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http://www.elsevier.com/locate/permissionusematerial Sachs J.P. (2014) Hydrogen Isotope Signatures in the Lipids of Phytoplankton. In: Holland H.D. and Turekian K.K.

(eds.) Treatise on Geochemistry, Second Edition, vol. 12, pp. 79-94. Oxford: Elsevier.

© 2014 Elsevier Ltd. All rights reserved.

12.4 Hydrogen Isotope Signatures in the Lipids of PhytoplanktonJP Sachs, University of Washington, Seattle, WA, USA

ã 2014 Elsevier Ltd. All rights reserved.

12.4.1 Introduction 7912.4.2 The Effect of dDwater on dDlipid 8012.4.3 The Effect of Biosynthesis on dDlipid 8212.4.4 The Effect of Species on dDlipid 8312.4.5 The Effect of Salinity on dDlipid 8312.4.6 The Effect of Temperature on dDlipid 8712.4.7 The Effect of Growth Rate on dDlipid 8912.4.7.1 Substrate-Limited Growth Rate Effects 8912.4.7.2 Light-Limited Growth Rate Effects 9212.4.8 Summary and Conclusions 92Acknowledgments 92References 92

12.4.1 Introduction

Key Points

• Experimental evidence indicates that the following factors influencealgal lipid dD values. The magnitude of the effect based on publishedstudies is indicated in parentheses:

• dDwater• Species (up to 160%)

• Lipid type (up to 170%)

• Salinity (þ0.9"0.2% per PSU)

• Growth rate (0 to #30% per day)

• Temperature (#2 to #8% per degree Celsius)

• Based on these findings, we provide the following guidance to thoseinterested in applying algal lipid dD values in paleoclimate or paleoen-vironmental studies:

• Use lipids unique to a family or species.

• Be mindful that changes in salinity will cause changes in lipid dDvalues that will be additive to those associated with changes inprecipitation and evaporation.

• Expect lipid dD values to decrease if temperature or growth rateincreases.

The ability to rapidly measure deuterium-to-hydrogenratios in small quantities of individual lipids is less than15 years old and has already led to significant advances inEarth science. Pioneered by John Hayes and Alex Sessions inthe late 1990s and early 2000s (Burgoyne and Hayes, 1998;Hayes, 2001; Sauer et al., 2001; Sessions, 2001; Sessions et al.,1999, 2001), compound-specific D/H analyses have begun tocontribute substantively to our understanding of lipid biosyn-thesis in bacteria (Sessions et al., 2002), algae (Zhang andSachs, 2007) and higher plants (Sessions, 2006), catabolismin bacteria (Valentine et al., 2004), trophic relationships inmarine food webs (Chikaraishi, 2006), climate during theQuaternary (Hu et al., 2003; Pahnke et al., 2007; Schefuszet al., 2005, 2011) and the Eocene (Pagani et al., 2006), sedi-ment transport in the ocean (Englebrecht and Sachs, 2005), thesource and fate of petroleum hydrocarbons (Pond et al., 2002),forensic science, pharmacology, and archaeology (Bensonet al., 2006), to name a few. And despite a burgeoning interestin applying lipid D/H ratios in these fields, there is much we do

not understand about the environmental factors that influenceD/H fractionation in eukaryotes and prokaryotes and the bio-chemical and biophysical processes within cells that cause D/Hfractionation. This article focuses on the hydrogen isotopesignatures in lipids produced by phytoplankton becausethose are the signatures most commonly used to evaluateenvironmental conditions in the past (see Chapter 12.5).

Because all biosynthetic products in phytoplankton andcyanobacteria appear to be depleted in deuterium relative toenvironmental water, ferredoxin–NADP reductase (FNR) is alikely source of D/H fractionation. This enzyme on the thyla-koid membrane in photosystem I (PS1) catalyzes the reduc-tion of NADPþ to NADPH. As such, it represents the firstopportunity newly liberated protons from the oxidation ofwater in PS2 can experience mass-based fractionation.NADPH thus produced is used to reduce CO2 in the Calvincycle, causing any fractionation by FNR to be reflected down-stream in all biomolecules. Since hydrogen derived fromNADPH is likely to be depleted in deuterium by #250% ormore (Luo et al., 1991; Schmidt et al., 2003), it is expectedthat lipids and other biosynthetic products will be depleted indeuterium compared to environmental water. Any change ingrowth conditions has the potential to alter D/H fraction-ation in algal cells by affecting the relative rates at whichPS2 (supply of Hþ) and PS1 (demand for Hþ) operate.Subsequent D/H fractionation during the transfer of H#

from NADPH to phosphoglyceric acid (PGA) and hydrogenexchange with cellular H2O will diminish the initialD-depletion (Hayes, 2001; Yakir, 1992; Yakir and Deniro,1990). Changes in nutrients, light, temperature, or salinitywill alter demand for Hþ by PS1, H# by PGA, and theexchange of hydrogen with cellular H2O, providing ampleopportunities for D/H fractionation.

