amount of CO2 ice in a martian polar cap can be setby the condition that it not liquefy at its base as aresult of hydrostatic pressure load, giving an upperlimit of ;60 mbar CO2 [C. Sagan, J. Geophys. Res.78, 1250 (1973)].

40. A. C. Lasaga, H. D. Holland, M. J. Dwyer, Science174, 53 (1971).

41. W. E. McGovern, J. Atmos. Sci. 26, 623 (1969).42. K. J. Zahnle, J. Geophys. Res. 91, 2819 (1986).43. These altitudes are comparable to those calculated

by setting the CH4 optical depth t 5 1 at l 5 1216 Å,the Lyman a wavelength. For a CH4 scale height H 58.4 km, CH4 cross section at 1216 Å s(CH4) 5 2 310217 cm22 (41), atmospheric surface number den-sity n(0) 5 2.6 3 1019 cm23, and [CH4] 5 1025, t 51 occurs at an altitude z 5 H ln{[CH4]s(CH4)n(0)H} '70 km. For present [CO2], CO2 adds only ;10% to tat 1216 Å.

44. In charged-particle irradiation experiments with2.5% H2O in CH4-NH3 atmospheres, organic sol-ids are also plentifully produced, involving alde-hydes, ketones, furans, and carboxylic acids asindications of oxygen chemistry [B. N. Khare, C.Sagan, J. E. Zumberge, D. S. Sklarew, B. Nagy.Icarus 48, 290 (1981)]. Similar results are found inUV irradiation experiments [K.-y. Hong, J.-H.Hong, R. S. Becker, Science 184, 984 (1974); B. N.Khare, C. Sagan, E. L. Bandurski, B. Nagy, ibid.199, 1199 (1978)].

45. G. D. McDonald, W. R. Thompson, M. Heinrich, B.N. Khare, C. Sagan, Icarus 107, 137 (1994).

46. V. R. Oberbeck, C. P. McKay, T. W. Scattergood, G.C. Carle, J. R. Valentin, Origins Life 19, 39 (1989).

47. M. Ackerman, in Mesospheric Models and RelatedExperiments, G. Fiocco, Ed. (Reidel, Boston, 1971),pp. 149–159.

48. B. N. Khare et al., Icarus 60, 127 (1984).49. C. Sagan and W. R. Thompson, ibid. 59, 133 (1984).50. W. A. Noyes and P. A. Leighton, The Photochemistry

of Gases (Reinhold, New York, 1941).51. G. Toupance, A. Bossard, R. Raulin, Origins Life 8,

259 (1977).52. This result derives from continuous-flow experi-

ments, in which the mean free path is much less thanthe radius of the reaction tube, so wall effects shouldnot be important [W. R. Thompson, G. D. McDon-ald, C. Sagan, Icarus 112, 376 (1994)].

53. K. J. Zahnle and J. C. G. Walker, Rev. Geophys.Space Phys. 20, 280 (1982).

54. D. J. DesMarais, in The Carbon Cycle and Atmo-spheric CO2: Natural Variations Archean to Present,E. T. Sundquist and W. S. Broecker, Eds. (AmericanGeophysical Union, Washington, DC, 1985), pp.602–611.

55. CH4 is present at 0.1 to 1% of [H2O] in comets [M. J.Mumma, P. R. Weissman, S. A. Stern, in Protostarsand Planets III, E. H. Levy and J. I. Lunine, Eds. (Univ.of Arizona, Tucson, 1993), pp. 1177–1252]. For animpactor mass flux ;5 3 1014g year21 around 4 Ga(11), exogenous CH4 supply provided ,1% of therequired flux.

56. C. F. Chyba, T. C. Owen, W.-H. Ip, in Hazards Due toComets and Asteroids, T. Gehrels, Ed. (Univ. of Ari-zona Press, Tucson, 1994), pp. 9–58.

57. G. A. Dawson, J. Geophys. Res. 82, 3125 (1977); D.L. Baulch et al., J. Phys. Chem. Ref. Data 11 (suppl.1), 327 (1982).

58. R. Stribling and S. L. Miller, Origins Life 17, 261(1987).

59. D. M. Hunten and T. M. Donahue, Annu. Rev. EarthPlanet. Sci. 4, 265 (1976).

60. S. C. Wofsy and M. B. McElroy, J. Geophys. Res. 78,2619 (1973).

61. Integrating the eddy diffusion equation fe 52Kn(z)[]f(z)/]z] for n(z) 5 n(0)exp(2z/H ), where f(z) is[NH3] at altitude z, gives fe 5 2[Kn(z)/H ][f(z) 2 f(0) #(K/H ) f(0)n(z).

62. C. Sagan, B. N. Khare, J. Lewis, in Saturn, T. Gehrelsand M. S. Matthews, Eds. (Univ. of Arizona Press,Tucson, 1984), pp. 788–807.

63. A. Henderson-Sellers and A. W. Schwartz, Nature287, 526 (1980).

64. C. Sagan, J. Theor. Biol. 39, 195 (1973); J. F. Kast-ing, K. J. Zahnle, J. P. Pinto, A. T. Young, Origins Life19, 95 (1989).

65. K. Rages and J. B. Pollack, Icarus 41, 119 (1980).66. While completing this paper, we learned that relat-

ed suggestions were made qualitatively by J. Kast-ing [J. Geophys. Res. 87, 3091 (1982)] and J. Love-lock [The Ages of Gaia (Norton, London, 1988),chap. 4].

67. B. N. Khare et al., Icarus 103, 290 (1993).68. P. Jenniskens, Astron. Astrophys. 272, 465 (1993).69. G. Chlewicki and J. M. Greenberg, Astrophys. J.

365, 230 (1990).70. J. M. Greenberg and J. I. Hage, ibid. 361, 260

(1990).71. H. J. Hagemann, W. Gudat, C. Kunz, J. Opt. Soc.

Am. 65, 742 (1975).72. W. W. Duley, Astrophys. J. 287, 694 (1984).73. E. Bussoletti, L. Colangeli, A. Borghesi, V. Orofino,

Astron. Astrophys. Suppl. Ser. 70, 257 (1987).74. F. Rouleau and P. G. Martin, Astrophys. J. 377, 526

(1991).

75. E. T. Arakawa, S. M. Dolfini, J. C. Ashley, M. W.Williams, Phys. Rev. B 31, 8097 (1985).

76. D. A. Anderson, Philos. Mag. 35, 17 (1977).77. S. C. Lee and C. L. Tien, Proceedings, Eighteenth

Symposium (International) on Combustion (Com-bustion Institute, Pittsburgh, 1981), pp. 1159–1166.

78. W. R. Thompson, B. G. J. T. P. Murray, B. N. Khare,C. Sagan, J. Geophys. Res. 92, 14933 (1987); B. N.Khare, W. R. Thompson, B. G. J. T. P. Murray, C. F.Chyba, C. Sagan, Icarus 79, 350 (1989).

79. This research was supported in part by NASA grantsNAGW-3273, NAGW-1896, and NAGW-1870. Wethank K. J. Zahnle, J. Bada, J. Kasting, W. R.Thompson, B. N. Khare, S. L. Miller, E. E. Salpeter,S. Epstein, J. W. Schopf, D. Layzell, D. Turpin, J. C.G. Walker, S. Soter, J. Goldspiel, P. Gogarten, L. A.Wilson, G. Bowen, C. de Ruiter, R. Hargraves, andG. D. McDonald.

Trichodesmium,a Globally Significant

Marine CyanobacteriumDouglas G. Capone, Jonathan P. Zehr, Hans W. Paerl,

Birgitta Bergman, Edward J. Carpenter

Planktonic marine cyanobacteria of the genus Trichodesmium occur throughout theoligotrophic tropical and subtropical oceans. Their unusual adaptations, from the mo-lecular to the macroscopic level, contribute to their ecological success and biogeo-chemical importance. Trichodesmium fixes nitrogen gas (N2) under fully aerobic con-ditions while photosynthetically evolving oxygen. Its temporal pattern of N2 fixationresults from an endogenous daily cycle that confines N2 fixation to daylight hours.Trichodesmium colonies provide a unique pelagic habitat that supports a complexassemblage of consortial organisms. These colonies often represent a large fraction ofthe plant biomass in tropical, oligotrophic waters and contribute substantially to primaryproduction. N2 fixation by Trichodesmium is likely a major input to the marine and globalnitrogen cycle.

Trichodesmium, a colonial marine cya-nobacterium (1) (Fig. 1), has intrigued nat-uralists, biologists, and mariners for wellover a century (2). These cyanobacteriahave been reported throughout the tropicaland subtropical Atlantic, Pacific, and Indi-an oceans, as well as the Caribbean andSouth China seas (Fig. 2) (3, 4). Moderninterest in Trichodesmium dates to the early1960s with the recognition that the biolog-ical productivity of large expanses of theocean is often limited by the availability ofnitrogen (5) and the observation that Tri-chodesmium is diazotrophic (that is, an N2

fixer). The current focus in assessing theglobal role of the upper ocean in assimilat-ing atmospheric CO2 has elevated the im-portance of quantifying marine N2 fixation.

