Bacterial and Eukaryotic Intact Polar Lipids.pdf

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Bacterial and eukaryotic intact polar lipids in theeastern subtropical South Pacific: Water-column

distribution, planktonic sources, and fatty acid composition

Benjamin A.S. Van Mooy ⇑, Helen F. Fredricks

Woods Hole Oceanographic Institution, Department of Marine Chemistry and Geochemistry, MS #4, Woods Hole, MA 02543, USA

Received 19 January 2010; accepted in revised form 17 August 2010; available online 27 August 2010

Abstract

Fatty acids are generally the most abundant lipid molecules in plankton, and thus play a central role in the cycling of organic matter in the upper ocean. These fatty acids are primarily derived from intact polar diacylglycerolipids (IP-DAGs),which compose cell membranes in plankton. The molecular diversity of IP-DAGs in the upper ocean remains to be fully char-acterized, and the advent of high performance liquid chromatography/electrospray ionization–mass spectrometry (HPLC/ESI-MS) approaches have now provided the opportunity to readily analyze IP-DAGs from marine planktonic communities.We used HPLC/ESI-MS to determine the concentrations of three classes of phospholipids (phosphatidlyglycerol (PG), phos-phatidylethanolamine (PE), and phosphatidylcholine (PC)), three classes of betaine lipids (diacylglyceryl trimethylhomoserine(DGTS), diacylglyceryl hydroxymethyl-trimethyl-b-alanine (DGTA), and diacylglyceryl carboxyhydroxymethylcholine(DGCC)), and three classes of glycolipids (monogalactosyldiacylglycerol (MGDG), digalactosyldiacylglycerol (DGDG),and sulfoquinovosyldiacylglycerol (SQDG)) in plankton filtered (>0.2 lm) from seawater collected within the euphotic zoneof the eastern South Pacific. The distributions of these IP-DAGs along the cruise transect provided important new insights ontheir tentative planktonic sources. Complementary data from our cruise, a principle components analysis of our IP-DAG con-centrations, observed fatty acid compositions of IP-DAG classes and published IP-DAG distributions in pure cultures of plankton suggest that heterotrophic bacteria were the dominant sources of PG and PE, while MGDG and SQDG originatedprimarily from Prochlorophytes. The origins of the other classes of IP-DAGs were less clear, although it is likely that PC,DGTS, DGTA, and DGCC were derived primarily from eukaryotic phytoplankton. The molecular distributions of fatty acidsattached to the different classes of IP-DAGs were generally distinct from one another, and suggest that reported distributionsof total fatty acids (as analyzed by gas chromatography) in the literature should be regarded as homogenized mixtures of dis-tinct molecular pools of fatty acids. 2010 Elsevier Ltd. All rights reserved.

1. INTRODUCTION

Fatty acids are commonly reported to be the most abun-dant lipidic components of plankton and particles in theocean, and it is clearly recognized that these fatty acidsare derived from a number of different groups of molecules

including intact polar diacylglycerols (IP-DAGs), triacyl-

glycerols (TAGs), wax esters, and sterol esters (Volkmanand Everitt, 1986; Wakeham and Canuel, 1988; Volkmanet al., 1989; Wakeham et al., 1997a). Yet the studies thathave independently examined TAGs, wax esters, and sterols(e.g. Wakeham and Canuel, 1988), indicate that IP-DAGsare a dominant molecular source of fatty acids in marineparticles. Thus, IP-DAGs as a group are likely to composethe most abundant group of lipid molecules in the sea.

In addition to being the primary membrane and energystorage molecules in plankton, fatty acids play important

0016-7037/$ - see front matter 2010 Elsevier Ltd. All rights reserved.

doi:10.1016/j.gca.2010.08.026

⇑ Corresponding author. Tel.: +1 508 289 2322; fax: +1 508 4572164.

E-mail address: [email protected] (B.A.S. Van Mooy).

www.elsevier.com/locate/gca

Available online at www.sciencedirect.com

Geochimica et Cosmochimica Acta 74 (2010) 6499–6516

http://dx.doi.org/10.1016/j.gca.2010.08.026mailto:[email protected]://dx.doi.org/016/j.gca.2010.08.026http://dx.doi.org/016/j.gca.2010.08.026mailto:[email protected]://dx.doi.org/10.1016/j.gca.2010.08.026


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roles in hormone production (Irigoien, 2004), cell–cell sig-naling (Vardi et al., 2008) and chemical defense (Miraltoet al., 1999). Fatty acids are a key component of the biolog-ical pump, where they compose 20% of the particulate or-ganic carbon export flux from the euphotic zone to thedeep-sea (Wakeham et al., 1980, 1984, 1997b; Goutxet al., 2007) and make a significant contribution to dis-

solved organic carbon in the deep-sea (Hwang and Druffel,2003).

The body of literature on fatty acids in marine planktonand particulate matter, was built largely through the appli-cation of gas chromatography (GC). Yet the IP-DAGsources of these fatty acids have largely escaped detailedexamination. IP-DAGs are not amenable to GC analysis,and analyzing fatty acids by GC requires that fatty acidsbe first liberated from IP-DAGs by using chemicalreactions in the laboratory. Recent studies employing highperformance liquid chromatography/electrospray ioniza-tion–mass spectrometry (HPLC/ESI-MS) have revealed abroad molecular diversity of IP-DAGs (e.g. glycolipids,

betaine lipids) in marine environments that extends far be-yond the more commonly known phospholipids (Rütterset al., 2002; Zink et al., 2003; Sturt et al., 2004; Suzumura,2005; Van Mooy et al., 2006, 2009; Schubotz et al., 2009 ).

Automated thin layer chromatography/flame ionizationdetection (TLC/FID) instruments (e.g. Iatroscan) (Volk-man and Everitt, 1986; Volkman et al., 1989; Lombardiand Wangersky, 1995; Striby et al., 1999) as well as HPLCmethods incorporating evaporative light scattering detec-tion (HPLC/ELSD) (Nordbäck et al., 1998) have providednovel insights on the distributions of IP-DAGs and otherfatty-acid-bearing molecules, and promised the degree of throughput required for comprehensive, quantitativeoceanographic studies. However, these two general meth-

ods are faced with the chromatographic challenges of resolving classes of IP-DAGs by their polarity alone ( Stribyet al., 1999), and unresolved classes of IP-DAGs are gener-ally reported simply as “polar lipids” (Volkman et al., 1989;Lombardi and Wangersky, 1995). While these methodsmay still detect the presence of ‘unknown’ classes of IP-DAGs (i.e. IP-DAGs for which no standard is availableto the analyst; e.g. Lombardi and Wangersky, 1995) theyprovide very little information to determine their identity.Finally, TLC/FID and HPLC/ELSD are strictly chromato-graphic methods, and do not independently provide struc-tural information about the constituent fatty acids of IP-DAGs.

In contrast, HPLC/ESI-MS methods offer high-throughput, quantitative analysis of IP-DAG classes, whilealso providing information on the fatty acids of these IP-DAGs. Importantly, HPLC/ESI-MS methods may alsobe readily configured to collect structural information onunknown lipids in the course of routine quantitative IP-DAG analyses. It is by this tactic that we first identified sul-folipids (SQDG) and betaine lipids in the ocean usingHPLC/ESI-MS (Van Mooy et al., 2006, 2009); these mole-cules had long been known to be major components of phy-toplankton membranes (Kato et al., 1996; Wada andMurata, 1998) and we identified them through a relativelystraightforward comparative analyses of retention times

and mass spectra from cultures of plankton known or ex-pected to contain these molecules.