Well established at this point is the fact that D/H ratios inalgal lipids are highly correlated with the D/H ratios of thewater in which the phytoplankton grew (Englebrecht andSachs, 2005; Sachs and Schwab, 2011; Sauer et al., 2001;Schouten et al., 2006; Schwab and Sachs, 2011; Zhang andSachs, 2007; Figure 1). Also well established is the fact that alllipids in autotrophic phytoplankton are depleted in deuterium

Treatise on Geochemistry 2nd Edition http://dx.doi.org/10.1016/B978-0-08-095975-7.01007-X 79

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by approximately 100–400% relative to the water in whichthey grew, owing to kinetic isotope effects (KIEs) during lipidsynthesis (Sessions et al., 1999). Furthermore, differing by100% or more can be different lipids produced by the samealga and the same lipid produced by different genera (Sessionset al., 1999; Zhang and Sachs, 2007). The biosynthetic steps inwhich the D/H fractionation occurs are just starting to beworked out (Sessions, 2006; Sessions et al., 2002; Zhang andSachs, 2007) but remain largely unknown. Of equal or greaterimportance for the application of hydrogen isotope techniquesin paleoceanography, geochemistry, and ecology are the envi-ronmental influences on the magnitude of that fractionation.Here again, the influence on D/H ratios in algal lipids of themost basic environmental parameters such as temperature,salinity, light levels, and nutrient concentrations remainslargely unknown.

Though some progress has been made in characterizing andunderstanding the hydrogen isotope signatures of lipids inphytoplankton, there remains much more that we do notknow than what we do know. Below is a review of the current

state of knowledge of the D/H signatures in algal lipids. First,we discuss the relationship between the hydrogen isotopiccomposition of environmental water and lipids, both withina single species and between different species. Next, we discussthe influence of water salinity on lipid D/H ratios, followed bya discussion of the effects of temperature, nutrients, light levels,and growth rates. We conclude with some thoughts on themost pressing and promising questions to be addressed withhydrogen isotopes in algal lipids.

12.4.2 The Effect of dDwater on dDlipid

Experiments with freshwater (Zhang and Sachs, 2007) andmarine (Englebrecht and Sachs, 2005) phytoplankton grownin batch culture indicate that lipid dD values (dDlipid) arehighly correlated with water dD values (dDwater). Zhang andSachs (2007) grew five species of freshwater green algae inbatch culture, including Eudorina unicocca, Volvox aureus, andthree strains of Botryococcus braunii (two A race, one B race),

–300

–200

–100

0

100

–100 100 300 500 –100 100 300 500

B. braunii, A racealkadienes

C27 C29 C31

–300

–200

–100

0

100

200

E. huxleyialkenones

Alk

adie

ne δ

D (‰

)

Alk

adie

ne δ

D (‰

)

Water δD (‰) Water δD (‰)

y = 0.726x-213.768R2= 0.999

y = 0.648x-210.707R2= 1.000

(a) (b)

–350

–330

–310

–290

–270

–50 –40 –30 –20 –10 0

(c)

Chesapeake Baydinosterol

Din

oste

rol δ

D (‰

)

Water δD (‰)

y = 1.87x -258R2= 0.95

Figure 1 dD values of (a) C27, C29, and C31 alkadienes in Botryococcus braunii cultures, (b) C37 alkenones in Emiliania huxleyi cultures, and (c)dinosterol in the Chesapeake Bay all track water dD values very closely. Data from Englebrecht and Sachs, 2005; Sachs and Schwab, 2011; Zhang andSachs, 2007.

80 Hydrogen Isotope Signatures in the Lipids of Phytoplankton



under controlled conditions in media containing five differentconcentrations of deuterium. The hydrogen isotopic ratios oflipids in the algae, including alkadienes, botryococcenes, hep-tadecenes, fatty acids, and phytadiene, were correlated withwater dD values with R2 values in excess of 0.99 for all lipidsin all species (Figure 2).

Similarly, Englebrecht and Sachs (2005) grew the marinecoccolithophorid Emiliania huxleyi in batch cultures with fivedifferent deuterium enrichments (Figure 3). The hydrogenisotope composition of C37 and C38 alkenones was linearlycorrelated with dDwater with R2 values greater than 0.99.A similar batch culture experiment with E. huxleyi performedby Paul (2002) and reported in Schouten et al. (2006) pro-duced essentially the same result (Paul, 2002; Schouten et al.,2006). The combined data sets are shown in Figure 3.

Since a wide diversity of lipids from six species of phyto-plankton, representing both marine and freshwater realms,and from both the acetogenic and the isoprenoid biosynthetic

pathways, have dDlipid values that are linearly correlated withenvironmental water dD values, it is reasonable to expect thatthis will be the case for all algal lipids under most circum-stances. Field studies across the environmental gradient fromseawater to river water and across large biogeographic zonescorroborate this supposition.