Although major advances in under-standing the biology of Trichodesmium haverecently occurred on diverse fronts, severalimportant questions remain largely unre-solved: (i) Where does Trichodesmium fit inthe broader scheme of cyanobacterial phy-logeny? (ii) How does Trichodesmium sus-tain simultaneous photosynthetic O2 evolu-tion with nitrogenase activity, and whydoes it fix N2 only during daylight periods?(iii) What physiological, morphological,and behavioral adaptations contribute toTrichodesmium’s ecological success in theoligotrophic marine environment? (iv)What environmental and ecological fac-tors control production and N2 fixation inTrichodesmium in situ, and to what extentdoes it contribute to productivity, nutrientcycling, and trophodynamics in tropicaland subtropical seas? (v) What is the over-

D. G. Capone is at the Chesapeake Biological Laborato-ry, Center for Environmental and Estuarine Studies, Uni-versity of Maryland, Solomons, MD 20688, USA. J. P.Zehr is in the Department of Biology, Rensselaer Poly-technic Institute, Troy, NY 12180, USA. H. W. Paerl is atthe Institute of Marine Sciences, University of North Caro-lina, Morehead City, NC 28557, USA. B. Bergman is inthe Department of Botany, Stockholm University,S-10691 Stockholm, Sweden. E. J. Carpenter is at theMarine Sciences Research Center, State University ofNew York, Stony Brook, NY 11794, USA.

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all importance of Trichodesmium N2 fixa-tion and primary production in the globalmarine N and C cycles?

Molecular and PhysiologicalFeatures

Open-ocean marine N-cycle studies and ba-sic N2 fixation research converged in 1961when Dugdale et al. first identified Tri-chodesmium as a putative light-dependentN2 fixer (6). However, as a nonheterocys-tous N2 fixer, Trichodesmium represented aclear exception to the prevailing dogma (7)and, in the absence of a pure culture orother definitive evidence, its ability to fixN2 was viewed with some caution (8). Acomparison of the DNA sequence of thegene that codes for the Fe protein of nitro-genase (nifH) obtained from natural popu-lations of Trichodesmium with that from aTrichodesmium isolate (9) provided the firstdirect evidence of the cyanobacterial originof the nitrogenase activity associated withTrichodesmium. Later immunolocalizationstudies showed that the nitrogenase proteinoccurs within Trichodesmium cells (10–12).

The diversity among forms of Trichodes-mium with respect to cellular and colonialmorphology has prompted extended debateconcerning its taxonomy and phylogeneticposition within the cyanobacteria. With

the relatively recent isolation and mainte-nance of cultures of the organism (13, 14)and the development of structural and mo-lecular biological approaches to character-izing natural populations, the taxonomicstatus of Trichodesmium populations hasgained firmer footing. Sequence analysis ofnifH and 16S ribosomal DNA (rDNA) hasenabled inference of the taxonomic identityof the Trichodesmium sp. isolates and hasprovided additional bases for distinguishingTrichodesmium species as well as the meansto determine the relative phylogenetic re-lation of Trichodesmium to other bacteriaand cyanobacteria (15, 16). The data areconsistent with differences in trichome di-mensions and colony morphology that hadbeen used in previous taxonomic schemes(15). The nitrogenase DNA sequences offield samples of T. thiebautii and both fieldand culture samples of T. erythraeum werevery similar (98% over 325 nucleotides), incontrast to comparisons of nitrogenase nifHgene sequences between species of othercyanobacterial genera, which are as low as75% similar. Trichodesmium nifH sequencesfrom three species form a deeply branchingcluster within the cyanobacteria (Fig. 3),implying a very early radiation in cyanobac-terial evolution. Although the nifH se-quence indicates that Trichodesmium sp.NIBB 1067 is distantly related to othercyanobacteria, it appears relatively closelyrelated to an oscillatorian on the basis of its16S rDNA sequence (17). The reason for

the differences in phylogenetic associationbased on the two genes is not yet clear butcould involve convergent evolution (innifH) or lateral transfer (of nifH or 16Sribosomal RNA genes). The rather distantrelation of the Trichodesmium nifH gene tothat of other cyanobacteria suggests thatTrichodesmium nifH evolution may be con-strained by the structural requirements foraerobic N2 fixation.

One of the most intriguing aspects ofTrichodesmium biology is the simultaneousoccurrence of N2 fixation and photosynthe-sis. Several hypotheses explaining how Tri-chodesmium might fix N2 aerobically havebeen proposed. First, it is possible that theproperties of Trichodesmium nitrogenase areunique with respect to their resistance to O2inactivation. Second, nitrogenase may betransiently modified to protect it from per-manent O2 deactivation, or it may be con-tinually synthesized to replace protein beinginactivated by O2. Third, there may beintracellular O2-consumptive processes thatmaintain O2 at concentrations compatiblewith N2 fixation. Finally, N2 fixation andphotosynthesis may be spatially segregatedin some manner—for instance, either with-in specific regions in a colony or by celldifferentiation within a trichome—suchthat N2 fixation and photosynthesis are mu-tually exclusive within individual cells.

Mapping of the Trichodesmium nitroge-nase operon and subsequent sequencing ofnitrogenase structural genes demonstratedA

B

Fig. 1. Examples of Trichodesmium colonies. (A)Fusiform or tuft of Trichodesmium culture IMS101; (B) radial or puff colony of T. thiebautii. Col-onies are typically ;2 to 5 mm in length (fusiform)or diameter (radial) and are composed of tens tohundreds of aggregated filaments (trichomes).Each trichome consists of tens to hundreds ofcells (typically ;100); cells are generally 5 to 15mm in diameter but can range up to 50 mm inlength (15) (photos by H. Paerl).

26.5°N

26.5°S

8

7 163

24

9

50

0 60 120 120 60 0180

Fig. 2. Location of process-oriented studies and distribution of Trichodesmium in the world’s oceansbased on maps compiled by Carpenter (3) with reference to surface nutrient and productivity distribu-tions from Berger et al. (85) to exclude areas of coastal upwelling and equatorial divergence. Dashed lineindicates approximate extent of Trichodesmium penetration into subtropical waters. Although Tri-chodesmium is found in waters colder than 20°C, growth and activity are usually restricted to watersabove 20°C (3, 4). See Table 1 for key to study locations.

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that the gene size and organization weresimilar to those of other heterocystous andnonheterocystous cyanobacteria (16). Mod-eling of the three-dimensional structure ofthe Fe protein from the deduced amino acidsequence did not reveal unique or unusualfeatures of the Trichodesmium nitrogenaseFe protein, relative to those of other cya-nobacteria, that might explain a higher O2tolerance (18). Moreover, elevated O2 con-centrations lead rapidly to inhibition of N2fixation, whereas decreased partial pressureof O2 often stimulates activity (19); hence,nitrogenase activity in Trichodesmium is O2-sensitive.

Continuous nitrogenase synthesis, as ob-served in other nonheterocystous cya-nobacteria (7), could replace nitrogenaseinactivated by O2 and thereby provide amechanism for simultaneous N2 fixationand photosynthesis. However, nitrogenaseactivity in Trichodesmium is sustained forseveral hours in the presence of the pro-tein synthesis inhibitor chloramphenicol(19, 20) and well after the disappearanceof nifH messenger RNA (21), which indi-cates a relatively low turnover rate of ni-trogenase protein during the active periodof N2 fixation. In some diazotrophs, nitro-genase may be protected from permanentinactivation by O2 through conformation-al changes or covalent modification (7).The Fe protein of Trichodesmium nitroge-nase can be modified in response to O2stress (19, 22), although the nature of themodification and whether this modifica-tion confers protection have yet to bedetermined.

Experimental evidence exists for the roleof O2 removal processes in maintaining lowintracellular O2 concentrations and therebyfacilitating contemporaneous N2 fixationand photosynthesis. Respiration rates inTrichodesmium appear high relative to othercyanobacteria, resulting in high compensa-

tion points (typically 100 to 200 mmol m22

quanta s21) (23). Several other biochemi-cal or physical means of O2 consumptionhave been suggested (24).

Fogg (25) first offered the provocativehypothesis that a spatial segregation of O2evolution and nitrogenase activity (analo-gous to that between vegetative cells andheterocysts) occurs within Trichodesmiumcolonies. He postulated that photosystem II(PS II)–associated O2 production took placein trichomes near the periphery of the colo-ny, whereas nitrogenase activity was con-fined to the inner portions of the colony thatlacked PS II. Experimental evidence sup-porting this idea was provided (26, 27) soonafter Fogg’s original suggestion. However,other findings suggest that nitrogenase andoxygenic photosynthesis may co-occur incells and that colony integrity is not anabsolute prerequisite for activity (28). Earlysuggestions that Trichodesmium might differ-entiate N2-fixing and photosynthesizingcells along individual trichomes have recent-ly gained support (29). If cellular delegationof activity through differentiation is conclu-sively demonstrated as a general mechanismin Trichodesmium, it would provide an in-valuable model for molecular-level studies ofa simple differentiated system.

The proposed mechanisms summarizedabove are not mutually exclusive. Trichodes-mium most likely uses several strategies topermit nitrogenase activity during photo-synthesis. Other mechanisms that enablethe co-occurrence of nitrogenase activityand oxygenic photosynthesis in this organ-ism may yet be identified. Regardless of howTrichodesmium is able to fix N2 in the light,it is equally curious that in natural popula-tions N2 fixation occurs only during the dayand is not performed during the night, as ischaracteristic for other nonheterocystouscyanobacteria (7). Saino and Hattori (30)first observed that nitrogenase was only ac-

tive in samples of Trichodesmium collectedduring daylight hours: Samples collected atnight were incapable of N2 fixation underartificial light. They suggested, 8 years be-fore circadian rhythms were identified inany prokaryote (31), that the daily cycle ofN2 fixation in Trichodesmium might be at-tributable to an endogenous rhythm. Re-search into the dynamics of the nitrogenasepool and nifH transcription in natural pop-ulations revealed a dynamic daily cycle ofsynthesis, activity, and degradation, directlycoupled to the light cycle (19–21), thusproviding a mechanistic explanation for theoriginal observations.