In this paper, we present a large-scale assessment of planktonic IP-DAGs in the euphotic zone ocean of the east-ern subtropical South Pacific. Our aim was to contribute in-sights on the molecular diversity of IP-DAGs and theirplanktonic sources. We are hopeful that, in conjunction

with the growing number of additional reports of IP-DAGsin natural planktonic communities (Van Mooy et al., 2006,2009; Ertefai et al., 2008; Schubotz et al., 2009; Vardi et al.,2009), our study will shed new light on the immense body of fatty acids literature covering the distribution, sources andbiogeochemical cycling of fatty acids in the sea.

2. METHODS

2.1. Sampling

Samples were collected in the South Pacific during theLeg II of the BIOSOPE campaign in November and

December of 2004 (Fig. 1). Seawater was obtained usingNiskin bottles mounted on a CTD-rosette from depths cor-responding to 100%, 50%, 15%, 3%, 1% and 0.3% levels of surface irradiance. Two-liter samples were taken from theNiskin bottles immediately upon their retrieval and filteredunder 200 mbar vacuum. Samples were filtered through0.2 lm pore size alumina membranes (Anodisc; Whatman);filtration was completed within 1 hour of retrieving thesamples. All membranes and filters were immediatelyplaced in envelopes made from combusted aluminum foil,and snap frozen in liquid nitrogen. Samples were trans-ported in a liquid nitrogen dry shipper to the laboratory

110˚W 100˚W 90˚W 80˚W 70˚W 60˚W60˚S

50˚S

40˚S

30˚S

20˚S

10˚S

S t n 1

1

S t n 1

5

S t n 1

7

S t n

1 8

EQ

Fig. 1. Map of locations where seawater samples were collected forIP-DAG analysis in the eastern subtropical South Pacific.

6500 B.A.S. Van Mooy, H.F. Fredricks / Geochimica et Cosmochimica Acta 74 (2010) 6499–6516


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in Woods Hole and transferred to liquid nitrogen upontheir arrival. Alumina membranes were combusted at450 C for 6 h prior to use.

2.2. IP-DAG extraction

The anodiscs were extracted using a modification of the

Bligh and Dyer method (Bligh and Dyer, 1959). Each ano-disc was placed in a combusted 50 ml glass centrifuge tube,and the following solvents were added: 6 mL phosphatebuffered saline (PBS), 15 mL methanol and 7.5 mL dichlo-romethane (Fisher Scientific). The samples were vortexed,sonicated for 15 min, and let stand overnight at 20 Cunder nitrogen. Next, 7.5 mL each of PBS and dichloro-methane were added, the sample vortexed again, and letstand for a few minutes to allow separation of the phases.The dichloromethane layer was removed to a vial andblown down to 100 lL with nitrogen, where it was trans-ferred to a 300 lL glass HPLC vial insert and blown downto near dryness, then 200 lL of 9:1 dichloromethane:meth-

anol was added. Finally, 20 l

L of this total lipid extract wasinjected for the analysis described below.

2.3. HPLC/MS analyses

IP-DAGs were analyzed using an LCQ Deca XP ion-trapmass spectrometerequippedwith an ESIsource connectedtoa Surveyor HPLC system (ThermoFinnigan San Jose, CA).A PVA-Sil column (YMC, Waters, S5, 120 Å, 150 mm 2 mm i.d.) was used with the following solvent gradient at0.5 mL min1: from100% A to85%A:15%B in20 min,thento 50% A and 50% B over 25 min, then hold for 2 min. Thecolumn was equilibrated with 100% A for 10 min at1 mL min1 prior to injecting the next sample. Eluent A

was 790:200:1.2:0.4 of n-hexane:2-propanol:formic acid:14.8 N NH4 (aq) and eluent B was 880:100:1.2:0.4 of 2-propa-nol:water:formic acid: 14.8 N NH4 (aq). The electrospraysource and the mass spectrometer were configured as pub-lished previously (Sturt et al., 2004) and programmed suchthat the base peak from each positive ion full scan (500– 2000 Da) was fragmented up to MS3.

2.4. IP-DAG standards and quantification

Diagnostic retention times were established from stan-dards of each IP-DAG class, as were characteristic positiveand negative ionization MS2 and MS3 spectra. Syntheticphosphatidlyglycerol (PG), phosphatidylethanolamine(PE), and phosphatidylcholine (PC), were obtained fromAvanti Polar Lipids Inc. (Alabaster, AL). Natural monoga-lactosyldiacylglycerol (MGDG) and digalactosyldiacylglyc-erol (DGDG) standards (from wheat) were obtained fromMatreya, LLC (Pleasant Gap, PA). Galactose has long beenrecognized to be the dominant, if not exclusive, sugar groupin IP-DAGs in cyanobacteria and algae (Janero and Barr-nett, 1981; Sato and Murata, 1982). As such, MGDG andDGDG are frequently reported without explicit verificationof their sugar content (reviewed by Wada and Murata, 1998;Dörmann and Benning, 2002). So although trace amounts of monoglucosyldiacylglycerol have been reported in cyano-

bacteria (0.6% in photoautotrophic Synechcystis Sato andMurata, 1982), we did not independently verify the molecu-lar distribution of sugars in either MGDG or DGDG. Sulfo-quinovosyldiacylglycerol (SQDG) was isolated fromSynechococcus WH8102 (kindly provided by E.A. Webb,University of Southern California) by using preparativeHPLC (described below).

Three classes of betaine lipids – diacylglyceryl trimethyl-homoserine (DGTS), diacylglyceryl hydroxymethyl-tri-methyl-b-alanine (DGTA), and diacylglyceryl carboxy-hydroxymethylcholine (DGCC) – were identified by theirpublished MS2 and MS3 spectra (Schubotz et al., 2009), rel-ative retention orders (Vogel et al., 1990; Vogel and Eichen-berger, 1992), and diversity of fatty acids (Vogel andEichenberger, 1992; Dembitsky, 1996; Kato et al., 1996).DGTS is significantly less polar than DGTA (Vogel et al.,1990) and elutes several minutes earlier than DGTA underour chromatographic conditions; DGTS also generally con-tains considerably shorter and less saturated fatty acids thanDGTA (Dembitsky, 1996; Kato et al., 1996). A DGTA stan-

dard was isolated from Chaetoceros affinis (kindly providedby S.T. Dyhrman, Woods Hole Oceanographic Institution)by using preparative HPLC (Van Mooy et al., 2009). We as-sumed that the DGTA standard was also suitable for thequantification of DGTS and DGCC. We feel that thisassumption is justified because, all three betaine lipids sharequaternary amine moieties,which, similar to PC,is where thecharge appears to reside under positive ionization. Further-more, DGTA and DGTS are structural isomers (Vogelet al., 1990). IP-DAGs were quantified in positive ionizationmode only. The molecular ion chromatograms were ex-tracted for each individual IP-DAG species at their appropri-ate retention time, integrated, and applied to externalstandard curves. The standard curves were composed of

triplicate measurements of IP-DAG standards at fourconcentrations. We found that instrument response fordifferent classes of IP-DAGs varied by a factor of 20(response factors are given in Electronic Annex EA-1) Therelative response factors (peak area (mole)1) followed theorder: PE > PG > MGDG > PC > DGDG DGTA >SQDG. In other words, the instrument was most sensitiveto PG and least sensitive to SQDG. The limit of quantifica-tion for these standards was on the order of 10 pmol on col-umn. A single mid-range standard was analyzed after everyfifth sample was analyzed as a check on the stability of theESI source. Based on the reproducibility of these standardanalyses, we estimate thatthe analyticalerror associatedwithour IP-DAG quantification was on the order of 10–15%.