Experiments performed in the Chesapeake Bay (CB) estuary(Maryland and Virginia, USA) indicate that the dD values of fourdifferent alkenones (both C37 and C38 di- and triunsaturatedvarieties) were correlated with the dD value of water across asalinity gradient of 19withR2 values of 0.7–0.8 (Figure 4; Schwaband Sachs, 2009, 2011). In addition, the dD values of the dino-flagellate sterol dinosterol (4a,23,24-trimethyl-5a-cholest-22E-en-3b-ol) were correlated with the dD values of water across asalinity gradient of 19 PSUwith anR2 valueof 0.9 (Figure 5; Sachsand Schwab, 2011). The consistently high correlation betweendDwater and dDlipid in an estuarine setting inwhich the assemblageof alkenone- and dinosterol-producing phytoplankton is likely to

(Martinque)

dD o

f bot

ryoc

occe

nes

(‰)

dD o

f nat

ural

ly o

ccur

ing

FAM

E (‰

)

dD o

f phy

tadi

enes

(‰)

dD o

f fre

e fa

tty

acid

s (c

orre

cted

, ‰)

dD of water at harvest (‰)

dD of water at harvest (‰) dD of water at harvest (‰)

dD of water at harvest (‰)

C30boty = 0.751x-277.948

(C30 bot)

y = 0.770x-148.222(C20:1)

y = 0.746x -165.893(C28:1)

y = 0.557x-282.096

y = 0.715x -162.069(C16)

y = 0.810x-153.424(C20:1)

y = 0.807x-167.963(C28:1)

y = 0.791x-169.872(C16)

C16

C18:1

C28:1

C20:1

y = 0.726x-329.067 (C34 bot [4])

y = 0.647x-359.821 (C34 bot iso?)

y = 0.730x-297.445(C31 bot)

C34bot(4)

C34bot(5) + ?

C34bot iso

C31bot

C32 + C33 bot

200

100

0

-100

-200

200

300

100

0

-100

-200

100

0

-100

-200

-300

200 C16

C18:1

C28:1

C20:1

300

(a) (b)

(d)(c)

100

0

-100

-200

0 100 200 300 400 500

-300

-400400 5003002001000

400 5003002001000 400 5003002001000

Figure 2 dD values of a variety of lipids from the green alga Botryococcus braunii (B race) grown in batch cultures were highly correlated with thedD value of the water. Reproduced from Zhang Z and Sachs JP (2007) Hydrogen isotope fractionation in freshwater algae: I. Variations among lipidsand species. Organic Geochemistry 38: 582–608.

Hydrogen Isotope Signatures in the Lipids of Phytoplankton 81



vary attests to the fact that isotopic changes of water are stronglyimprinted on lipids in phytoplankton.

Two other studies looked at the relationship betweendDwater and the dD value of lipids common to many algalspecies in surface sediments from lakes across large biogeocli-matic zones in western Europe (Sachse et al., 2004) and theeastern United States (Huang et al., 2002, 2004). In theEuropean lake survey, Sachse et al. (2004) found a high corre-lation between the dD value of the C17 normal alkane (nC17)and the dD value of climatological average rainfall. In the studyof eastern US lakes, Huang et al. (2002) found a high correla-tion between the dD values of palmitic acid (nC16:0), nC17, andphytol in surface sediments and lake water dD values across adiversity of climates and limnologies (Figure 6). However,these lipids are produced by many different plankton andhigher plants, so their sources in sediments are likely to bemixed. A final example of the near-universality of the imprintof water dD values on algal lipids comes from the fact that eventotal lipid extracts (TLEs) from a wide diversity of freshwater

and saline lakes have dD values that are highly correlated withdDwater (Figure 7) (Nelson and Sachs, unpublished).

12.4.3 The Effect of Biosynthesis on dDlipid

Much of the difference in the slopes and intercepts of therelationships depicted in Figures 1–6 can be attributed to thebiosynthetic pathway from which particular lipids are produced,an observation first made by Estep and Hoering (1980)(Estep and Hoering, 1980) and expanded upon by Sessionset al. (1999). Lipids produced by the two isoprenoid path-ways, 1-deoxyxylulose 5-phosphate/2-C-methyl-D-erythritol 4-phosphate (DOXP/MEP) and mevalonic acid (MVA), havebranched carbon chains (e.g., sterols, phytol, botryococcenes,and phytadienes) and are depleted in deuterium by approxi-mately 100–200% relative to lipids synthesized by the aceto-genic pathway which have a straight chain of carbon atoms(e.g., fatty acids, alkenones, alkadienes, and the leaf wax

Englebrecht and Sachs (2005)300

-10 90 190

dD37:2= 0.724 ! dDH2O-226R2= 0.991 (n = 13)

