°N) and ~1.2 Tg N in the South Atlantic (0–40°S). In tropical area, despite low growth rates, the relatively high biomass in these regions implies that it may account for a quantitatively important input of organic C and N, and data from several studies have supported this idea [38,41,69,71,78,79]. Furthermore, the typical value of N2-fixation rates under bloom conditions could up to 1500 μmol N/m2·d [78]. However, Großkopf et al. [80] deemed that the most widely used method of measuring oceanic N2-fixation rates underestimates the contribution

42

Z. Jiang et al./Acta Ecologica Sinica 35 (2015) 37–45

Table 3 N2-fixation rates (μmol N/m2·d) of Trichodesmium spp. in China marginal seas and other global seas/oceans determined by different methods. Region

Time

Method

Fixation rate

Reference

Kuroshio in southern ECS (25°–25.5°N)

Mar 1995 May 1995 Sep 1995 Jan 1996 Mar 1995 Jul 1994 Sep 1995 Summer 1977 Jan 2004 Apr 2004 Aug 2004 Oct 2004 Winter 2006 Spring 2005 Summer 2002 Winter 2004–2007 Spring 2004, 2005 Summer 2004, 2006 Autumn 2004–2006 Oct, Dec 1972 Sep-Oct 2002 Jul-Aug 2003 May 1995 Bloom, 1995 Background, 1995 Nov 2001 Jan–Feb 2003,2004; Dec 2004 Apr 2000; Mar 2003, 2004 1975–1998 Feb–Mar, Aug 1974 Feb–Mar, Aug 1974 Aug 1974 May 1994 Jul 2000 1990–1992 May 1994 1994–2003 1994–1996 Oct 2006 1995–1999

CV1

6.2 59.4 3.4 0.1 0.3 33.6 2.1 126 0.8 92 10 5.5 2.4 ± 0.5 34.7 ± 11.0 168.1 ± 167.6 1.2 ± 0.5 7.3 ± 5.2 12.6 ± 5.7 5.0 ± 0.6 134 169 55 35 ± 7.4 129 40 1108.15 ± 973.8 24.75 ± 5.9 720.36 ± 425.3 117a 77 ± 9.7 4.2 ± 4.0 6.2 ± 4.0 278 ± 129 84.5 ± 17.7 84 ± 43 73 ± 22 239 ± 38 41.1 11 200

[10]

Upwelling in southern ECS (25.5°–26°N)

SE ECS (10°–25°N) Sanya Bay, SCS (18°11′–18°18′N, 109°20′–109°30′E)

Kuroshio (19°–24°N,121°–124°E)

Northern SCS basin (18°–22°N)

N. Pacific (21°N, 159°W) N. Pacific (19°–25°N, 154°–160°W) N. Pacific (18°–28°N, 170°E–154°W) Arabian Sea (7°–10°N) Arabian Sea (Along 65°E) N. Arabian Sea Arabian Sea (10°–22°N, 65°–72°E)

Coastal waters of Tanzania (~6°S, 39°E) Caribbean Sea (12°–22°N) Caribbean passages (22°–23°N) Sargasso Sea (22°–36°N) NE Caribbean Northwestern Gulf of Mexico Station ALOHA (22°45′N, 158°W) SW North Atlantic (14°–22°N) N. Atlantic (0–30°N, 25°–75°W) Station BATS (31°50′N, 64°10′W) N. Atlantic(25.4°–34.1°N, 13.9°–36.7°W) Tropical N. Atlantic (0°−15°N, 20°–30°W)

C2H2 C2H2

15N

2

15

N2

C2H2 C2H2 C2H2 C2H2 C2H2 CV2

C2H2 C2H2

C2H2 15 N2 C2H2 C2H2 C2H2 15 N2 STAE AV

[75] [26]

[15]

[76] [43] [1] [30] [6]

[17] [28]

[1] [7] [77] [1] [78] [36] [20] [18]

a μmol N/m3·d. STAE: size- and temperature-adjusted estimates; CV1: calculated value according to C2H2 measurements conducted in Kuroshio near southern Japan; CV2: calculated value according to C2H2 measurements conducted in the Arabian Sea [30]; AV: assumed value from abundance, cell nitrogen content, cell per filament ratio, and cell nitrogenspecific N2-fixation rate.

of diazotrophs relative to a newly developed method [74]. Their data show that in areas dominated by Trichodesmium, the established method underestimates N2-fixation rates by an average of 62%. Capone et al. [1] assessed a global annual input of ~80 Tg N by Trichodesmium fixation based on the biological rate measurements and it is thought that ~40–60% of this amount is directly attributable to non-bloom Trichodesmium [81]. However, according to the Trichodesmium data of satellite remote sensing during 1998–2003, Westberry and Siegel [82] estimated the annual global N fixation rates by their blooms is ~42 Tg N, while under nonbloom conditions is an additional ~20 Tg N. The geochemical estimates of total pelagic N 2 fixation are 100–200 Tg N/yr [44,45,83,84]. Recently, Großkopf et al. [80] estimated that the global marine N2-fixation rate derived from direct measurements may increase to 177 ± 8 Tg N/yr. Thus, Trichodesmium is a significant source of fixed N to the ocean, especially the blooms. This suggested that Trichodesmium is likely the dominant organism in the global ocean new N budget, although other diazotrophs (e.g., Richelia and unicellular cyanobacteria) also play an important role in N2 fixation [43,85,86]. In addition, a substantial fraction of the N2 fixed by diazotrophs can be released as dissolved organic nitrogen (DON), whose subsequent remineralization represents an additional source of new N for the ecosystem [87]. For example, geochemical estimates of N2 fixation, which integrate over wide spatial and temporal