2.5. Preparative HPLC

Culturing Synechococcus WH8102 and C. affinis underphosphorus-limiting conditions resulted in IP-DAG ex-tracts that were heavily enriched in SQDG and DGTA,respectively. Culture media and conditions have been de-scribed (Van Mooy et al., 2009). The analytical HPLC con-ditions described above (Section 2.3) were sufficient to yieldchromatographically pure fractions of SQDG and DGTA.Our preparative HPLC routine began by first running theculture extracts three times on the HPLC/ESI-MS to assure

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center of the ultraoligotrophic South Pacific subtropicalgyre (Claustre et al., 2008), which is characterized by in-tense stratification, a deep chlorophyll maximum (DCM)at 200 m that is overlaid by surface waters with verylow (0.02 lg L1) chlorophyll a concentrations (Raset al., 2008), inorganic nitrogen concentrations that are be-low detection limits (Raimbault et al., 2008), and abundant(>100 nmol L1) dissolved phosphate (Moutin et al., 2008).Stations 15 through 18 bracket the transition between thegyre and the Chilean coastal upwelling zone (Claustreet al., 2008). At Station 15 and stations just to the east,the DCM was particularly intense. At Stations 17 and 18,the DCM was shallower, and high nitrate low chlorophyll

(HNLC) conditions were observed there (Raimbaultet al., 2008).

3.2. Overview of IP-DAGs in the eastern South Pacific

Total lipid extracts from 23 samples were examined fortheir intact polar lipid content. We identified 9 major clas-ses of IP-DAGs that comprise the focus of this paper (Table1): three classes of phospholipids, phosphatidylglycerol(PG), phosphatidylethanolamine (PE), and phosphatidyl-choline (PC); three classes of betaine lipids diacylglyceryltrimethylhomoserine (DGTS), diacylglyceryl hydroxy-methyl-trimethyl-b-alanine (DGTA), and diacylglyceryl

carboxyhydroxymethylcholine (DGCC); and three classesof glycolipids, monogalactosyldiacylglycerol (MGDG),digalactosyldiacylglycerol (DGDG) and sulfoquinovosyl-diacylglycerol (SQDG) (Table 1). These 9 major IP-DAGclasses were present in every sample, with the exceptionof samples from Station 11 where DGDG was absent. Fur-thermore, the 9 major classes IP-DAGs dominated the total

IP-DAG distribution such that their presence in our sam-ples accounted for 97 ± 2% of the total peak area in thechromatograms on average. The sum of the 9 major IP-DAG classes was highly linearly correlated (r2 = 0.72,

p < 0.01) with the sum of heterotrophic bacteria, cyanobac-teria, and eukaryotic phytoplankton determined by flowcytometry and pigment concentration (Grob et al., 2007;Ras et al., 2008). Other readily identifiable phospholipidsincluded methylated forms of PE (Schubotz et al., 2009).Ornithine lipids were also observed at a few stations (Schu-botz et al., 2009; Van Mooy et al., 2009) as were cerebro-sides (Schubotz et al., 2009; Vardi et al., 2009). Severalother molecules showed mass spectra with common fatty

acid or diglyceride moiety ions, but were not readily identi-fiable by us.Intact polar di- and tetra-ether glycerolipid molecules

derived from planktonic Archaea (DeLong et al., 1998;Schouten et al., 2000) were not detected by us. This resultwas not entirely unexpected because these organisms aregenerally thought to be scarce in surface waters comparedto bacteria (DeLong et al., 1999; Karner et al., 2001),although archaeal abundance data were not available fromour cruise to actually make this comparison. In addition,the work of Nishihara and Koga (1987) along with a morerecent study by Huguet et al. (2010) has shown that the tra-ditional Bligh and Dyer (1959) extraction method we em-ployed is almost completely ineffective at extracting intact

polar lipids from archaea. In an examination of planktonin the Black Sea, Schubotz et al. (2009) used a soxhletextraction method, which according to Huguet et al.(2010) is markedly more effective for extracting intact polarlipids from marine archaea than the Bligh and Dyer meth-od. Yet Huguet et al. (2010) also reported that archaealpolar lipids were absent in the euphotic zone.

3.3. Planktonic phospholipids

The concentrations of the major phospholipids (i.e. PG,PE and PC) ranged from 582 pmol L1 of PC in surfacewaters at Station 18, to 16 pmol L1 of PG in the deepest

waters of Station 11, but the majority of concentrationswere confined to between 100 and 300 pmol L1 (Fig. 3).Compared to the major glycolipids and betaine lipidsincluded in this study, the major phospholipids composed20 ± 5% of total IP-DAGs on a molar basis when averagedacross the entire dataset. Looking across the data from theentire cruise, PC concentrations were distinctly greater thanPG concentrations (n = 23; p < 0.01), but there were noother significant differences between the concentrations of the three classes of phospholipids.

In general, concentrations of individual major phospho-lipids tended to decrease with increasing depth in the watercolumn; with only a few exceptions, the greatest concentra-

Fig. 2. Water columnprofiles of environmentalproperties along thecruise transect. Top: temperature (C). Middle: chlorophyll aconcentrations (lg L1). Bottom: phosphate concentrations(nmol L1).



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Table 1Overview of IP-DAGs analyzed.

Type Class Structure

Phospholipid Phosphatidylglycerol (PG)

HO OP

OH

O

O

OO R1

O

O

O

R2

Phosphatidylethanolamine (PE)

H3NO

PO

O

OO R1

O

O R2

O

Phosphatidylcholine (PC)

NO

PO

O

OO R1

O

O R2

O

Betaine lipid Diacylglyceryl trimethylhomoserine (DGTS)

O O R1

O

O

O

R2

N

OO

Diacylglyceryl hydroxymethyl-trimethyl-b-alanine (DGTA)

O O R1

O

O

O

R2

N

OO

Diacylglyceryl carboxyhydroxymethylcholine (DGCC)

O O R1

O

O

O

R2

O

OO

N



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Table 1 (continued )

Type Class Structure

Glycolipid Monogalactosyldiacylglycerol (MGDG)

O O R1

O

O R2

O

O

OH

HO

HO

HO

Digalactosyldiacylglycerol (DGDG),

O O R1

O

O R2

O

O

OH

HO

HO

HO 2

Sulfoquinovosyldiacylglycerol (SQDG)

O O R1

O

O R2

O

O

OH

HO

HO

S

O

O

O

R1 and R2 represent acyl (i.e. fatty acid) groups, which define species within each class of IP-DAGs.Charges are those expected at seawater pH.

PG concentration (pmol L-1

)

0 50 100 150 200 250 300

D e p t h ( m )

0

50

100

150

200

250

Stn 11

Stn 15

Stn 17Stn 18

PE concentration (pmol L-1

)

0 100 200 300 400 500

PC concentration (pmol L-1

)

0 100 200 300 400 500 600

Fig. 3. Profiles of concentrations of phospholipids. Locations of stations shown in Fig. 1.



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tion of these molecules at any given station was observed atdepths shallower than 50 m. One notable exception to thiswas an apparent deep maximum in phospholipid concentra-tions at 150 m depth at Station 11, which coincided with thetop of the deep chlorophyll maximum. Concentrations alsotended to increase from west to east along the transect.

3.4. Planktonic betaine lipids

All three of the known microbial betaine lipids were ob-served: DGTS, DGTA, and DGCC. The concentrationsranged from 1392 pmol L1 DGTA in the surface watersat Station 18 to undetectable levels of DGCC in the deepestwaters at Station 11 (Fig. 4). On average, betaine lipidscomposed 30 ± 13% of the total major IP-DAGs. Examin-ing the dataset as a whole, a distinct hierarchy in theabundance of the betaine lipids was apparent, whereDGTA > DGTS > DGCC (n = 23; p < 0.05).