290

dDH2O (vs. VSMOW)

dDC

37:2

alk

enon

e (v

s. V

SM

OW

)

390 490 590

200

100

0

-100

-200

-300

Paul (2002)

Figure 3 dD values of alkenones versus dD values of growth water from batch culture experiments with the marine coccolithophorid E. huxleyi. Dataare from Englebrecht and Sachs (2005) and Paul (2002), as reported in Schouten et al. (2006).

y = 0.84x-156, R2= 0.80

y = 0.79x-173, R2= 0.72

y = 0.75x-179, R2= 0.75

y = 0.98x-183, R2= 0.80-240

-220

-200

-180

-160

-140

-10-20-30-40

Water dD (‰)

Alk

enon

e dD

c37:3

c37:2

C38:3

C38:2

Figure 4 dD values of four C37 alkenones in suspended particles were highly correlated with dD values of water along a $175 km section ofthe Chesapeake Bay estuary. Reproduced from Schwab VF and Sachs JP (2011) Hydrogen isotopes in individual alkenones from the ChesapeakeBay estuary. Geochimica et Cosmochimica Acta 75: 7552–7565.




n-alkanes, n-alkanols, and n-alkanoic acids) (Sauer et al., 2001;Sessions et al., 1999; Zhang and Sachs, 2007; Figures 8 and 9).

KIEs in multiple steps within a lipid biosynthetic pathwaycontribute to the observed net D-depletion in lipids. Theseinclude hydrogenation (by NADPH and H2O), desaturation,decarboxylation, elongation, and methylation, among others.The proton in NADP(H) is estimated to have a dD value of#250 to #600% relative to environmental water, so any stepin which hydrogen derived from NADPH is added will causeisotopic depletion of the product (Hayes, 2001; Luo et al., 1991;Schmidt et al., 2003). This was demonstrated by Chikaraishiet al. (2009) by determining the dD values of phytol and itsprecursors in cucumber cotyledons (Chikaraishi et al., 2009).Desaturation of fatty acids and alkenones results in unsaturatedproducts that are substantially depleted in D relative to precur-sors (Chikaraishi et al., 2004; D’andrea et al., 2007; Schwab andSachs, 2009, 2011) (cf. Figure 4) implying a large KIE associated

with some desaturases. The same was inferred for some decar-boxylase and methylase enzymes involved with lipid synthesis.For example, decarboxylation of the C30:1 fatty acid to producethe C29 alkadiene in B. braunii (A race) resulted in a D-depletionof the product by as much as 40%, and the C31–C34 botry-ococcenes in cultured B. braunii (B race) were all depleted inD relative to the C30 homologue from which they are synthe-sized (Zhang and Sachs, 2007). Elongation of carbon chainsmay cause some D/H fractionation but is likely to be minorcompared to the previously discussed processes (Chikaraishiet al., 2004; Zhang and Sachs, 2007).

12.4.4 The Effect of Species on dDlipid

It is the sum total of the aforementioned (and other) KIEs thatdictate the dD value of algal lipids. Because there are a series ofenzyme-mediated reactions leading to the synthesis of anylipid, the physiological state of a cell will almost certainlyinfluence the extent to which each individual KIE is expressedin the final product. The type and relative abundance of differ-ent biochemicals in different species of algal cells are alsoexpected to result in different isotopic depletions of a particularlipid relative to environmental water. For example, under sim-ilar batch culture growth conditions, two species of Chloro-phyceae (E. unicocca and V. aureus) and three species ofTrebouxiophyceae (B. braunii) produced palmitic acid (nC16:0

fatty acid) that differed by 160% relative to water (Zhang andSachs, 2007; Figure 10).

In addition to the magnitude of the KIE expressed in eachstep of lipid synthesis differing between species, it is possiblethat the enzymes used for a particular reaction can differbetween species, as is the case for fatty acid desaturase enzymesin eukaryotic phytoplankton and cyanobacteria (Chi et al.,2008). Ubiquitous lipids such as palmitic acid (C16:0), with amultitude of aquatic and terrestrial sources, may therefore notbe good targets for D/H-based paleohydrologic reconstructions.

12.4.5 The Effect of Salinity on dDlipid

A decrease in D/H fractionation with increasing salinity wasfirst observed by Schouten et al. (2006) in alkenones fromcultured coccolithophorids (E. huxleyi and G. oceanica) overthe salinity range 25–35 PSU. Our field data from ChristmasIsland and the CB support that finding.