A primary criterion for establishing theendogenous nature of N2 fixation in Tri-chodesmium—its persistence over several cy-cles in constant light—has recently beenprovided in Trichodesmium sp. IMS 101(32). The circadian clock, which appears tobe set by illumination patterns, is likely tobe of adaptive significance in ensuring thatsynthesis is induced in anticipation of thelight period, thus optimizing the efficiencyof light-driven N2 fixation in the relativelystable environment of tropical and subtrop-ical seas. This is the first endogenousrhythm to be confirmed in a prokaryoteother than a coccoid cyanobacterium (31).

Ecology

Trichodesmium primarily inhabits surfacewaters of oligotrophic, tropical, and sub-tropical oceans, and is encountered inhigh abundance in western boundary cur-rents (for example, the Gulf Stream, Ku-roshio), in tropical portions of the centralgyres, and in several ocean margin seas (3)(Fig. 2). The water column of these envi-ronments is generally very stable, with theupper mixed layer often around 100 m.This zone is characterized by low nutrientconcentrations, very clear waters, anddeep light penetration.

In an environment where the densitiesof microorganisms are very low and thewaters highly transparent, Trichodesmium isunusual in that it is visually prominent,especially during surface blooms. The ob-served abundance of Trichodesmium in nu-trient-depleted waters prompts the questionof how it is uniquely adapted to this envi-ronment. Characteristics of Trichodesmiumthat appear to contribute to its success inthe oligotrophic open ocean include its ca-pability to fix N2; its natural buoyancy,which positions it in the upper water col-umn; a photosynthetic apparatus adapted toa high-light regime; and a relatively lowgrowth rate, which, coupled with a lack ofmajor grazers, allows it to maintain relative-ly high biomass.

Despite its ability to fix N2 and its hab-

Fig. 3. Evolutionary relationof the genus Trichodesmiumto other diazotrophs on thebasis of nifH amino acid se-quence data. The evolution-ary distances between de-duced amino acid sequenc-es were used to create aphenogram using the neigh-bor-joining method ofPHYLIP (96). The Fe proteingene (nifH) of Trichodes-mium was amplified fromcultures by means of thepolymerase chain reaction(9). Trichodesmium nifH sequences cluster relatively distantly from other cyanobacterial genera, imply-ing that the Trichodesmium genus diverged early in evolution. The nifH DNA sequences obtained fromfield collections of T. erythraeum and T. thiebautii are very similar (98%) and could be distinguished byonly a few signature nucleotides (short distances among Trichodesmium species nifH sequencescannot be distinguished here).

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itation in the high-light environment of thenear surface, Trichodesmium has growthrates that are low relative to those of manyeukaryotic phytoplankton (4). Even in cul-ture, the doubling times of Trichodesmiumare slow (;3 to 5 days), which suggests thata relatively low growth rate may in fact bean adaptation for exploiting the high-ener-gy but low-nutrient conditions of the oligo-trophic oceans (33). Because of its diazotro-phic capacity, Trichodesmium growth ratesare presumably limited by the availability ofnon-N nutrients. Field and laboratory datasuggest that Fe is a key factor limiting Tri-chodesmium N2 fixation and growth rates(34–36).

A key characteristic of Trichodesmium isthe presence of gas vesicles, which providebuoyancy (37) and help maintain Trichodes-mium populations in the upper surface wa-ters. As has been noted in other planktoniccyanobacteria, the buoyancy of Trichodes-mium is a dynamic property: A daily cycle ofrising and sinking of colonies is often ob-served, and this may be a result of cellballasting through the progressive increaseof relatively dense carbohydrate and proteinaccumulating from photosynthesis throughthe day (38).

Wind stress at the surface affects thequalitative distribution of Trichodesmiumpopulations. Relatively high and steadywinds of the Trade Wind belts mix plank-ton populations throughout the upper eu-photic zone, but natural buoyancy counter-acts mixing to the bottom of the mixedlayer; population densities are generallygreatest at relatively shallow depths (20 to40 m) in the upper water column (3, 4)(Fig. 4).

Localization of the population in thehigh-irradiance upper water column is anadaptation that may provide a solution tothe constraint of high compensation points(23) (equivalent to compensation depths of;50 to 70 m in oligotrophic waters) andthe added energetic demands of N2 fixation.Pigment composition and photosyntheticparameters indicate a photosystem adaptedfor both maximum efficiency and photopro-

tection at high light (4, 39). Moreover, Feenters the open ocean primarily throughatmospheric deposition, and location ofthese organisms in the upper water columnmay be advantageous with respect to Feacquisition (35).

When wind stress is low for an extendedperiod, the intrinsic buoyancy of Trichodes-mium can have striking consequences, lead-ing to the development of extensive surfaceblooms or “red tides” (3, 39, 40). Thesephenomena, which range in actual colorfrom yellow to brown, are easily observed bysatellite (Fig. 5) (41, 42). The accumula-tion of the population at the surface for anextended period may result in photoinhibi-tion (43) and, possibly, photooxidativedamage (3, 4, 39); however, bloom organ-isms appear to be metabolically active,growing (40, 41), and relatively resistant tophotoinduced damage (39).

Although physical processes most oftendominate oceanic plankton distributions,large surface accumulations of Trichodes-mium may affect the bulk physical andchemical properties of surface waters. Me-soscale sea-surface features of the extentand intensity characteristic of some Tri-chodesmium blooms likely modify light pen-etration and the quality of the in situ lightfield (43) as well as heat and gas transferacross the ocean-atmosphere interface (44).Organic and inorganic nutrients accumu-late during blooms and can affect subse-quent phytoplankton succession and pro-ductivity (40).

Ecologically, Trichodesmium affects thestructure and function of the oligotrophicocean by contributing to its productivityand trophodynamics, as well as by provid-ing a unique pelagic habitat. Upon closeinspection, many colonies reveal a micro-cosm including bacteria, other cyanobac-teria, protozoa, fungi, hydrozoans, andcopepods (45). Nitrogen and carbon fixedby Trichodesmium enters the food web, butnot necessarily through increased C and Nflux into classical food chains. Some spe-cies of Trichodesmium produce a toxin thatdeters grazing by calanoid and cyclopoid

copepods, considered to be the major graz-ers in these systems (46). A specializedgroup of harpacticoid copepods appearadapted to the toxin and are capable ofdirectly grazing Trichodesmium (47), al-though their quantitative role in the con-sumption of Trichodesmium is unknown.Although there are anecdotal reports ofthe presence of Trichodesmium in the gutsof gelatinous zooplankton and fish (4),this does not appear to be a quantitativelyimportant fate for Trichodesmium biomass,and much of the C and N fixed by Tri-chodesmium likely enters upper trophiclevels by other pathways. In natural pop-ulations of Trichodesmium, a large propor-tion of recently fixed N is released asorganic N (48). Besides providing a meansof exchange between N2-fixing and non–N2-fixing cells in the colonies, this exu-dation may be an important source of Cand N for microbes inhabiting the colo-nies (49) or free-living in the water col-umn, and ultimately may be a source ofinorganic N for phytoplankton.

Biogeochemistry

With its cosmopolitan distributionthroughout much of the oligotrophic trop-ical and subtropical oceans (3) and its ca-pacity to form extensive surface blooms(40–42), Trichodesmium may be one of themost globally important cyanobacteria andphytoplankton. However, there is insuffi-cient quantitative information to substanti-ate this inference, possibly because of theunique problems associated with assessingthe biomass and productivity of Trichodes-mium by traditional methods as well as theinherent difficulty in sampling ephemeralblooms (50). As a result, the contributionof Trichodesmium to marine C and N inputhas generally been considered relativelysmall (51). Recent data, however, suggestotherwise.

Accurately quantifying N2 fixation inthe seas has direct bearing on our under-standing of C flux in the tropical oceans.According to the “new production” concept

Fig. 4. Average depth distributionsof trichomes of T. thiebautii andvolumetric N2 fixation (A), and chlo-rophyll a (Chl a) biomass and prima-ry productivity relative to micro-plankton (B), for a series of stationstaken in the southwestern tropicalNorth Atlantic in May 1994 (Fig. 2,map code 9) (97). Bars indicate6SE of the mean.

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(52), in steady-state marine systems theamount of organic N removed from thesystem (that is, the euphotic zone) shouldequal external (“new”) N inputs enteringthat system. Although N2 fixation is a po-tential source of new N, NO3

2 from depthis generally considered the quantitativelydominant source of new N in most open-ocean systems (52). Areal rates of totalproduction in tropical oligotrophic regionsare typically very low (53), and the relativeamount of production dependent on new N(that is, “new” production) is thought to besmall; nonetheless, the export of organic Nand C from the upper water column ofoligotrophic regions is important in the ma-rine C and N budgets because the total areainvolved is vast (53).

The increase in atmospheric CO2 hasprovided added impetus to the quest toquantify oceanic primary production and itsrate of removal to depth (54). N2 fixationand vertical NO3

– flux from depth havedifferent potentials for supporting primaryproduction and effecting net removal ofCO2 from the atmosphere. Vertical NO3

2

flux occurs with a concurrent upward flux ofCO2 and PO4

3– from depth, often close tothe stoichiometric requirement of phyto-plankton. Thus, relative to N2 fixation,NO3

– derived from depth has limited capac-ity for effecting net removal of atmosphericCO2. N2 fixation represents a source of newN entering the system that can account fora net sequestering of atmospheric CO2 into

export production (55).Recently, several independent lines of

evidence have prompted speculation thatmarine N2 fixation has been severely un-derestimated and may play a larger role inglobal C and N fluxes. Large imbalances inthe estimated N budgets for the North At-lantic, central North Pacific, Indian, andglobal oceans have prompted speculation asto unknown or poorly quantified N inputs,and N2 fixation may provide the missingsource of N (56).