Similar to phospholipids, the concentrations of betainelipids tended to be highest in surface waters and decreasedwith increasing depth in the water column. Furthermore,the concentrations were higher in the eastern stations of the transect than in the western stations. At Station 15 therewere maximums in all three betaine lipids at 50 m depth,which were much more pronounced than observed in the

profiles of the phospholipids at that station.

3.5. Planktonic glycolipids

Themajor glycolipids we observed were MGDG, DGDGand SQDG. The concentrations ranged from 1560 pmol L1

of SQDG at the deep chlorophyll maximum at Station 15 toundetectable levels of DGDG (i.e.


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observed for betaine lipids, there was a distinct hierarchy inthe abundance of glycolipids: SQDG > MGDG > DGDG(n = 23, p < 0.05).

Concentrations of MGDG and SQDG were strongly lin-early correlated with one another (R2 = 0.79; p < 0.01), andwere generally several-fold more abundant than DGDG.The water profiles of MGDG and SQDG did not decrease

with depth in a manner similar to phospholipids. Instead, atStations 11 and 15 in the gyre we observed pronounceddeep maximums that roughly correlated with the DCMs;at Stations 17 and 18 in HNLC region of the transect, deepmaximums in SQDG were also observed, but the concen-trations of MGDG were comparatively similar throughoutthe euphotic zone. The water column profiles of DGDGwere distinct from that of the other two major glycolipids.At Station 15, there appeared to be a maximum at theDCM similar to MGDG and SQDG, but surface concen-trations were also relatively high. At Stations 17 and 18,the DGDG profiles more closely resembled those of phospholipids.

3.6. Substitute lipid ratios

Plankton in the open ocean have been shown to substi-tute non-phosphorus lipid for phospholipids under condi-tions of phosphorus limitation (Van Mooy et al., 2006,2009), and we calculated molar ratios of these non-phosphorus lipids to their corresponding phospholipids(Fig. 6). SQDG:PG and betaine lipid:PC ratios (BL:PC)in the upper mixed layer (0.70) loaded variable.

The abundances of all three classes of phospholipids andall three classes of betaine lipids were significantly positivelyloaded on principal component one (PCA1-1), whichaccounted for 45% of the variability in the data we includedin the PCA (Fig. 7). The abundances of eukaryotic

picophytoplankton, picocyanobacteria from the genus Syn-echococcus, and heterotrophic bacteria were also signifi-cantly positively loaded on PCA1-1. In contrast, theconcentrations of MGDG, SQDG, chlorophyll a and pico-cyanobacteria from the genus Prochlorococcus were signifi-cantly positively loaded on the second principal component

SQDG:PG ratio (mol/mol)

0 5 10 15

D e p t h ( m )

0

50

100

150

200

250

Stn 11

Stn 15

Stn 17

Stn 18

Betaine lipids:PC ratio (mol/mol)

0 5 10

Fig. 6. Profiles of concentration ratios of SQDG, PG, betaine lipids, and PC. Locations of stations shown in Fig. 1.

Principal component 1 (PCA1-1;45%)

-0.5 0.0 0.5 1.0 1.5

P r i n c i p a l c o m p o n e n

t 2 ( P C A 1 - 2 ; 2 4 % )

-0.5

0.0

0.5

1.0

1.5

Pro

Euk

Syn

Het Bact

MGDGSQDG

PG

PE

PC

DGTS

DGCC

DGTA

Chl a

PO4

NO3

T

Fig. 7. Results of PCA1. The percentages in parentheses indicatethe amount of variability accounted for by each principle compo-nent. Pro = Prochlorococcus cells abundances; Syn = Synechococ-

cus cell abundances; Euk = eukaryotic phytoplankton abundance;Het Bact = heterotrophic bacteria abundances; Chl a = chloro-phyll a concentrations.



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(PCA1-2), which accounted for 24% of the variability. Tem-perature, phosphate, and nitrate were the only factors sig-nificantly loaded on a third principal component (notshown), which accounted for 19% of the variability in thedataset.

3.8. Diversity of IP-DAG diglyceride moieties

Our analysis of diglyceride moieties showed, as ex-pected, that the most common marine planktonic fattyacids identified by GC (i.e. 14:0, 16:0, 16:1, 18:0, 18:1,18:2, 20:5, 22:6; Wakeham and Canuel, 1988; Volkmanet al., 1989; Viso and Marty, 1993; Wakeham et al.,1997a) are also represented in our analyses of IP-DAGsby HPLC/ESI-MS (Table 2). These fatty acids could befound attached to almost any IP-DAG, however each classof IP-DAG tended to have characteristic combinations of fatty acids. PG and PE were dominated by 16- and 18-car-bon fatty acids, and were the only IP-DAGs to containreadily-identifiable 17-carbon fatty acids. PC contained

substantially greater proportions of long-chain polyunsatu-rated fatty acids (e.g. 20:5 and 22:6) than the other phos-pholipids. MGDG and DGDG were unique from otherIP-DAGs in that they contained both 14-carbon fatty acidsand polyunsaturated 18-carbon fatty acids. SQDG was sim-ilar to MGDG and DGDG in that it contained abundant14-carbon fatty acids; however the polyunsaturated 18-carbon fatty acids were absent. DGTS contained manydozens of different combinations of fatty acids, making itthe most diverse class of IP-DAGs by far; the diglyceride

moieties contained mainly 32, 34 and 36 carbon atoms withmultiple unsaturations, but there was little consensus onspecific fatty acids. The fatty acids of DGTA were also di-verse, but were characterized by a dominance of 20:5 and22:6. Only three different fatty acids contributed substan-tially to DGCC: 14:0, 16:0 and 22:6.

Examining the average number of carbon atoms in the

diglyceride moieties of IP-DAGs from across the transectshowed that SQDG, PE, PC, DGTA, and DGCC wereall distinct from one another (n = 23; p < 0.05; Fig. 8).The number of carbon atoms in diglyceride moieties of MGDG, DGDG, and DGTS composed a distinct set atthe low end of the range; PG and DGTS also formed a dis-tinct set. Furthermore, variation in the range of the numberof carbon atoms among the classes of IP-DAGs is striking.For example, SQDG is overwhelmingly composed of spe-cies with diglyceride moieties of 30 atoms: the mean we ob-served was 29.8 carbon atoms and a range of only 29.0 to30.1. On the other end of the spectrum, DGTA containeddiglyceride moieties with a mean of 40.4 carbon atoms

and a range between 38.9 and 44.0.Similar overall trends were observed in the number of double bonds in the diglyceride moieties, although therewerefewer distinctions between classes of IP-DAGs (Fig. 9).Nonetheless, fatty acids of SQDG are distinctly saturated:thediglyceride moieties of this class averaged barelyone dou-ble bond (e.g. with fatty acids of zero and one double bond).Again on the other end of the spectrum, the fatty acids of DGTA were distinctly unsaturated with a mean of 8.7doublebonds and a range between 7.4 and 12.0.

Table 2

Summary of observed diglyceride moieties within each class of IP-DAGs. The first term (e.g. 32:1) is the number of carbon atoms and doublebonds in the diglyceride moiety as whole. The first term in parenthesis (e.g. 23%) is the average abundance of the diglyceride moiety withineach class of IP-DAGs. The additional term(s) in parenthesis are the specific pair(s) of individual fatty acids that make up the diglyceridemoiety.