A decrease in D/H fractionation (increase in a) as salinitiesincreased occurred in several algal and cyanobacterial lipids inpondsonChristmas Islandwith salinities of 17–149 PSU in June–July 2005 (Sachse and Sachs, 2008; Figure 11). Fractionationfactors (a) for TLEs were between 0.797 and 0.908 and covariedwith salinity according to the relationship a¼0.0007*Sþ0.79(R2¼0.74, n¼32), indicating a 0.7% decrease in D/H fraction-ation per unit increase in salinity (Sachse and Sachs, 2008).Fractionation factors for individual lipids increased in concert(Figure 11). Slopes of the regression of a onto salinity for thenC17 alkane from algae (Sachse et al., 2004), diploptene frombacteria (Elvert et al., 2001; Wakeham, 1990), and phytenefrom algae and/or bacteria (Boudou et al., 1986; Hefter et al.,1993; Schouten et al., 2001) were similar to one another

y = 1.87x-258R2= 0.95

y = 1.35x-284R2= 0.86

Sediment

Water dD (‰)

Din

oste

rol d

D (‰

)

-45-350

-330

-310

-290

-270

-35 -25 -15 -5

Particles

Figure 5 dD values of the dinoflagellate sterol dinosterol were linearlycorrelated with water dD values in the Chesapeake Bay estuary whenmeasured in both suspended particles (circles) and surface sediments(diamonds). Reproduced from Schwab VF and Sachs JP (2011)Hydrogen isotopes in individual alkenones from the Chesapeake Bayestuary. Geochimica et Cosmochimica Acta 75: 7552–7565.

-80 -60 -40 -20 0Water dD (‰, VSMOW)

Pal

miti

c ac

id d

D (‰

, VS

MO

W) dDPA=-167.0 + 0.939 ! dDH2O

R = 0.894

20 40-100-300

-260

-220

-180

-140

-100

Figure 6 dD value of palmitic acid from surface sediments of a transectof lakes in the eastern United States (Huang et al., 2002).




(0.00076–0.0011) and to the subset of TLEs from which the bio-markers were purified, 0.00083 a units per PSU salinity(Figure 11). The higher y-intercept (or a at zero salinity) fornC17 relative to phytene and diploptene in Figure 11 is consistentwith the widely noted D-depletion in isoprenoid lipids relative toacetogenic lipids (Hayes, 2001; Sessions et al., 1999; Zhang andSachs, 2007). A y-intercept for the TLEs that is intermediatebetween acetogenic nC17 and isoprenoid phytene and diplopteneis expected since TLEs contain a mixture of the two lipid types(Buhring et al., 2009).

In the CB, dD values of dinosterol, a dinoflagellate sterol(Volkman et al., 1998), also indicate that D/H fractionation

decreases linearly as salinity increases (Sachs and Schwab,2011). The slope of the regression of a onto salinity was0.00099 in suspended particles and significant at the 95% con-fidence level (Figure 12). This translates into a decrease in D/Hfractionation of 0.99% per PSU increase in salinity. This sensi-tivity of D/H fractionation to salinity is similar to those forphytene (1.1% per PSU), nC17 alkane (0.80% per PSU), andTLEs (0.70% per PSU) from Christmas Island. Regardless of thebiosynthetic pathway, environment, or source of the lipid, theD/H fractionation associated with salinity appears to be rela-tively constant in the diverse settings of the CB estuary and thehypersaline ponds on Christmas Island.

The higher slope of the linear regression of adino/water ontosalinity (Figure 4(b)) in sedimentary dinosterol relative toparticulate dinosterol can be explained by high dinoflagellateproduction during summer when river flows and runoff arelowest. The particulate dinosterol and water samples weretaken during high flow conditions in springtime when surfacewater dD values and salinities in the CB are expected to berelatively low compared to the summer. In other words, thewater overlying sediments in May is likely to be characterizedby lower dDwater and salinity values than would typically occurin summer when dinoflagellate production peaks. Annual flux-weighted dDdino values are thus expected to be higher thandDdino values measured in May in response to the combinedinfluence of higher dDwater values and diminished D/H frac-tionation in higher salinity water during summer.

Unlike dinosterol, D/H fractionation in alkenones fromsuspended particles in the CB was unchanged as salinityincreased from 10 to 29 PSU (Schwab and Sachs, 2011; Fig-ure 13). We have no satisfactory explanation for the differentbehavior of alkenones relative to alkenones in cultured cocco-lithophorids, dinosterol in the CB, or a variety of lipids insaline ponds on Christmas Island. One possibility is that achange in the species of alkenone producers occurs along thelength of the estuary such that a change in the magnitude ofD/H fractionation imparted to alkenones by the differentspecies cancels out the effect of salinity on D/H fractionation.

y = 0.9389x-151.44R2= 0.8189

-250.00

-240.00

-230.00

-220.00

-210.00

-200.00

-190.00

-180.00

-100.00 -80.00 -60.00 -40.00dD Water

dD T

LE

Figure 7 dD values of total lipid extracts (TLEs) from surface sediments in 12 saline lakes from around the globe are well correlated with dD values ofthe lake water. Salinities of all lakes were less than 60 ppt (Nelson and Sachs, unpublished).