For nonupwelling regions of the tropicaland subtropical North Atlantic, the verticalflux of NO3

–, assumed to represent the bulkof the influx of new N into the euphoticzone (52), is estimated to range from 30 to150 mmol N m22 day21 (57). However,such estimates of new N inputs are ofteninsufficient to satisfy the calculated demandfor new N and organic export from theeuphotic zone (57), which suggests the ex-istence of sources of combined N in addi-tion to vertical NO3

– flux. Other biogeo-chemical approaches also reveal N demandin, or losses from, the upper water columnexceeding current estimates of N inputs,further indicating unquantified inputs (58).

Independent evidence that N2 fixationmay be of greater relevance than currentlybelieved derives from studies of the naturalisotopic abundance of 15N in surface partic-ulate organic nitrogen (PON). Trichodes-

mium has a low d15N, typical of N2-fixingorganisms (59). Relative to the isotopiccomposition of PON in more nutrient-richareas, the d15N of suspended particles andzooplankton in surface waters of the west-ern and central tropical Pacific and in theCaribbean is often low and inversely relatedto the abundance of Trichodesmium. Thissuggests that the isotopically light PON inthese waters is a result of N2 fixation byTrichodesmium (59).

Relative to conventional assessments ofplanktonic standing crop, primary produc-tivity, and inorganic N uptake, there arefew direct estimates of Trichodesmium bio-mass and its rate of N2 fixation (50) (Table1) and even fewer estimates of its contribu-tion to primary production. The most com-prehensive studies to date have been madein the subtropical Sargasso Sea and in thetropical Caribbean and far western Pacific(Kuroshio, South and East China Sea) (Fig.2), ocean margin regions that may not berepresentative of the tropical oceans. Morelimited data are available for the tropicalNorth Atlantic and the vicinity of Hawaii.Virtually no information exists for largeexpanses of the tropical oceans, particularlyin the Southern Hemisphere. Nonetheless,several of the studies conducted over thepast two decades have provided direct evi-dence of the importance of Trichodesmiumrelative to other phytoplankton (60). With

Table 1. Summary of direct areal estimates of N2 fixation, as originally reported or as derived. Studiesbased on C2H2 reduction determinations of N2 fixation used a 3:1 conversion from C2H2 reduced to N2fixed, unless otherwise noted. N, number of discrete observations.

Location

Areal estimates

Mapcode

Refer-ence

DateAverage 6 SE(mmol N m22

day21)N

Subtropical28°N, 155°W (North Pacific) Aug 73 33 1 1 (95)27° to 34°N (NW Sargasso Sea) Sep–Oct 73 1.4* 6 0.47 9 2 (93)22° to 36°N (Sargasso Sea) Aug 74 6.2* 6 4.0 7 3 (92)22° to 23°N (Caribbean

passages)Feb–Mar 74,

Aug 744.2* 6 4.0 10 3 (92)

30°N (Atlantic transect) May–Jun 75 0.29* 6 0.13 5 4 (94)Tropical

0° to 24°N, 45° to 66°W (SWNorth Atlantic)

Fall 64 41† 6 7.6 19 5 (65)

Spring 65 108† 6 24 15 5 (65)21°N, 159°W (North Pacific) Oct, Dec 72 134 2 6a (62)

12° to 22°N (Caribbean) Feb–Mar 74,Aug 74

77* 6 9.7 12 3 (92)

10° to 25°N (SE East China Sea) Summer 77 126 32 7 (63)23°N, 158°W (North Pacific) Jun 90,

Feb 9185 2 6b (89)

7° to 10°N (Arabian Sea) May 95 35 6 7.4 9 8 (81)14° to 22°N (SW North Atlantic) May 94 73 6 22 12 9 (64)NE Caribbean May 94 278 6 129 3 9 (64)Tropical, grand average 106 6 24

*Data as originally presented using a 6.3 :1 conversion ratio. †Data based on direct 15N2 uptake. Average ratesfrom 0 and 15 m are assumed over the top 20 m and have been increased by 50% to account for activity below 20 m,on the basis of data for the region from Fig. 4 and (64).

A

B

118°E 120°E

20°S

20°S

Fig. 5. (A) Image of chlorophyll and (B) relativeTrichodesmium abundance derived from a coast-al zone color scanner (CZCS) image of the north-western coast of Australia in the vicinity of theDampier Archipelago and confirmed by contem-poraneous sea-truth data [adapted from (42)]. Aprotocol based on reflectivity and absorption at550 nm was used. Chlorophyll is reported as de-tected by CZCS, with lowest to highest chloro-phyll a concentrations ranging from purple (,0.05mg m23) to red (.3.0 mg m23). For Trichodes-mium, dark colors indicate its absence; lighter col-ors (light blue through orange) indicate its pres-ence. Differences in color represent varying re-sponses to the protocol, not necessarily differenc-es in Trichodesmium concentration.

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respect to subtropical waters such as theSargasso Sea, previous studies have detectedappreciable populations of Trichodesmiumonly for limited periods during the summer,and the calculated contributions of Tri-chodesmium to C and N input were relative-ly small (Table 1) (61).

In tropical studies, despite low growthrates, the relatively high biomass of Tri-chodesmium in these regions implies that itmay account for a quantitatively importantinput of organic C, and data from severalstudies have provided support for this idea(60) (Fig. 4). Not surprisingly, the mostdata exist for estimates of N2 fixation (Ta-ble 1). The average rate of N2 fixationreported for a suite of stations throughoutthe Caribbean Sea was 77 mmol N m22

day21. One station in the tropical NorthPacific yielded an N2 fixation rate of 134mmol N m22 day21 (62–65). We calculated126 mmol N m22 day21 on the basis of data(63) for the China Sea and Kuroshio cur-rent near the southern tip of Japan (Table1). Our recent results from the open tropi-cal Atlantic Ocean (Fig. 4) are generallyconsistent with earlier data from this andother tropical regions (Table 1). Scaling theaverage (nonbloom) tropical rate of 106mmol N m22 day21 to tropical oligotrophicwaters that cover about 150 3 106 km2 (53)yields an annual input of ;80 Tg N, con-siderably greater than previous estimates ofpelagic N2 fixation input (3, 40).

These estimates of N input through N2fixation in tropical areas are roughly equiv-alent to the estimates of vertical NO3

2 fluxgiven above and suggest that these twoprocesses are of comparable importance innew N input. Moreover, recent evidenceindicates that actual eddy diffusivities are atthe lower range of those values currentlyassumed (66). If this is correct, it wouldlower the estimate of N input by verticalflux of NO3

2 and correspondingly increasethe relative importance of upper water col-umn N2 fixation to overall N input. Theextreme depth of the euphotic zone in theoligotrophic open ocean (.100 m), and theabsence of measurable NO3

– throughoutmuch of the upper water column of thehighly oligotrophic tropical areas, imply anuncoupling between the reservoirs of NO3

–

at depth and productivity in the near sur-face. Any NO3

2 that penetrates up fromdepth is likely to be assimilated at the chlo-rophyll maximum, typically located nearthe 1% light level near the nitracline.

Although numerous blooms have beendocumented and their spatial extent andbiomass density have been estimated (40–42), the effect of a given bloom on C and Ninput has been determined only on a fewoccasions. For three studies that directlymeasured N2 fixation in surface blooms, the

input of N in the bloom was about threetimes that occurring throughout the rest ofthe water column (67).

Taken together, these observationsstrongly indicate that N2 fixation is an im-portant component of the marine N budgetthat needs to be considered in basin-scalestudies of N cycling and in calculations ofglobal N budgets. In addition to N2 fixa-tion, other potentially significant “new” Ninputs, such as atmospheric deposition ofdissolved inorganic and organic N, havebeen poorly quantified or ignored in thepast (68). More accurate assessment of theircontributions to the oceanic N cycle willfurther help to rectify current discrepanciesin basin-scale and oceanic N budgets.

Future Research and Prospects

Research efforts from a molecular to a glob-al perspective provide a new basis for un-derstanding the biology and ecology of Tri-chodesmium and inferring its role in globalbiogeochemical cycles. Physiological, ge-netic, and immunological evidence haveconfirmed Trichodesmium, rather than asso-ciated microorganisms, as the main N2 fixerin its colonial aggregates. Trichodesmiumfixes N2 aerobically, possibly in the samecell and at the same time as it evolves O2through photosynthesis, making it a uniqueand valuable model organism for the studyof N2 fixation in photosynthetic organisms.As one of the few prokaryotic systems iden-tified as having an endogenous rhythm, Tri-chodesmium has a relative biochemical sim-plicity that makes it an attractive system foridentifying the genetic and physiologicalfactors regulating components of its biolog-ical clock.

Multiple characteristics have been iden-tified that contribute to the ability of Tri-chodesmium to fix N2 aerobically, but a sin-gle critical component that allows simulta-neous photosynthesis and nitrogenase ac-tivity of nitrogenase has yet to be found,and seemingly conflicting results need to bereconciled. The dichotomy in growth rateestimates based on C or N in natural pop-ulations needs also to be resolved. Theavailability of robust cultures will greatlyimprove our knowledge of the biochemicalfunctioning of these unusual diazotrophs.