Phospholipids

PG 32:1 (23%; 16:0/16:1), 34:2 (23%; 16:1/18:1 and 16:0/18:2), 36:2 (20%; 18:1/18:1), 32:2 (10%; 16:1/16:1), 33:1 (10%; 16:0/17:1),32:2 (7%; 16:0/18:1)

PE 34:2 (24%; 16:1/18:1 and/or 17:1/17:1), 32:1 (13%; 16:1/18:1), 38:6 (13%; 16:0/22:6), 35:2 and/or 36:9 (11%, nd), 32:2 (8%; 16:1/16:1), 44:12 (6%; 22:6/22:6), C33:1 or C34:8 (5%, nd), 40:6 (5%; 18:0/22:6), 39:0 or 40:7 (4%; nd)

PC 38:6 (38%; 16:0/22:6), 36:6 (17%; 14:0/22:6), 40:9 (9%; nd), 40:10 (5%; 18:4/22:6), 34:5 (5%; nd), 44:12 (5%; 22:6/22:6), 37:6 (4%;15:0/22:6), 42:11 (4%; 20:5/22:6), 40:11 (3%; 18:5/22:6)

Glycolipids

MGDG 36:9 (22%; 18:4/18:5), 30:2 (19%; 14:0/16:2), 30:1 (19%; 14:0/16:1), 36:10 (17%; 18:5/18:5), 32:1 (7%; 14:0/18:1 and/or 16:0/16:1),30:3 (7%; 14:1/16:2), 36:8 (3%; 18:4/18:4)

DGDG 30:2 (30%; 14:0/16:2), 32:5 (14%; 14:0/18:5), 30:1 (12%; 14:0/16:1), 36:10 (11%; 18:5/18:5), 34:7 and/or 33:0 (8%, nd), 32:1 (8%;14:0/18:1 and/or 16:0/16:1), 32:4 (7%; 14:0/18:4)

SQDG 30:2 (28%; 14:0/16:2), 30:0 (26%; 14:0/16:0), 30:1 (24%; 14:0/16:1), 28:0 (17%; 14:0/14:0), 32;5 (5%; nd)

Betaine lipids

DGTS 34:5 (15%; nc), 34:4 (7%; nc), 28:0 (6%; 14:0/14:0), 36:5 (6%; nc), 32:1 (6%; nc), 34:2 (5%; nc), 32:4 (5%; nc), 36:4 (5%; nc), 34:1(5%; nc), 36:2 (4%; nc), 30:1 (4%; nd), 32:2 (4%; nd), 34:3 (4%; nd), 36:6 (4%; nc), 33:0 and/or 34:7 (4%; nd) 34:6 (3%; nd), 34:8(2%; nc)

DGTA 44:12 (28%; 22:6/22:6), 38:6 (15%; 16:0/22:6), 44:11 (8%; nd), 42:11 (8%; 20:5/22:6), 40:10 (6%; 20:5/20:5), 36:6 (5%; 14:0/22:6),38:5 (5%, nd), 34:5 (4%, nd), 39:0 and/or 40:7 (3%; nd), 34:2 (2%, nc), 36:6 (2%; 16:0/20:5), 30:1 (2%; nd), 36:2 (1%; 18:1/18:1),28:0 (1%, nd), 34:4 (1% nc)

DGCC 38:6 (52%; 16:0/22:6), 44:12 (30%; 22:6/22:6), 36:6 (18%; 14:0/22:6)

nd = no MS3 data available to determine probable fatty acid pairs.nc = no consensus on fatty acids pairs (i.e. many different fatty acid pairs).



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3.9. Results of PCA2

The second PCA (PCA2) was intended to increase ourunderstanding of the controls on the composition of diglyc-eride moieties. Given that diglyceride moieties of specificclasses of IP-DAGs contain specific numbers of carbonatoms anddouble bonds andthat specific IP-DAGs are asso-ciated with specific groups of plankton, we elected not to in-clude cell abundances in PCA2. In addition to thenumbers of diglyceride carbon atoms and double bonds for each class of IP-DAGs, the only environmental data we included were thetemperatures andthe depths of thewaterswe sampled. Thesetwo environmental variables are the canonical physical vari-ables known to effect changes in the chain-length and unsat-uration of fatty acids by microorganisms in order to maintainmembrane fluidity (Sinensky, 1974; DeLong and Yayanos,1985; Bartlett and Bidle, 1999).

Temperature was significantly positively loaded on thefirst principal component (PCA2-1) while depth was signif-icantly negatively loaded on the second principal compo-nent (PCA2-2; Fig. 10). PCA2-1 and PCA2-2 accountedfor 42% and 24%, respectively, of the variability in the dataused in PCA2.

4. DISCUSSION

Bacterial and eukaryotic membrane lipids are composedprimarily of IP-DAGs. Harvey et al. (1986) confirmed thefinding by White et al. (1979) that phospholipids outsideof living cells are readily degraded by microbes and are ex-tremely labile. Importantly, Harvey et al. (1986) also ob-served that the degradation of phospholipids yielded bothfree fatty acids and lyso-phospholipids (i.e. phospholipidsminus one fatty acid) indicating that cleavage of the acylmoieties at the reactive ester bond (i.e. phospholipase A1and A2 pathways) was an important, if not dominant, modeof degradation for these molecules. Since this ester bond iscommon to all IP-DAGs, and phospholipases A1 and A2

Principal component 1 (PCA2-1; 42%)

-1.0 -0.5 0.0 0.5 1.0

P r i n c i p a l c o m p o n e n

t 2 ( P C A 2 - 2 ; 2 4 % )

-1.0

-0.5

0.0

0.5

1.0MGDG

SQDG

SQDG

DGCC

DGTA

DGTS

DGTS

PE

PG

PC

T

Depth

Fig. 10. Results of PCA2. The hollow symbols represent thevariance in number of diglyceride carbon atoms; filled symbolsrepresent the relative number of diglyceride double bonds.

M G D G

D G D G

S Q D G

P G

P E

P C

D G T S

D G T A

D G C C

28

30

32

34

36

38

40

42

44

46

A c y l c a r b o n a t o m s

Mean

Std.Dev.

Range

aa

b

c

d

e

f

g

a,c

Fig. 8. Distribution of the number of acyl carbon numbers indifferent classes of IP-DAGs. Letters indicate statistically distinctgroups ( p < 0.05) as indicated by ANOVA and post-hoc Tukeytest.

M G D G

D G D G

S Q D G

P G

P E

P C

D G T S

D G T A

D G C C

0

2

4

6

8

10

12

14

A c y l d o u b l e b o n d s

Mean

Std.Dev.

Range

aa

a,d

b

c

d

ee

f

Fig. 9. Distribution of the number of acyl double bonds in differentclasses of IP-DAGs. Letters indicate statistically distinct groups( p < 0.05) as indicated by ANOVA and post-hoc Tukey test.



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are known to be effective against both phospholipids andnon-phosphorus lipids (Matos and Pham-Thi, 2009), it isreasonable to expect that all classes of IP-DAGs we ana-lyzed were relatively labile (although it is unlikely that theresidence time of all classes of IP-DAGs outside if livingcells were identical to one another). Thus we have madethe critical assumption that living plankton were the pri-

mary sources of IP-DAGs in our samples. As such, wehypothesized that higher concentrations of IP-DAGs wouldbe observed where concentrations of microbial cells werehighest. This hypothesis appears to be supported: we ob-served a general west-to-east increase in IP-DAGs alongour cruise transect (Figs. 3–5), which corresponded to thegeneral increases in microbial carbon (Grob et al., 2007;Ras et al., 2008). There was significant correlation betweenthe classes of IP-DAGs ( p < 0.05 for 31 out of 36 possiblepair-wise comparisons; not shown), and a multivariateexploratory statistical method was required in order toidentify higher-order trends in the dataset. Since, to a firstapproximation, IP-DAGs are expected to scale linearly

with cell abundance, PCA was deemed appropriate forexamining the relationships between the abundance of dif-ferent classes of IP-DAGs and different classes of microbialcells.