scales and take into account also the production of DON, are in the range 15–56 Tg N/yr for the whole Atlantic Ocean [88]. 5.3. Function in carbon and nitrogen biogeochemical cycles Trichodesmium is the important source of new N and main supplier of new production in the subtropical and tropical Atlantic [36,81,89] and Pacific Ocean [21,45]. N source in oceans is mainly from the upwelling nitrate and biological N fixation. The nitrate input to oceans usually accompanies by the release of CO2, which dramatically decreases the oceanic C net-absorption. However, the biological N fixation enhances the oceanic C net-absorption [1,83]. And their N fixation is the major driving force of C transferring to deep sea [83]. Therefore, Trichodesmium plays a key role in global oceanic C and N geochemistry cycles. The significantly increasing of pCO2 influences the global climate change, and the oceans are the biggest absorption implement for CO2. The global N fixation by Trichodesmium is directly related to CO2 absorption of ocean [1,83,90]: when the N fixation increases, the C absorption will increase, too. And because the N absorption by marine phytoplankton has positive correlation with the absorption of dissolved inorganic carbon, it is just the negative feedback effects to climate. Otherwise, the expected rise in global sea surface temperature and pCO2, leading to increased stratification, enhanced

Z. Jiang et al./Acta Ecologica Sinica 35 (2015) 37–45

acidification, decreased mixed layer depth and nutrient availability, has been suggested to result in an increase in nitrogen fixation [91–94]. For example, Boyd and Doney [91] predicted a future increase of N2-fixation by 27% (80–94 Tg/yr) due to a floristic shift toward diazotrophy by Trichodesmium caused by combined effects of mixed layer depth, stratification, and nutrient distribution. According to the cultural Trichodesmium in different levels of pCO2, Levitan et al. [93] also expected the elevated CO 2 to increase diazotrophic N2 fixation and growth, thereby enhancing inputs of new N and increasing primary productivity in the oceans. Spungin et al. [59] suggested that elevated pCO2 could provide Trichodesmium with a competitive dominance that would extend its niche, particularly in P-limited regions of the tropical and subtropical oceans. However, based solely on the observed dependence of Trichodesmium growth on temperature, Breitbarth et al. [48] forecasted the increase in temperature may also result in a reduction of the area characterized by optimum nitrogen fixation and growth. 6. Prospect In conclusion, the N2-fixation of Trichodesmium is a crucial process of air–sea interaction, and deeply influences the oceanic C and N biogeochemical cycle, as well as their responses to global change. The research on Trichodesmium in China marginal seas has rarely been found, and existing literature is only limited on their species composition and abundance distribution in certain areas. The available reports show that the ECS and SCS both have abundant Trichodesmium. However, we do not have an overall understanding of their population structure, distribution pattern, and major restrict factors in China until now. And we also know little about their contribution to the primary/new production and N input in China marginal seas. Besides, their growth and distribution are profoundly influenced by the water masses and circulation. The present documents could not explain the relationship between Trichodesmium (e.g., distribution, N fixation, and harmful bloom) and hydrodynamics sufficiently. The problems mentioned earlier all need further study to deeply understand the contribution and function of Trichodesmium in C and N biogeochemical cycles in China marginal seas. Besides, the rapid developmental genomics provide a good tool to explore the molecular-level responses of Trichodesmium to the environmental forcings (e.g., temperature increase, P and Fe limitation, and light regulation), and thereby better interpret their physiological representation (e.g., growth, photosynthesis, N fixation rate, nutrient and light availability, and vertical migration) and function in biogeochemical cycles of ECS and SCS. Moreover, the molecular biological methods would help to understand the marine diazotrophic community structure and contribution to the N fixation, as unicellular cyanobacteria and bacterioplankton and endosymbiont Richelia that are expressing nitrogenase (nifH) genes can also support a significant fraction of total new production in oligotrophic waters [13,85,86]. Recently, Zhang et al. [95] found relatively high N2 fixation rate (>40 μmol N/m2·d) in the Yellow Sea Cold Mass, which was highly possible attributed to the unicellular diazotrophs. Also, Chen et al. [13] showed that unicellular diazotrophs contributed 65% of the total N2 fixation in the SCS and 50% in the Kuroshio. Therefore, we should pay more attention on the Trichodesmium and other diazotrophic groups in China marginal seas. Acknowledgements This project was supported by the National Basic Research Program of China (2010CB428903), National Marine Public Welfare Research Project of China (201305009, 201205015 and 201405007), and National Natural Science Foundation of China (41206103 and 41206104).

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