A variety of recognized morphologicaland functional features demonstrate Tri-chodesmium to be well adapted to the N-poor oligotrophic ocean environment. Ofparticular ecological relevance may be itscyclic patterns of vertical migration, N2fixation, and photoprotective processes.However, we have yet to develop an inte-grative physiological model directly linkingthese ecological behaviors in cultures or innatural populations. More rigorous defini-

tion of the interplay of physical and chem-ical limiting factors will provide importantconstraints for modeling and predicting thein situ dynamics of these populations.

The accumulating evidence strongly in-dicates a much more important role of Tri-chodesmium in oceanic biogeochemistrythan it is currently afforded. Synoptic esti-mates of the areal and temporal extent ofTrichodesmium populations and, particular-ly, blooms by new and planned satellitesensors [for example, the ocean color andtemperature scanner (OCTS) and the sea-viewing wide field-of-view sensor (Sea-WiFS)], coupled with the development ofin situ methods of enumerating these pop-ulations (for example, fluorescence-basedoptical plankton counters and laser-inducedfluorescence imagers), will further refineand advance our knowledge of occurrencein tropical and subtropical seas. This in-formation will be useful in determiningthe potential role of blooms in the modi-fication of system-scale features such asheat, material flux, and albedo. The morecomprehensive database on Trichodesmiumpopulation biomass and distribution de-rived from satellite studies, along with ex-panded direct information on its in situcontributions to C and N cycling, will alsoallow for more precise extrapolation of itsoceanic contributions to new N inputs aswell as the inclusion of Trichodesmium asan explicit component of large-scale mod-eling of oceanic productivity.

Variations in oceanic productivity overglacial-interglacial time scales have beendirectly related to subtle variations in theextent of denitrification and cyanobacterial(that is, Trichodesmium) N2 fixation in thesea (69). The possibility exists for use ofnucleotide probes of nifH to examine paleo-ecological trends of Trichodesmium popula-tions in oceanic sediment cores (70). De-veloping a firm understanding of the dy-namics and controls of Trichodesmium pop-ulations in the contemporaneous ocean,along with information on its populationtrends over geological time scales, will leadto important new insights about controls ofoceanic productivity.

REFERENCES AND NOTES___________________________

1. The genus Trichodesmium is composed of a smallnumber of marine cyanobacterial species within theorder Oscillatoriales and was first described byEhrenberg (71). As the name Trichodesmium implies(from the Greek, tricho: hair, desmos: chain), they arefilamentous. Trichodesmium species are most oftenobserved as fusiform or spherical colonies, althoughthey also occur as single trichomes (72, 73). Theynotably lack heterocysts, the morphologically andbiochemically differentiated cells specialized for fix-ing N2, which characterize filamentous cyanobacte-ria of the orders Nostocales and Stigonematales [R.W. Castenholz and J. B. Waterbury, in Bergey’sManual of Determinative Bacteriology, J. T. Staley,

SCIENCE z VOL. 276 z 23 MAY 1997 z www.sciencemag.org1226

Ed. ( Williams and Wilkins, Baltimore, 1989), pp.1710–1727].

2. The Red Sea obtained its name from the colorationimparted by T. erythraeum (71) [H. J. Carter, Ann.Mag. Nat. Hist. 3, 182 (1863)]. Darwin, while aboardthe H.M.S. Beagle [C. Darwin, Journal of Researchesinto the Natural History and Geology of the CountriesVisited During the Voyages of the H.M.S. BeagleRound the World Under the Command of Capt. FitzRoy, R.N. (Clowes, London, 1845)], as well as Cap-tain J. Cook and his naturalist, J. Banks, aboard theH.M.S. Endeavour, each commented on Trichodes-mium’s appearance and abundance in tropical seas[J. C. Beaglehole, Ed., The Endeavor Journals ofJoseph Banks 1768–1771 (Angus and Robertson,Sydney, Australia, 1962), vol. II: The Journals of Cap-tain James Cook on His Voyages of Discovery, vol. I:The Voyage of the Endeavor 1768–1771 (Cam-bridge Univ. Press, Cambridge, 1955), pp.404–405]. Because of its buoyancy and tendency toform colonies, it is easily observed near the sea sur-face by the naked eye. The colonies, long referred toby mariners as “sea sawdust,” aggregate at the sur-face during periods of low wind stress to form con-spicuous surface slicks or blooms.

3. E. J. Carpenter, in Nitrogen in the Marine Environ-ment, E. J. Carpenter and D. G. Capone, Eds. (Aca-demic Press, New York, 1983), pp. 65–103.

4. iiii, Mar. Biol. Lett. 4, 69 (1983); J. R. Gallon etal., Arch. Hydrobiol. Suppl. 117, 215 (1996).

5. See (74) for a summary of historical data on N limita-tion. Iron (Fe) availability has been shown to limit thegrowth of phytoplankton in some oceanic regionscharacterized as high nutrient, low chlorophyll(HNLC) [J. Martin et al., Nature 371, 123 (1994)].

6. R. C. Dugdale, R. D. W. Menzel, J. H. Ryther, DeepSea Res. 7, 297 (1961). In parallel with the expansionof interest in the marine N cycle, a revolution in fun-damental research on N2 fixation also occurred [R.W. F. Hardy and U. D. Havelka, Science 188, 633(1975)].

7. At the time, nitrogenase was recognized to be inac-tivated by O2. It was generally held that cyanobacte-ria protected nitrogenase from O2 evolved throughphotosynthesis by spatial segregation of nitrogenaseinto specialized cells named heterocysts, whichlacked PS II and did not evolve O2. Nonheterocys-tous forms were thought to be incapable of N2 fixa-tion. After 1970, several other nonheterocystouscyanobacteria were found to have nitrogenase; how-ever, unlike Trichodesmium, their nitrogenase wasonly active under conditions of reduced O2 concen-tration, or by temporally segregating N2 fixation tononphotosynthetic periods (that is, at night) [P. Fay etal., Nature 220, 810 (1968); P. Fay, Microbiol. Rev.56, 340 (1992); J. R. Gallon, New Phytol. 122, 571(1992); B. Bergman, J. R. Gallon, A. N. Rai, L. Stal,FEMS Microbiol. Rev. 19, 139 (1997)].

8. G. E. Fogg, Ecol. Bull. 26, 11 (1978); B. F. Taylor etal., Arch. Microbiol. 88, 205 (1973).

9. J. P. Zehr and L. A. McReynolds, Appl. Environ.Microbiol. 55, 2522 (1989); J. P. Zehr et al., ibid. 56,3527 (1990).

10. H. W. Paerl, J. C. Priscu, D. L. Brawner, ibid. 55,2965 (1989).

11. B. Bergman and E. J. Carpenter, J. Phycol. 27, 158(1991).

12. C. Fredriksson and B. Bergman, Microbiology 141,2471 (1995).

13. Two cultures are currently available, both apparentlystrains of T. erythraeum [J. P. Zehr, S. Braun, Y.Chen, M. T. Mellon, J. Exp. Mar. Biol. Ecol. 203, 158(1996)]: strain NIBB 1067, isolated by K. Ohki and Y.Fujita [Mar. Ecol. Prog. Ser. 7, 185 (1982)], and Tri-chodesmium IMS 101, isolated by Prufert-Bebout etal. (14). The former has been resistant to transplan-tation, whereas the latter has been disseminated toseveral laboratories. Additional isolates may now beavailable.

14. L. Prufert-Bebout, H. W. Paerl, C. Lassen, Appl. En-viron. Microbiol. 59, 1367 (1993).

15. The abundance and location of intracellular struc-tures such as gas vacuoles, as well as the form andsize of cells and of aggregates or colonies, variessubstantially; taxonomic identifications are based on

such characteristics [K. Anagnostides and J. Kom-arek, Arch. Hydrobiol. Suppl. 80, 327 (1988)]. De-tailed cytomorphological analysis of natural popula-tions of Trichodesmium in the Caribbean and Sar-gasso seas, which concluded that there are severaldistinct morphotypes of Trichodesmium [S. Jansonet al., J. Phycol. 31, 463 (1995)], is supported byboth nifH (16) [J. Ben-Porath, E. J. Carpenter, J. P.Zehr, ibid. 29, 806 (1993)] and 16S rDNA (S. Janson,K. Virgin, B. Bergman, S. Giovannoni, in preparation)sequence analysis.

16. J. P. Zehr, K. Ohki, Y. Fujita, J. Bacteriol. 173, 7055(1991); G. Sroga, U. Landegren, B. Bergman, M.Lagerstrom-Fermer, FEMS Microbiol. Lett. 136, 137(1996).

17. A. Wilmotte, J.-M. Neefs, R. De Wachter, Microbiol-ogy 140, 2159 (1994).

18. J. P. Zehr, D. Harris, B. Dominici, J. Salerno, FEMSMicrobiol. Lett., in press.

19. J. P. Zehr, M. Wyman, V. Miller, L. Duguay, D. G.Capone, Appl. Environ. Microbiol. 59, 669 (1993).

20. D. G. Capone, J. M. O’Neil, J. Zehr, E. J. Carpenter,ibid. 56, 3532 (1990).

21. M. Wyman, J. P. Zehr, D. G. Capone, ibid. 62, 1073(1996).

22. K. Ohki, J. P. Zehr, Y. Fujita, J. Gen. Microbiol. 138,2679 (1992).

23. E. J. Carpenter and T. Roenneberg, Mar. Ecol. Prog.Ser. 118, 267 (1995); (75, 76). The compensationpoint is the light intensity at which photosynthetic O2production equals respiration.