Ale

xand

rium

Isoc

hrys

is

Asc

ophy

llum

n-alkyl C skeletons

dD (‰

)

Isoprenoid C skeletonsSterolsSesqui- and triterpenoidsPhytol and phytene

n-alkanesn-alcohols

-400

-300

-200

-100

0

Fatty acids

Fucu

s

Zost

era

Spa

rtin

a

Dau

cus

Met

hylo

cocc

us

Figure 8 Acetogenic lipids (open symbols) are usually enriched inD relative to isoprenoid lipids in a variety of plants, algae, and bacteria.Reproduced from Sessions AL, Burgoyne TW, Schimmelmann A, andHayes JM (1999) Fractionation of hydrogen isotopes in lipidbiosynthesis. Organic Geochemistry 30: 1193–1200.




The magnitude of the species effect in batch cultures of twococcolithophorids (E. huxleyi and Gephyrocapsa oceanica) wasabout 30% in Schouten et al. (2006), which, coincidentally, isjust the right magnitude to counter a 1% decrease in D/Hfractionation per unit increase in salinity in the CB if theassemblage of alkenone producers was dominated by E. huxleyiat low salinity, and G. oceanica at high salinity. Another possi-ble explanation for the lack of change in a for alkenones inthe CB estuary is that greater osmoregulation capacity incoastal haptophytes may result in a diminished sensitivity ofalkenone–water D/H fractionation to salinity changes.

A linear decrease in D/H fractionation as salinity increases isclear from the Christmas Island saline pond lipids, the CBdinosterol, and Schouten et al. (2006)’s coccolithophorid cul-ture experiments. Unexplained is why Schouten observed athreefold greater sensitivity of D/H fractionation to changes in

salinity in alkenones from batch culture (3.3% per PSU) thaneither of our field studies indicate for a wide diversity of lipids(0.7–1.1% per PSU). The most likely explanation for this dis-crepancy is that different growth rates for the coccolithophoridsat different salinities in the batch cultures imparted additionalD/H fractionation in the alkenones. This effect is discussed indetail later in Section 12.4.7.

The mechanism or mechanisms by which increasingsalinity causes decreasing D/H fractionation between waterand lipids in algae and cyanobacteria remains unknown.Controlled experiments with continuous cultures are neededso that all growth and environmental conditions can be heldconstant while salinity is varied. Such experiments are under-way in our laboratory at the University of Washington, andpresumably in other laboratories. In the meantime, we offeredthree hypotheses to explain why increased salinity resulted in

Acetogenic

Water dD (‰)

300

300

Palmitic acid

Phytadiene

C16:0 FAME

C34 botryococcene

C30 botryococcene

5004002001000

200

100

0

-100

-200

170‰

• 170‰ differencein D/H fractionationfor different lipids in

B. braunii (B race)

-300

-400

Lipi

d dD

(‰)

Isoprenoid

Figure 9 Acetogenic lipids (C16:0 fatty acid and C16:0 fatty acid methyl ester) from B. braunii (B race) were enriched in deuterium by about 170%relative to isoprenoid lipids (botryococcenes and phytadiene). Adapted from Zhang Z and Sachs JP (2007) Hydrogen isotope fractionation in freshwateralgae: I. Variations among lipids and species. Organic Geochemistry 38: 582–608.

160‰

400

300

200

100

0

-100

100 200 300dD of water at harvest (‰)

dD o

f C16

fatt

y ac

ids

(cor

rect

ed, ‰

)

400 500

160‰ differencein palmitic acid

between families ofgreen algae

V. Aureus (15 "C)

V. Aureus (25 "C)B. Braunii (A race)B. Braunii (B race)

E. unicocca

0

-200

Figure 10 The dD value of a single lipid, palmitic (C16:0) acid, varied by 160% among five families of freshwater green algae. Adapted from Zhang Zand Sachs JP (2007) Hydrogen isotope fractionation in freshwater algae: I. Variations among lipids and species. Organic Geochemistry 38: 582–608.




decreased D/H fractionation between algal lipids and extracel-lular water in Sachs and Schwab (2011) and Sachse and Sachs(2008). They are as follows:

Hypothesis 1 Higher salinities cause decreased water trans-port across the cell membrane, resulting in greater recyclingof internal water, and an increasingly D-enriched internal

water pool from which lipids are synthesized (Kreuzer-Martinet al., 2006; Sachse and Sachs, 2008).

Water transport across the cell membrane via aquaporinsand Naþ channels is restricted upon exposure to high salinitiesto prevent Naþ toxicity (Boursiac et al., 2005; Pomati et al.,2004), while at the same time, the volume of water in cyano-bacterial cells has been shown to decline by up to 25% duringsalt stress (Allakhverdiev et al., 2000a,b). Lower transport ratesof water across the cell wall and a decreased pool of internalwater would act in concert to continuously enrich the dD valuesof cellular water as NADPH is continuously replenished withD-depleted hydrogen. Estimates of the dD values of hydrogenfor NADPþ protonation range from #250 to #600% (Luoet al., 1991; Schmidt et al., 2003). Lipids (and all other biosyn-thetic products) would reflect the increasingly elevated dDvalues characterizing the internal water. Consequently, it is pos-sible that D/H fractionation between dinosterol and CB waterdecreases from the headwaters to the mouth of the bay as aresult of decreased water transport across dinoflagellate cellmembranes, diminution of their internal water pool, and anincrease in the dD value of internal water as water is recycledrather than replenished from outside the cell, and D-depletedhydrogen is withdrawn for NADPH production.