4.1. Planktonic sources of phospholipids

Of the three major types of IP-DAGs (Table 1), phospho-lipids have received the most attention in the marine watercolumn, but, despite this, knowledge of the planktonic ori-gins of the specific classes of phospholipids is still limited(Suzumura, 2005). The result of PCA1 showed that the con-centrations of all three classes of phospholipids, and theabundances of heterotrophicbacteria,eukaryotic picophyto-

plankton, and Synechococcus were all significantly related toone another (Fig. 7). The phospholipids PG and PE are thesole classes of IP-DAGs in Pelagibacter ubique (Van Mooyet al., 2009), the cultivated representative of theSAR11 clade(Rappé et al., 2002), which dominates open ocean environ-ments (Morris et al., 2002; Carlson et al., 2009). In addition,PG and PE have long been known to dominate the mem-branes of other, more readily cultivatable strains of marineheterotrophic bacteria (Oliver and Colwell, 1973). Marinestrains of anoxygenic aerobic photoheterotropic bacteriaalsomaybe richin PG(Van Mooy etal.,2009), andbacterio-chlorophyll analyses showed that these cells were presentthroughout the transect (Ras et al., 2008). Anoxygenic aero-bic photoheterotropic bacteria are not distinguished fromstrictly heterotrophic cells by flow cytometry and are ac-counted for in heterotrophic bacterial cell counts. Sinceapproximately 8 out of every 10 microbial cells were hetero-trophic bacteria along the cruise transect (Grob et al., 2007),we assert that the linkage between PG, PE and heterotrophicbacteria cells identified by PCA1 is the result of the domi-nance of these twoclasses of phospholipids in themembranesof heterotrophic and photoheterotrophic bacteria. Thisassertion is supported by the fatty acid composition of PGand PE: these two classes of IP-DAG were the only onesfound to contain odd-chain fatty acids, which are prevalentinbacteria(Table 2). Furthermore, the long-chainpolyunsat-

urated fatty acids (i.e. 20:5 and 22:6) that are abundant ineukaryotic phytoplankton (e.g. Volkman et al., 1989) wereabsent in PG and relatively minor PE (Table 2).

While heterotrophic bacteria may have been the domi-nant source of PG and PE in our samples, they were almostcertainly not the sole source. PG is found in cyanobacteriaand eukaryotic phytoplankton, where it is an essential com-

ponent of the thylakoid membranes (Sato et al., 2000b;Gombos et al., 2002) that house the light-dependent reac-tion centers required for photosynthesis. Recent work onthe crystal structure of photosystem II (PSII), the photosys-tem where water is oxidized to dioxygen, indicates that PGis essential for its assembly, stability and maintenance (Gus-kov et al., 2009). It has also been shown that PSII com-plexes devoid of PG have impaired electron transportactivity (Sakurai et al., 2006) and that PG deficient mutantsof cyanobacteria and other autotrophic bacteria displaymarked photophysiological deficiencies (Benning and Som-erville, 1992; Gombos et al., 2002). Yet despite its impor-tance, PG is a comparatively minor component of

thylakoid membranes (Wada and Murata, 1998) and gener-ally composes only a few percent of total IP-DAGs in pic-ocyanobacteria (Van Mooy et al., 2006) and even less ineukaryotic picophytoplankton (Van Mooy et al., 2009).In eukaryotes, PE often functions as an intermediate inthe synthesis of PC (Nelson and Cox, 2000; Yang et al.,2004). Consistent with this, PE is generally either several-fold less abundant than PC in eukaryotic algae or entirelyabsent (Eichenberger and Gribi, 1997; Kunzler and Eichen-berger, 1997; Bigogno et al., 2002; Khozin-Goldberg andCohen, 2006), but there are some exceptions (Janero andBarrnett, 1981). As mentioned above, we observed that afew of the less abundant PE species contained long-chainpolyunsaturated fatty acids (Table 2), and thus it is possible

that a small fraction of PE we observed in our samples wascontributed by eukaryotic plankton.

As with PG and PE, the phospholipid PC was positivelyloaded on PCA1-1. PC is consistently the dominantphospholipid in the eukaryotic phytoplankton we haveexamined (Van Mooy et al., 2009). Furthermore, PC hasbeen reported to be a major IP-DAG in zooplankton, fromprotozoa, to copepods, to krill (e.g. Patton et al., 1972;Mayzaud et al., 1999; Lund and Chu, 2002). So althoughthe abundances of protists and larger zooplankton werenot included in PCA1, they, in addition to eukaryotic phy-toplankton, undoubtedly contributed to the concentrationsof PC we observed along the cruise transect. A eukaryoticsource for PC is further evidenced by the fatty acid compo-sition of PC, which invariably contained the 22:6 fatty acidderived from eukaryotes. Van Mooy et al. (2008) were ableto attribute as much as 14.9% of phospholipid synthesisalong this cruise transect to phytoplankton, and thuseukaryotic phytoplankton could easily be an importantsource of PC. Marine photoheterotrophic bacteria, whichare common in the open ocean (Koblı́žek et al., 2007),may also contain abundant PC (Van Mooy et al., 2006).However, ornithine lipids, which are also present in photo-heterotrophic bacteria, were scarce in our samples (VanMooy et al., 2009). Although the diagnostic pigment forphotoheterotrophic bacteria, bacteriochlorophyll a, was



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generally three orders of magnitude less than chlorophyll afrom phytoplankton (Ras et al., 2008). Therefore, at thistime we can reasonably assert that the concentrations of PC we observed on our transect were more strongly influ-enced by contributions from eukaryotic phytoplanktonand zooplankton than by bacteria.

Comparing the concentrations of the three phospholip-

ids with concentrations of and turnover rates of particulatephosphorus determined by Duhamel et al. (2007), providesinsights on the lability of phospholipids. On average, thesum of all three phospholipids accounted for 4 ± 1% of the total particulate phosphorus, and there were no obviousspatial trends in the individual values of this percentage.This percentage is well within the ranges reported from pre-vious studies, where phospholipid concentrations weredetermined using different methods (Suzumura and Ingall,2001, 2004; Suzumura, 2005). We had previously reportedthat phospholipid synthesis accounted for an average of 14 ± 5% of total phosphate uptake by plankton (Van Mooyet al., 2008). As the latter percentage is higher than the for-

mer, phospholipids clearly turned over at rates that are sub-stantially faster than other molecular pools of planktonicphosphorus. This result is consistent with the long-standingview that phospholipids are among the most labile bio-chemicals in the sea.

4.2. Planktonic sources of betaine lipids

Betaine lipids were originally thought to be present onlyin eukaryotes (Dembitsky, 1996), yet are now also recog-nized to also be present in bacteria (Benning et al., 1995;Geiger et al., 1999; López-Lara et al., 2003). However, com-pared to phospholipids and glycolipids, there are compara-bly fewer reports of betaine lipids in the microbiological

literature. At this point, culture studies suggest that betainelipids are ubiquitous in eukaryotic phytoplankton (Katoet al., 1996; Van Mooy et al., 2009), which could explainthe significant positive loading of all three betaine lipidsalong with eukaryotic phytoplankton on PCA1-1. Yet beta-ine lipids were recently reported in the aphotic zone of theBlack Sea (Schubotz et al., 2009). This could reflect inputsfrom phytoplankton in the euphotic zone, but if betaine lip-ids do indeed originate primarily from living biomass, thenthis observation suggests that these molecules are derivedfrom some source other than phytoplankton; Schubotzet al. (2009) argue that the presence of odd-chain fatty acidsin these betaine lipids from the aphotic zone are indicativeof a bacterial source for these IP-DAGs. Indeed, we ob-served that heterotrophic bacterial abundances were alsosignificantly loaded on PCA1-1 (Fig. 7).