24. These include the Mehler reaction [photoreduction ofO2 by PS I and a source of adenosine triphosphate(76)]; uptake hydrogenase [S. Saino and A. Hattori,Mar. Biol. (Berlin) 70, 251 (1982)]; elevated cyto-chrome oxidase [B. Bergman, P. J. A. Siddiqui, E.J. Carpenter, G. A. Peschek, Appl. Environ. Micro-biol. 59, 3239 (1993)]; photorespiration [W. Li, H.Glover, I. Morris, Limnol. Oceanogr. 25, 447 (1980)];and superoxide dismutase activity (K. Cunninghamand D. G. Capone, in preparation).

25. G. E. Fogg, in Algal Physiology and Biochemistry, W.D. P. Stewart, Ed. (Blackwell, Oxford, 1974), pp.650–682.

26. Autoradiographic localization of photosynthetic14CO2 incorporation in both natural and cultured Tri-chodesmium provided evidence for spatial segrega-tion in colonies [E. J. Carpenter and C. C. Price IV,Science 191, 1278 (1976); H. W. Paerl and P. T.Bland, Appl. Environ. Microbiol. 43, 218 (1982); H.W. Paerl, J. Phycol. 30, 790 (1994)]. Moreover, mi-croelectrode measurements of small-scale O2 gradi-ents in actively photosynthesizing Trichodesmiumcolonies at times revealed distinct internal hypoxic oranoxic microzones (27) [H. W. Paerl and B. M. Be-bout, Science 241, 442 (1988)].

27. H. W. Paerl and B. M. Bebout, in Marine PelagicCyanobacteria: Trichodesmium and Other Dia-zotrophs, E. J. Carpenter, D. G. Capone, J. Rueter,Eds. (Kluwer Academic, Dordrecht, Netherlands,1992), pp. 43–59.

28. The fluorescence signature of PS II light-harvestingpigments was found in all cells [E. J. Carpenter et al.,Mar. Ecol. Prog. Ser. 65, 151 (1990)]; nitrogenaseoccurs with carboxysomes, ribulose-1,5-bisphos-phate carboxylase-oxygenase (RuBisCO), and phy-coerythrin in the same cell (77, 78). Paerl et al. (10)detected nitrogenase by immunolocalization in ahigh proportion of cells within the colonies they ex-amined; more recent studies of several species ofTrichodesmium species collected in various watersconsistently found that nitrogenase is synthesized ina smaller subset (10 to 40%) of cells, randomlyspread through the colony (11, 12, 77). Freetrichomes occur in nature [for example, (73)], al-though the specific rate of nitrogenase activity asso-ciated with free filaments in situ may be considerablyless than in intact colonies (72). Colonies disaggre-gated under N2 and returned to aerobic conditions,or colonies disaggregated under air and assayedunder N2, exhibited nitrogenase activity comparableto whole colonies (D. G. Capone et al., in prepara-tion). Cultures of Trichodesmium growing aerobicallyas individual trichomes fix N2 at relatively high rates(14) [K. Ohki and Y. Fujita, Mar. Biol. (Berlin) 98, 111

(1988)]. However, cultures that exhibit concurrentnitrogenase activity with photosynthesis in single fil-aments (13, 14) are grown at present at subsaturat-ing light levels, which may allow for cellular respira-tion to maintain sufficiently low O2 concentrations fornitrogenase activity.

29. Bryceson and Fay (79) first suggested spatial segre-gation of nitrogenase activity and photosynthesis incells along a trichome, noting the absence of car-boxysomes (sites of RuBisCO) in three to five zonesalong trichomes of T. erythraeum, which also hadhigher reducing potentials. Recent immunolocaliza-tion studies of several species of field-collected Tri-chodesmium (78) [C. Fredriksson and B. Bergman,Protoplasma 197, 76 (1997)] and the Trichodes-mium IMS 101 isolate (C. Fredriksson, H. Paerl, B.Bergman, in preparation) reported that within individ-ual trichomes, nitrogenase is localized along eachtrichome in subsets of several cells with distinct ul-trastructural features.

30. T. Saino and A. Hattori, Deep Sea Res. 25, 1259(1978). Subsequent examination of nitrogenase pro-tein dynamics in field populations found active pro-tein only present in the cells during the day (20).Nitrogenase synthesis was initiated in the morning,and activity commenced with the onset of photosyn-thesis. The daily cycle of synthesis is regulated at thelevel of transcription, as nifH transcripts were detect-ed before the beginning of the light period precedingthe appearance of the protein and were present onlyuntil midday in natural populations and in Trichodes-mium sp. IMS 101; this indicated that the diel cycle ofsynthesis was cued by a factor other than light (21).Moreover, there appears to be a subcellular processthat results in cessation of activity during the lateafternoon. The extant Fe protein component of nitro-genase is converted to an inactive form through anas yet uncharacterized posttranslational proteinmodification mechanism and “turned off.” Immuno-localization studies confirmed that nitrogenase wassynthesized and degraded over the day-night cycle(12).

31. For N2 fixation in a coccoid cyanobacterium, see N.Groebblaar, T. C. Huang, H. Y. Lin, T. J. Chow,FEMS Microbiol. Lett. 37, 173 (1986). Circadianclocks have also been found in association with pho-tosynthetic genes of another Synechococcus strain[for example, T. Kondo et al., Science 266, 1233(1994)]. L. Stal and W. Krumbein [Arch. Microbiol.143, 67 (1983)] reported persistence of N2 fixation inconstant light for one cycle in a filamentous benthicoscillatorian that normally confines nitrogenase ac-tivity to dark periods.

32. Y. Chen, J. P. Zehr, M. Mellon, J. Phycol. 32, 96(1996). A daily cycle of nitrogenase induction andactivity persisted for up to six cycles in constant lightwith a free-running clock of about 26 hours. A dielvariation in the compensation point has also beenreported; this may be driven by an endogenousrhythm in respiration, because photosynthetic pa-rameters do not exhibit a comparable daily cycleunder continuous illumination (75).

33. C. S. Reynolds, in Molecular Ecology of Aquatic Mi-crobes, I. Joint, Ed. (NATO ASI Series, vol. G 38;Springer-Verlag, Berlin, 1995), pp. 115–132. Lowgrowth rates and high biomass are common strate-gies for algae inhabiting physically stable, nutrient-poor environments. Interestingly, growth rates ofnatural populations of Trichodesmium based on CO2assimilation are generally somewhat more rapid thanthose based on N2 uptake (3, 4).

34. It had been speculated that molybdenum was a pri-mary limiting factor for Trichodesmium [R. W.Howarth and J. J. Cole, Science 229, 653 (1985)],although no substantial evidence has been provided.In contrast, because nitrogenase is an Fe-rich pro-tein, the cell quota for Fe of Trichodesmium is con-siderably higher than in other phytoplankton (35).Trichodesmium is capable of rapid Fe uptake, al-though it does not appear to produce siderophores(35). Fe additions can stimulate Trichodesmiumgrowth and activity [J. G. Rueter et al., J. Phycol. 26,30 (1990); H. W. Paerl, L. Prufert-Bebout, C. Guo,Appl. Environ. Microbiol. 60, 1044 (1994)]. Tri-chodesmium growth in situ also requires phospho-

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rus, and the source of its phosphorus requirementhas been the focus of some attention (4, 36).

35. J. G. Rueter et al., in (27), pp. 289–306.36. D. M. Karl et al., ibid., pp. 219–237.37. C. VanBaalen and R. M. Brown, Arch. Microbiol. 69,

79 (1969); A. Walsby, Br. Phycol. J. 13, 103 (1978);E. Gantt et al., Protoplasma 119, 188 (1984). Tri-chodesmium gas vesicles, which can survive pres-sures at depth in excess of 200 m, have the highestcollapse pressure of any cyanobacterial vesicles yetexamined.

38. T. Villareal and E. J. Carpenter, Limnol. Oceanogr.35, 1832 (1990); K. M. Romans, E. J. Carpenter, B.Bergman, J. Phycol. 30, 935 (1994).

39. M. Lewis, O. Ulloa, T. Platt, Limnol. Oceanogr. 33, 92(1988). Phycobiliproteins, which have a high efficien-cy of transfer of light energy, appear to be the prima-ry light-harvesting pigments of photosynthesis (4) [S.Shimura and Y. Fujita, Mar. Biol. (Berlin) 31, 121(1975)]. Trichodesmium also has relatively high con-centrations of carotenoids (3, 4) [K. Hogetsu and M.F. Watanabe, in Studies on the Community of MarinePelagic Blue-Green Algae, R. Marumo, Ed. (OceanResearch Institute, Tokyo, 1975), pp. 36–40], aswell as ultraviolet (UV )-absorbing compounds thatmay each confer protection from photooxidativedamage (80) [A. Subramaniam, thesis, State Univer-sity of New York, Stony Brook (1995)]. Phycobilipro-tein complexes may also play a photoprotectant roleunder high-light conditions by reducing their quan-tum efficiency and by converting some phycourobilin(lmax 5 495 nm) to phycoerythrin (lmax 5 595 nm),thereby reducing absorption at lower wavelengths(80).

40. V. P. Devassy, P. M. Bhattathiri, S. Z. Qasim, IndianJ. Mar. Sci. 7, 168 (1978); R. Santhanam et al., ibid.23, 27 (1994); (36). For a summary of historical re-ports of extensive blooms over the last several de-cades, see E. J. Carpenter and D. G. Capone, in (27),pp. 211–217.