Hypothesis 2 Elevated salinities cause growth rates to decline,resulting in diminished D/H fractionation between lipids andextracellular water.

Many studies on phytoplankton and algae indicate thatgrowth rates generally decrease upon exposure to elevated salin-ities (Cifuentes et al., 2001; Clavero et al., 2000; Herbst andBradley, 1989). Though different algae are adapted to a range ofoptimal salinities, virtually all experience a decline in growthrate once a threshold salinity is passed. Laboratory culturestudies on the marine diatom Thalassiosira pseudonana (Zhanget al., 2009) and the marine coccolithophorids E. huxleyi and

Salinity

a lip

id–w

ater

00.65

0.7

0.75

0.8

0.85

0.9

0.95

50 100 150

Xmas nC17

y = 0.81 + 0.0008x R2= 0.79

y = 0.76 + 0.0012x R2= 0.95

y = 0.77 + 0.00074x R2= 0.48

y = 0.81 + 0.00069x R2= 0.85

y=0.65 + 0.0011x R2= 0.91

y = 0.72 + 0.00082x R2= 1

y = 0.78 + 0.00076x R2= 0.66

y = 0.68 + 0.00099x R2= 0.57

Xmas nC17:1Xmas nC18Xmas nC18:1Xmas phyteneXmas diplopteneXmas TLECB dinosterol

Figure 11 alipid–water versus salinity relationships for total lipid extracts and lipid biomarkers in Christmas Island saline ponds (open symbols) anddinosterol from the Chesapeake Bay estuary (orange squares). Dinosterol is an isoprenoid lipid from dinoflagellate algae, and nC17 (circles) and nC17:1are acetogenic lipids from algae and cyanobacteria. Diploptene (triangles) is an isoprenoid from bacteria, and phytene (diamonds) is an isoprenoid lipidfrom bacteria and algae. Data from Sachse and Sachs (2008) and Sachs and Schwab (2011).

353025Salinity

20151050.680

0.690

0.700

0.710a

0.720

0.730

0.740Sediment

Suspended particles

a = 0.00174±0.000321 *

a = 0.000990±0.000229 *salinity + 0.685±0.004,R2= 0.57

salinity + 0.685±0.005, R2

= 0.77

Figure 12 adinosterol/water values in suspended particles (filled circles,N¼16) and surface sediment (open circles, N¼11) plotted againstsalinity. Error bars for adino/water represent propagated errors reported fordDdino and dDwater measurements. Linear regressions for suspendedparticles (solid lines) and sediments (dotted lines) shown with 95%confidence intervals. Reproduced from Schwab VF and Sachs JP (2011)Hydrogen isotopes in individual alkenones from the Chesapeake Bayestuary. Geochimica et Cosmochimica Acta 75: 7552–7565.




G. oceanica (Schouten et al., 2006) indicate substantial decreasesin D/H fractionation expressed in a variety of lipids as growthrates are reduced, either through purposeful nitrogen limitation(Zhang et al., 2009) or some combination of salinity and tem-perature (Schouten et al., 2006). If growth rates of dinosterol-producing dinoflagellates declined along the length of the CB, itis possible that D/H fractionation between dinosterol and CBwater decreased in concert.

Hypothesis 3 Upon exposure to high salinities, rapid produc-tion of osmolytes draws D-depleted hydrogen from an internalpool of water, leaving it, and subsequent lipids synthesizedfrom it, enriched in deuterium.

When subjected to osmotic stress, cells from all domains oflife accumulate organic solutes to counteract the externalosmotic pressure (Borowitzka and Brown, 1974; Borowitzkaet al., 1977; Brown, 1978; Galinski, 1995; Grant, 2004; Mackayet al., 1984; Reed and Stewart, 1985; Reed et al., 1986; Roberts,2004, 2005; Ventosa et al., 1998). Also called osmolytes orcompatible solutes (because they provide osmotic balance with-out interfering with the metabolic functions of the cell; Ventosaet al., 1998), these small organic molecules span a wide diversityof structures and can be zwitterionic, uncharged, or anionic(Roberts, 2005). Furthermore, they can represent a very substan-tial fraction of cellular hydrogen, reaching up to 10–20% of thedry weight of cells in some hypersaline bacteria (Ventosa et al.,1998). If hydrogen is shuttled off from internal pools of waterand NADPH to rapidly synthesize osmolytes in response toosmotic stress, residual pools of those substances would be leftenriched in deuterium, and all subsequent biosynthetic productswould reflect this enrichment. Thus, if dinoflagellates produceincreasingly greater concentrations of osmolytes as salinitiesincrease seaward in the CB, their internal pools of water andNADPH might become increasingly D-enriched, as would thedinosterol synthesized from that water, resulting in less D/Hfractionation relative to CB water.