DGTS is the only betaine lipid that has been reported inbacteria; gene sequences similar to those encoding the BtaAprotein that catalyzes the defining step of DGTS synthesisare fairly widespread among the a-Proteobacteria (López-Lara et al., 2003), but so far as we know betaine lipidsthemselves have only been found in two species of bacteria:Rhodobacter sphaeroides (Benning et al., 1995) and Sinorhi-zobium meliloti (Geiger et al., 1999). Both species are pho-toheterotrophic bacteria, however, based on previousresearch conducted in the Sargasso Sea, we have argued

that heterotrophic bacteria are unlikely sources for betainelipids in the oligotrophic open ocean (Van Mooy et al.,2009). As far as we know, betaine lipids have never beenobserved in any of the cyanobacterial genera that dominatethe ocean (Van Mooy et al., 2009), although DGTS hasbeen reported in epi- and hypolithic cyanobacteria fromLake Kinneret and the Dead Sea (Ř ezanka et al., 2003).

We suggest that Synechococcus clustered with the betainelipids (and phospholipids) in PCA1 because Synechococcustends to be more abundant in higher nutrient waters thatalso favor eukaryotic phytoplankton (Cavender-Bareset al., 2001). Synechococcus were minor components of the microbial community throughout the transect (Grobet al., 2007) and thus were unlikely to contribute signifi-cantly to the total concentrations of IP-DAGs.

In an exhaustive survey of betaine lipids in algal cul-tures, Kato et al. (1996) suggested that DGTS was confinedto green alga. Indeed, the pronounced peak in DGTS at adepth of 50 m at Stations 18 (Fig. 4), corresponded with apeak in chlorophyll b, which is a diagnostic pigment for

green algae (Ras et al., 2008). However, other work hasshown the presence of DGTS in other classes of phyto-plankton (reviewed by Dembitsky, 1996), and, for example,we have observed traces of DGTS in the PrymnesiophyteEmiliana huxleyi (Van Mooy et al., 2009). Although DGTScontained a broad diversity of fatty acids (Table 2), thenumber of carbon atoms and double bonds were similarto both the PE and MGDG; as stated above, PE was likelyto be of bacterial origin, and MGDG, which was likelydominated by input from cyanobacteria and eukaryoticphytoplankton (Figs. 8 and 9). Although we have dis-counted heterotrophic bacteria and cyanobacteria assources of betaine lipid, the broad distribution of fatty acidsin DGTS cast some doubt on the interpretation that DGTS

is primarily from eukaryotic phytoplankton. Future work isclearly required to more definitively determine the source orsources of DGTS in the upper ocean.

Kato et al. (1996) found DGTA to be present in culturesof Prymnesiophytes and Cryptophytes, but the pigmentassociated with the latter group, alloxanthin, was absentin the samples we examined (Ras et al., 2008) and thusCryptophytes were unlikely to be a major source of DGTA.DGTA was generally about twice as abundant as DGTS,and peaks in DGTA at Stations 17 and 18 (Fig. 4) couldhave been derived from the diatoms that were abundantthere (Gómez et al., 2007). We found DGTA in the diatomChaetoceros gracilis (Van Mooy et al., 2009), and DGTAwas the only IP-DAG that contained significant 20:5 fattyacid, which can be the dominant fatty acid in diatoms(Volkman et al., 1989; Zhukova, 2004). The greater abun-dance of longer-chain fatty acids in DGTA compared toDGTS, has been consistently observed in eukaryotic phyto-plankton (Dembitsky, 1996). So although it is difficult tomake any concrete conclusions on the origins of DGTAin our samples, known sources of DGTA, along withobserved fatty acid contents, suggest that eukaryoticphytoplanktonic are an important source of this class of IP-DAG.

Literature on the origin of DGCC is very sparse. DGCCwas originally isolated and described from Pavlova lutheri ,



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a prymnesiophyte alga (Kato et al., 1994). We have ob-served it to be abundant in the prymnesiophyte E. huxleyi ,but also in the diatom Thallasiosira pseudonana (Van Mooyet al., 2009). We observed only three different diglyceridemoieties in DGCC molecules along the transect, and eachof these contained 22:6 fatty acids (Table 2), which are par-ticularly dominant lipids in E. huxleyi (Volkman et al.,

1989; Pond and Harris, 1996). Based on reported distribu-tions of 19’-hexanoyloxyfucoxanthin and fucoxanthin (Raset al., 2008), prymnesiophytes were present in relativelyhigher abundance than diatoms throughout the transect.Thus we posit that prymnesiophytes were an importantsource of DGCC.

4.3. Planktonic sources of glycolipids

In contrast to the phospholipids and betaine lipids, theglycolipids MGDG and SQDG were not significantlyloaded on PCA1-1 and instead were significantly loadedon PCA1-2 (Fig. 7). These glycolipids are among the canon-

ical “

chloroplast lipids”

(Wada and Murata, 1998), andthus it is not unexpected that chlorophyll a concentrationswere also significantly loaded on PCA1-2. The profiles of MGDG and SQDG show maximums at the DCM at Sta-tions 11 and 15; SQDG also shows a peak at 50 m at Sta-tion 11 (Fig. 5). These depths are marked by maximumsin the concentrations of zeaxanthin and divinyl chlorophylla, which are derived from Prochlorophytes (Ras et al.,2008). Similarly, Grob et al. (2007) reported maximums inthe cell abundances of Prochlorococcus at these locationsin the water column. While the other lipids generally in-creased in concentration from west to east, MGDG andSQDG lipid are strikingly less abundant at Stations 17and 18 compared to Station 15 (Figs. 3–5). So although

these chloroplast lipids are undoubtedly present in eukary-otic phytoplankton, our PCA1 results suggest that Prochlo-rophytes are the dominant sources of these lipids in theeastern subtropical South Pacific. Furthermore, we havenoticed SQDG to be present in only vanishingly small con-centrations in some eukaryotic phytoplankton (Van Mooyet al., 2009), and both 20:5 and 22:6 fatty acids were absentin both MGDG and SQDG.

Although the glycolipid DGDG is also among thecanonical chloroplast lipids, the concentration profiles of DGDG, were much more similar to those of phospholipidand betaine lipids than to SQDG or MGDG ( Figs. 3–5).We suggest that contributions from organisms other thanProchlorophytes impacted the distribution of DGDG onour transect. Indeed, DGDG has been linked with thesynthesis of DGTS in some algae (Khozin-Goldberg andCohen, 2006). Yet the fatty acid composition of DGDGis more similar to SQDG and MGDG than any other classof IP-DAG. So although picocyanobacteria were the ex-pected source of DGDG (Wada and Murata, 1998), the pri-mary planktonic sources of DGDG are unclear.