41. The high reflectivity of gas vesicles and the high con-centrations of phycobiliproteins in the cells provideuseful features for remote sensing (42) [S. Tassan,Int. J. Remote Sens. 16, 3619 (1995); C. Dupouy, in(27), pp. 177–191]. Capone et al. (81) recently en-countered a bloom in the Arabian Sea that lastedabout 7 days and was retrospectively observed inimages from AVHRR satellite overpasses in themorning but not in the afternoon, consistent withshipboard observations that the bloom sank slightly(to ;0.5 m) each afternoon. From the satellite imag-ery, the areal extent of this bloom was conservativelyestimated to be ;1 3 106 km2. Trichodesmiumblooms have also been observed from the U.S.space shuttle [for example, D. A. Kuchler and D.Jupp, Int. J. Remote Sens. 9, 1299 (1988)].

42. A. Subramaniam and E. J. Carpenter, Int. J. RemoteSens. 15, 1559 (1994).

43. R. L. Oliver and G. G. Ganf, J. Plankton Res. 10,1155 (1988). Trichodesmium, with its UV-absorbingcompounds and phycobiliproteins, displays higherabsorption than eukaryotic phytoplankton in the UVand blue regions of the spectrum (80).

44. Episodic phytoplankton blooms may contribute tofeedback in the warm water pool climate system inthe Pacific [D. A. Siegel et al., J. Geophys. Res.100, 4885 (1995)]. Cyanobacterial blooms in theBaltic Sea increase sea surface temperature locallyby 1.5°C [M. Kahru, J.-M. Leppanen, O. Rud, Mar.Ecol. Prog. Ser. 191, 1 (1993)]. Phytoplankton bio-mass may influence the flux of moisture [S. Sathy-endranath et al., Nature 349, 54 (1991)]. The denseaccumulation of planktonic biomass at the inter-face may increase viscosity [I. R. Jenkinson,Oceanol. Acta 16, 317 (1993)] providing turbulentdamping and increasing diffusion rates across theair-sea interface.

45. J. O’Neil and M. Roman, in (27), pp. 61–73.46. S. Hawser et al., J. Appl. Phycol. 4, 79 (1992).47. Some harpacticoid copepods are able to graze Tri-

chodesmium to .75% of their body weight per day[J. O’Neil and M. Roman, Hydrobiologia 292–293,235 (1994)] and can consume 33 to 45% of colony Nper day [J. O’Neil, P. M. Metzer, P. Glibert, Mar. Biol.(Berlin) 125, 89 (1996)].

48. From 25 to 50% of recently fixed N by freshly collect-ed colonies could be accounted for by the net re-lease of amino acids [D. G. Capone, M. D. Ferrier, E.J. Carpenter, Appl. Environ. Microbiol. 60, 3989(1994)] or by the accumulation of 15N-dissolved or-ganic N (P. Glibert and D. Bronk, ibid., p. 3996).

49. H. W. Paerl, B. M. Bebout, L. E. Prufert, J. Phycol.25, 773 (1989).

50. Cruises through regions in which Trichodesmiumblooms occur are infrequent, and conventionaloceanographic cruises seldom allow time for oppor-tunistic sampling of blooms. Moreover, because oftheir size and buoyancy, traditional methods of mi-croplankton sampling are strongly biased againstaccurate assessment of Trichodesmium even undernonbloom conditions. Trichodesmium is oftenviewed as a nuisance organism in plankton net–based sampling focused on microplankton collec-tion and quantitation, as high densities clog nets. Netsampling, unless performed carefully at low towingspeeds and with fine-mesh nets (#100 mm), will dis-rupt the relatively delicate colonies. Routine proto-cols for plankton counting, measuring chlorophylland primary productivity on samples drawn from wa-ter bottles, may also impose biases. Because colo-nies are buoyant, they rapidly rise to the top of watersample bottles (sample water is drawn from the bot-tom). Relatively small sample volumes are used inmost of these procedures (;300 ml and often less)and would likely undersample large colonial phyto-plankton, such as Trichodesmium, that typically oc-cur at concentrations at or below one colony per liter(under nonbloom conditions). Prescreening of water,commonly performed to remove large zooplankton,also removes Trichodesmium colonies, thereby elim-inating their contribution to estimates of biomass orC fixation. Moreover, unless discharged by acidtreatment, gas vesicles prevent cells from sinking inUtermohl settling chambers (used for counting phy-toplankton) or sedimenting during centrifugation.Results from conventional oceanographic phyto-plankton surveys in tropical areas should thereforebe viewed cautiously with regard to their accuracy inquantitatively estimating Trichodesmium abundanceand total C and N fixation unless sampling and han-dling protocols were specifically adapted to accountfor these organisms. Recent satellite evidence (41,42) reinforces the proposition that planktonic dia-zotrophs are undersampled.

51. D. G. Capone and E. J. Carpenter, Science 217,1140 (1982); J. N. Galloway et al., Global Biogeo-chem. Cycles 9, 235 (1995). The relatively smallglobal estimate is attributable to the general accep-tance of Carpenter’s (3) calculation for Trichodes-mium N input by N2 fixation, which, for large expans-es of the ocean, depended on historical planktonsurveys that likely underestimated abundances ofTrichodesmium (50).

52. Primary productivity is dependent on two distinctsources of N: “recycled” N, which results from thecycling of autochthonous organic and inorganic ma-terial, and “new” N, which is defined as allochtho-nous N that arrives de novo into a system. Sources ofnew N include NO3

– advecting or diffusing up fromthe large reservoirs of NO3

– at depth, combinedforms of N from wet or dry atmospheric deposition,and N entering into the system through N2 fixation[R. C. Dugdale and J. J. Goering, Limnol. Oceanogr.12, 196 (1967)]. Relative to other sources of new N,nitrate from depth is generally considered the mostimportant (82–84).

53. D. G. Capone, Mitt. Int. Ver. Theor. Angew. Limnol.25, 105 (1996); (82, 85).

54. The oceans likely play a key role in the global C cycle[U. Siegenthaller and J. Sarmiento, Nature 365, 119(1993)]. It is postulated that the marine biota serve asa “biological pump” for sequestering C into the deepocean [A. R. Longhurst and W. G. Harrison, Prog.Oceanogr. 22, 47 (1989)].

55. D. M. Karl et al., Nature 373, 230 (1995); (82). IfPO4

3– limits Trichodesmium, its supply from depthwould co-occur with deep CO2. However, Fe is cur-rently thought to be the most critical limiting nutrientfor Trichodesmium (34–36). In this regard, the atmo-spheric supply of Fe would ultimately constrain Tri-

chodesmium’s capacity to sequester CO2.56. A budget for the upper water column of North Pacific

waters near Hawaii (36) required an input by N2 fix-ation about equivalent to vertical N flux in order toachieve balance. Michaels et al. (86) estimated aninput of N between 50 and 75 Tg year21 to accountfor observed excesses of combined N in the NorthAtlantic; M. Furnas et al. [Great Barrier Reef Re-search Commission Res. Pub. 36 (1995)] invokedextensive N2 fixation to balance nutrient budgets fortwo sections of the Australian shelf–Coral Sea alongthe Great Barrier Reef. The Arabian Sea budget pre-sented by K. Somasundar et al. [Mar. Chem. 30, 363(1990)] indicates a deficit of ;28 Tg N, which theysuggest may be partially offset by N2 fixation. Globalbudgets generally reveal an excess of losses overinputs (51) [J. Christensen et al., Global Biogeo-chem. Cycles 1, 97 (1987); D. G. Capone, in Micro-bial Production and Consumption of GreenhouseGases: Methane, Nitrogen Oxides and Halometh-anes, W. B. Whitman and J. E. Rogers, Eds. (Amer-ican Society for Microbiology, Washington, DC,1991), pp. 255–275]. D. Hansell and T. Y. Water-house [EOS (suppl.) 76, OS17 (1996)] reported ananomaly in surface total organic N in the oligotrophicNorth Pacific, which they attributed to N2 fixation.

57. Vertical NO3– flux is computed as the product of the

NO3– gradient at the nitracline and the vertical eddy

diffusivity (Kz). Nitrate gradients used to calculate thisterm do not vary widely within the major oceanicbasins (74). Although there is considerable debateregarding appropriate values for diapycnal eddy dif-fusivities, values in the range of 1 3 1025 to 5 31025 m22 s21 are commonly reported or used (74,83, 86). Eddy diffusivities for open-ocean systemshave been estimated by various approaches, andlarger values have been reported [for example, (74,88)]. Depth-integrated uptake of 15NO3

–, used toestimate new N demand in the euphotic zone, oftenexceeds NO3

– flux into the upper water column intropical waters (83, 87) [F. King and A. H. Devol,Limnol. Oceanogr. 24, 645 (1979)]. However, tracerincubation methods may cause a substantial nutrientperturbation in highly nutrient-depleted waters,thereby artificially stimulating NO3

2 uptake [P. M.Glibert and D. G. Capone, in Nitrogen Isotope Tech-niques, R. Knowles and T. H. Blackburn, Eds. (Aca-demic Press, New York, 1993), pp. 243–272]. More-over, although it is often assumed that photoinhibi-tion of nitrification prevents recycling of NH4

1 toNO3

2 within the euphotic zone [R. Olsen, J. Mar.Res. 39, 227 (1981)], nitrification occurs in the lowereuphotic zone and some NO3

2 uptake may be con-sidered recycled production [B. B. Ward, K. A. Kil-patrick, E. H. Renger, R. W. Eppley, Limnol. Ocean-ogr. 34, 493 (1989)].

58. Estimates of net productivity inferred from O2 dy-namics in the upper water column exceed estimatesof input fluxes (88); PON export can exceed conven-tional inputs (36, 89).

59. Typically a d15N of 0 to 2 per mil, where d15N equals[(15N:14Nsample/15N:14Nair) 2 1] 3 1000. Organicproduction dependent on N2 fixation results in bio-mass with an N isotopic ratio relatively low in 15N andsimilar to that of the available pool of N2 (that is, d15Nnear zero). This is distinct from organic matter pro-duction dependent on fixed forms of N, which areusually heavier (that is, a greater relative ratio of 15Nto 14N), and thereby provides a distinct signature fortracing the movement of recently fixed N into thefood web. Light PON is most often found in areaswhere Trichodesmium is abundant (84, 90) [E. Wadaand A. Hattori, Nitrogen in the Sea: Forms, Abun-dances, and Rate Processes (CRC Press, Boca Ra-ton, FL, 1991); M. A. Altabet et al., Nature 354, 136(1991); S. A. Macko, L. C. Entzeroth, P. L. Parker,Naturwissenschaften 71, 374 (1984); M. M. Mullin,G. H. Rau, R. W. Eppley, Limnol. Oceanogr. 29, 245(1984)]. Recently, L. Tupas, M. P. Sampson, and D.M. Karl [Eos (suppl.) 76, OS86 (1996)] reported aninput of light surface PON settling into deep (for ex-ample, 1500 m) sediment traps during the summerat the Hawaiian Ocean Time Series (HOT ) stationnorth of Hawaii, which they attributed to Trichodes-mium. Surface PON had a substantially lighter signal,

SCIENCE z VOL. 276 z 23 MAY 1997 z www.sciencemag.org1228

relative to deeper PON, during the bloom we ob-served in the Arabian Sea (81). Isotopically light PONmay also result from other processes, such as up-take of recycled ammonium [D. M. Checkley and L.C. Entzeroth, J. Plankton Res. 7, 553 (1985)], exportof 15N-enriched fecal material from the euphoticzone, or strong isotopic fractionation during nitrateuptake (84) [T. A. Villareal, M. A. Altabet, K. Culver-Rymsza, Nature 363, 709 (1993)], or use of atmo-spherically derived anthropogenic N emissions, suchas NOx and NH3/NH4

1 (91). However, these pro-cesses are unlikely in the highly oligotrophic openocean (90).

60. Trichodesmium spp. can represent a major fraction(;60%) of total chlorophyll and are important in pri-mary production and N input in the Caribbean Sea(92). In the Indian Ocean, Trichodesmium can reacha biomass of up to 30 mg of chlorophyll a per cubicmeter (79). E. J. Carpenter and K. Romans [Science254, 1356 (1991)] concluded that Trichodesmium isthe most abundant phytoplankton in the tropicalNorth Atlantic. C. S. Davis, S. M. Gallager, and A. R.Solow [ibid. 257, 230 (1992)], using towed videomicroscopy, found very high abundance of Tri-chodesmium in North Atlantic waters. Trichodes-mium accounted for a mean of 17% of total chloro-phyll a throughout the year at the HOT station (58).Richelia intracellularis (a common endosymbiont ofdiatoms) [T. Villareal, in (27), pp. 163–175] can alsocontribute substantially to new N inputs (E. J. Car-penter et al., in preparation).

61. Subtropical studies (93–95) found that even duringthe summer, when Trichodesmium was relativelyabundant and active, areal rates of N2 fixation weregenerally low compared with tropical waters.

62. K. R. Gunderson et al., Pac. Sci. 30, 45 (1976).63. T. Saino, thesis, University of Tokyo (1977).64. D. G. Capone et al., in preparation.65. J. J. Goering et al., Limnol. Oceanogr. 11, 614

(1966).66. T. Osborn, personal communication; K. L. Polzin, J.

M. Toole, J. R. Ledwell, R. W. Schmitt, Science 276,93 (1997). J. R. Ledwell et al. [Nature 364, 701(1993)] estimated Kz values of 1.1 3 1025 m2 s21 ata site 1200 km west of the Canary Islands during anextended (several months) SF6 tracer experiment; K.A. VanScoy and D. E. Kelley [Eos (suppl.) 76, OS40(1996)] recently reported an integrated, basin-widebulk Kz upper limit of 3 3 10–5 m2 s–1 on the basis ofa two-decade data set of tritium penetration.

67. For a bloom described in (65), we derived an input in

the top 0.5 m of 115 mmol N m–2 day–1, comparedwith the average of 41 mmol N m–2 day–1 for other,nonbloom stations during the same period. Similarly,a surface accumulation with input estimated to be;300 mmol N m22 day21 was reported in (62). For abloom encountered in the early spring intermonsoonin the central Arabian Sea (81), we estimated thebloom contribution of N through N2 fixation to be;100 mmol N m–2 day–1, compared with 35 mmol Nm22 day21 through the rest of the water column.

68. H. W. Paerl, Nature 316, 747 (1985); R. Duce et al.,Global Biogeochem. Cycles 5, 193 (1991); S. Cornellet al., Nature 376, 243 (1995); (91).

69. M. B. McElroy, Nature 302, 328 (1983); P.Falkowski, ibid., in press; L. A. Codispoti, in Produc-tivity of the Ocean: Present and Past, W. H. Berger,V. S. Smetacek, G. Wefer, Eds. ( Wiley, New York,1989), pp. 377–394.

70. J. P. Zehr and D. G. Capone, Microbiol. Ecol. 32,263 (1996).

71. C. G. Ehrenberg, Ann. Phys. Chem. 18, 477 (1830).72. T. Saino and A. Hattori, in The Kuroshio IV, A. Y.

Takenouchi, Ed. (Saikon, Tokyo, 1979), pp. 697–709.

73. R. Letelier and D. Karl, Mar. Ecol. Prog. Ser. 133,263 (1996).

74. J. J. McCarthy and E. J. Carpenter, in (3), pp. 487–512.

75. T. Roenneberg and E. J. Carpenter, Mar. Biol. (Ber-lin) 117, 693 (1994).

76. T. Kana, Limnol. Oceanogr. 38, 18 (1993).77. P. Siddiqui, E. J. Carpenter, B. Bergman, J. Phycol.

28, 320 (1992).78. S. Janson, E. J. Carpenter, B. Bergman, FEMS Mi-

crobiol. Lett. 118, 9 (1994).79. I. Bryceson and P. Fay, Mar. Biol. (Berlin) 61, 151

(1981).80. A. Subramaniam, E. J. Carpenter, D. Karentz, P. G.

Falkowski, in preparation.81. D. G. Capone et al., in preparation.82. R. Eppley and B. Peterson, Nature 282, 677 (1979).83. M. R. Lewis, W. G. Harrison, N. S. Oakey, D. Hebert,

T. Platt, Science 234, 870 (1986).84. M. A. Altabet, Deep Sea Res. 35, 535 (1988).85. W. Berger et al., Scripps Inst. Oceanogr. Ref. 87-

30:1 (1987).86. A. Michaels et al., Biogeochemistry 35, 181 (1996).87. L. Legendre and M. Gosselin, Limnol. Oceanogr. 34,

1374 (1989).88. W. J. Jenkins, Nature 331, 521 (1988).89. D. M. Karl et al., in preparation.

90. E. J. Carpenter, B. Fry, H. R. Harvey, D. G. Capone,Deep Sea Res. 44, 27 (1997).

91. H. W. Paerl and M. L. Fogel, Mar. Biol. (Berlin) 119,635 (1994).

92. E. J. Carpenter and C. Price, Limnol. Oceanogr. 22,60 (1977).

93. E. J. Carpenter and J. J. McCarthy, ibid. 20, 389(1975).

94. J. J. McCarthy and E. J. Carpenter, J. Phycol. 15, 75(1979).

95. T. Mague et al., Mar. Biol. (Berlin) 41, 213 (1977).96. J. Felsenstein, PHYLIP Phylogeny Inference Pack-

age (version 3.5c), Department of Genetics, Univer-sity of Washington, Seattle.

97. Water samples were collected at various depths overthe upper 200 m by CTD (conductivity, temperature,depth)-rosette. Trichodesmium population abun-dance and depth distribution were obtained by filter-ing whole water bottles onto 5- or 10-mm filters,followed by direct counting by epifluorescent mi-croscopy. For N2 fixation, colonies of Trichodes-mium were obtained by plankton tows at 10 to 15 mand assayed using the C2H2 reduction method.Samples were incubated on deck under full sunlight(100%) or with neutral density screening to approxi-mate light levels of 1, 10, 28, and 55% of surfaceirradiance. For estimates of N2 fixation, we assumeda conversion ratio of 4 mol of C2H4 produced permole of N2. Volumetric rates of N2 fixation were es-timated by scaling the rates determined on isolatedcolonies with reference to the population density ob-served at the depth corresponding to the light level ofincubation. Chlorophyll of the ,202-mm fraction wasdetermined by standard methods, whereas Tri-chodesmium chlorophyll was estimated by deter-mining chlorophyll a per cell and scaling that to totaltrichome counts at each depth. Primary productivitywas determined using 14C-bicarbonate on assays of,202-mm bulk water from each depth or, as for N2fixation, on isolated colonies and scaled to trichomeconcentrations at specific depth.

98. We thank M. Mulholland, J. Burns, L. Duguay, J.Montoya, and two anonymous reviewers for theircomments, A. Parrella for technical assistance, andF. Younger for artwork. We particularly thank A. Sub-ramaniam for discussion and help on the remotesensing images. D.G.C. thanks B. Taylor for intro-ducing him to Trichodesmium. Supported by theNSF Divisions of Ocean Sciences and EnvironmentalBiology.

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