4.4. Controls on SQDG:PG and BL:PC ratios

Phytoplankton are known to substitute non-phosphorusSQDG and betaine lipids for PG and PC, respectively,

when dissolved phosphate in their environment is scarceand becomes limiting (Benning et al., 1995; Sato et al.,2000a; Van Mooy et al., 2009). We have shown previouslythat shifting SQDG:PG and BL:PC ratios in phytoplank-ton is reflected in the SQDG:PG and BL:PC ratios of thewhole planktonic community, despite the contributions of heterotrophic bacteria to PG and PC. Since, phosphate

concentrations at our stations were relatively high (Moutinet al., 2008), and did not limit primary or bacterial produc-tion (Bonnet et al., 2008; Van Wambeke et al., 2008), wewould not predict SQDG:PG and BL:PC ratios in phyto-plankton to vary considerably. For the most part, the re-sults from the upper euphotic zone (


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lipids and PC, are more sensitive to phosphate limitationthan the SQDG-containing Prochlorophytes. Yet all threephospholipids and betaine lipids were significantly loadedon the same principle component (PCA1-1) and were inde-pendent of phosphate concentrations. Thus, variation incommunity structure is the most likely source of the ob-served variation in BL:PC with depth.

4.5. Controls on composition of diglyceride moieties

When examining samples at the community level, thedistribution of fatty acids associated with each class of IP-DAGs is constrained by the spectrums of fatty acidsproduced by the different species of plankton that contrib-ute to the sample. Indeed, even within genera, there isstrong genetic control on the distributions of fatty acidsavailable for attachment to IP-DAG headgroups (Chiet al., 2008). Yet at the species level, the canonical controlson fatty acid composition are temperature and pressure;organisms respond to changes in these to variables by alter-

ing their fatty acid composition in order to maintain mem-brane fluidity (Sinensky, 1974; DeLong and Yayanos, 1985;Wada et al., 1994; Bartlett and Bidle, 1999; Fang et al.,2000). Although our sampling was focused primarily inthe surface mixed layer and bracketed a fairly small rangeof both of these parameters, we sought to understandwhether temperature and pressure affected the distributionof fatty acids within classes of IP-DAGs. There was clearvertical structure in IP-DAG distributions at each of ourstations (Figs. 3–5), and, thus, the timescales of mixing wereclearly longer than timescales of IP-DAG synthesis. There-fore, there is no a priori reason to expect that vertical seg-regation in fatty acid composition due to temperature andpressure effects could not also be possible.

If pressure and temperature impacted only the composi-tion of fatty acids, then we might expect that this effectshould manifest itself consistently across all classes of IP-DAGs. However, PCA2 did not yield any clustering patterncommon to all of the IP-DAGs (Fig. 10), suggesting a lackof common temperature or pressure control on fatty acidcomposition. Since higher pressure and lower temperatureshould affect greater unsaturation but shorter chain-length,we expected that to these two properties of the IP-DAGsshould cluster in opposite quadrants of the PCA2 diagram.Yet both the chain-length and unsaturation of each class of IP-DAG tended to cluster together. This result suggeststhat the average chain-length and unsaturation of IP-DAGswas an inherent biochemical property of each class of IP-DAGs that was unaffected by temperature or pressure.However, lipid phase transition temperatures may differby as much as 20 C between different classes of IP-DAGs,even if they possess identical fatty acids (reviewed by Wil-liams, 1998). So it is possible that plankton were regulatingmembrane fluidity through shifts in relative abundances of different classes of IP-DAGs; we were not able to isolatethis effect, if it was present, from the large vertical varia-tions in community structure by using the statistical meth-ods employed in this study.

Our examination of variations in the average number of diglyceride carbon atoms and double bonds showed that

the classes of IP-DAGs tended to contain characteristicdiglyceride moieties (Figs. 8 and 9). It is likely that the dif-ferences in the fatty acids between IP-DAGs has at leastsome physiological basis: within a single species of planktondifferent classes of IP-DAGs almost invariably contain dif-ferent distributions of fatty acids (Vogel and Eichenberger,1992; Kato et al., 1995; Gombos et al., 1996; Khozin-Gold-

berg and Cohen, 2006). However, the same class IP-DAGscan harbor different distributions of fatty acids in differentspecies of plankton (e.g. Vogel and Eichenberger, 1992;Khozin-Goldberg and Cohen, 2006). Thus, in addition toplankton physiology, the fatty acids in IP-DAGs are alsoa reflection of community composition. We can only spec-ulate on how the balance of intracellular and intercellularprocesses affected the fatty acid composition of IP-DAGswe observed on our cruise. Yet it is clear that previouslypublished profiles of “total fatty acids” from the oceanare a reflection of mixtures of IP-DAGs containing distinctdistributions of fatty acids.

5. CONCLUSIONS

IP-DAGs in the surface waters of the eastern South Pa-cific were dominated by three phospholipids, three betainelipids and three glycolipids; concentrations of all 9 majorclasses of IP-DAGs were generally in the range of a fewhundred picomolar. A principle components analysis of IP-DAG concentrations, analyses of the constituent fattyacids contained in the IP-DAGs, and review of available lit-erature allowed us to tentatively identify broadly-definedsources of most of the 9 major classes of IP-DAGs. PGand PE appeared to be derived primarily from heterotro-phic bacteria, while Prochlorophytes were dominant con-tributors to SQDG and MGDG. Owing in part to the

abundance of long-chain polyunsaturated fatty acids,DGTA, DGCC and PC appeared to originate primarilyfrom eukaryotic plankton. DGTS is also likely to derivefrom eukaryotic phytoplankton, although it contained abroader and more ambiguous distribution of fatty acids.Yet, it is important to recognize that these aforementionedlinks between classes of IP-DAGs and classes of planktonare based only on simple correlations between the abun-dances of different groups of plankton, the concentrationof IP-DAG classes, and fatty acid compositions of IP-DAGs. There is substantial biological diversity within thesegroups of plankton, and it is likely that the various differentmembers of these groups contribute IP-DAGs in varyingdegrees, which may depend on as of yet undiscovered con-nections to environmental conditions. Finally, the potentialfor microzooplankton to contribute to variations in IP-DAG distributions in the water column was not considered,primarily for want of data from our cruise or from culturestudies.

Many of the classes of IP-DAGs we encountered on ourcruise contained distributions of diglyceride moieties thatwere distinct from one another. It has been recognized sincethe inception of fatty acids research in the oceans that spe-cific types of plankton contribute varying proportions of specific fatty acids to the total pool of fatty acids in envi-ronmental samples. Our results show that the total pool



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of fatty acids should also be regarded as a mixture of fattyacids from classes of IP-DAGs with characteristic distribu-tions of diglyceride moieties.

ACKNOWLEDGMENTS

We offer our gratitude to the numerous investigators who con-

tributed to the comprehensive, high-quality, publically-availableBIOSOPE cruise database. Specifically we thank T. Moutin, S.Duhamel, P. Raimbault, C. Grob, O. Uloa, and J. Ras for obtain-ing the data we used in the this paper. We also thank H. Claustreand A. Sciandra for the invitation to join the cruise and for leadingthe BIOSOPE project. The assistance of the crew of the R/V L’Ata-lante was invaluable, as was the camaraderie with T. Moutin, O.Prášil, M. Twardowski and F. Van Wambeke. The acquisition of BIOSOPE data was funded through Centre National de RechercheScientifique – Institut National des Sciences de l’Univers grants.Comments on an earlier draft of this manuscript by L. Hmelo,K. Longnecker, and K. Popendorf were invaluable, as was the in-put for three anonymous reviewers. Participation in this cruise byB.V.M. was supported by the WHOI Ocean Life Institute. Analysisof the samples from this cruise was supported by a grant to B.V.M.

from the U.S. National Science Foundation (OCE-0646944).

APPENDIX A. SUPPLEMENTARY DATA

Supplementary data associated with this article can befound, in the online version, at doi:10.1016/j.gca.2010.08.026.

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