Both hypotheses 1 and 3 call on a change in the internalwater dD value as salinity changes and would thus predict thatall biosynthetic products would experience a decrease in D/Hfractionation relative to external water since water is the sourceof hydrogen for all biochemicals in phytoplankton. Hypothe-sis 2 calls upon a salinity-induced reduction in growth rate asthe source of lowered D/H fractionation in dinosterol (andother lipids) relative to water, but does not necessarily predictthat all biosynthetic products be increased in dD. These aretestable hypotheses that future work ought to address.

12.4.6 The Effect of Temperature on dDlipid

Hydrogen isotope fractionation in algal lipids appears toincrease as temperature increases. Zhang et al. (2009) showedthat acetogenic lipids from two species of freshwater greenalgae (E. unicocca and V. aureus) grown in batch culture at15 &C were enriched in deuterium by 20–40% relative tothose grown at 25 &C (Zhang et al., 2009; Figure 14). The lipidsincluded palmitic acid (C16:0) in both species (Figure 14(a)),the naturally occurring methyl esters of palmitic acid in bothspecies, stearic acid (C18:0) and its naturally occurring methylester in E. unicocca, and heptadecene (C17:1ene) in E. unicocca(Figure 14(b)). Yet with only two temperatures investigated, itwould be imprudent to generalize these results.

Further support for an increase in D/H fractionation inalgal lipids as temperature increases comes from two indepen-dent laboratory culture studies with marine coccolithophorids(Figure 15). Schouten et al. (2006) cultured E. huxleyi in batchcultures at three temperatures and three salinities and found anincrease in D/H fractionation (decrease in alpha) in alkenonesof about 0.4–1% per degree Celsius increase in temperature(Figure 15). The sensitivity of D/H fractionation to temperatureappears to have been even larger in alkenones from G. oceanica,in which dD values decreased by 3–4% per degree Celsius

302826242220Salinity

MeC37:3 in sediments




and filters

and filters

and filters

and filters

181614121080.800

Alk

enon

e–w

ater

D/H

frac

tiona

tion

0.810

0.820

0.830

0.840

0.850

0.860

Figure 13 Hydrogen isotope fractionation between four alkenones and environmental water (aalkenone/water) as a function of measured surfacewater salinity in the Chesapeake Bay estuary, May 2006. Suspended particle values are in black. Surface sediment values are in gold. No influenceof salinity on D/H fractionation in alkenones is observed in this data. Reproduced from Schwab VF and Sachs JP (2011) Hydrogen isotopes inindividual alkenones from the Chesapeake Bay estuary. Geochimica et Cosmochimica Acta 75: 7552–7565.




increase in temperature (Figure 15). This is the same tempera-ture sensitivity we observed in lipids from E. unicocca andV. aureus cultures (see Figure 14). Although Schouten et al.(2006)’s G. oceanica alkenone data are from just two tempera-tures, they are compelling because three cultures were grown ateach of those temperatures, one at each of three salinities, andbecause an independent set of threeG. oceanica batch cultures byWolhowe et al. (2009) yields a similarly high temperaturesensitivity. D/H fractionation in alkenones increased by 5%(i.e., alpha decreased by 0.0049) per degree Celsius in thatstudy (Figure 15) (Wolhowe et al., 2009).

Contradicting these two studies are the results from the CBwhere no change was observed in apparent D/H fractionation infour different alkenones over the surfacewater temperature rangeof 15&–20 &C (Schwab and Sachs, 2011; Figure 16). But as dis-cussed in the section on salinity, where it was shown that nochange in D/H fractionation between alkenones and water was

evident in the CB (Figure 13), other factors may be affecting thehydrogen isotope trends in alkenones along the Bay, such as achange in the assemblage of alkenone producers along the salin-ity gradient.

Zhang et al. (2009) discuss three mechanisms by whichtemperature may influence D/H fractionation during lipid syn-thesis: its influence on enzyme activities, KIEs, and hydrogentunneling. Lipid biosynthesis is catalyzed by a variety of enzymeswhose conformation facilitates reaction with substrates. Whenthe enzyme is in the native (active) state, the reaction rateincreases with temperature. Temperature changes may inducethe (partial) replacement of an enzyme by an isoenzyme withbetter heat- or cold-tolerance (Steele and Fry, 2000). Jahnke et al.(1999) reported that the soluble methane monooxygenase(sMMO) and particulate methane monooxygenase (pMMO)isozymes expressed different carbon isotopic fractionation fac-tors (5% difference) in methanotrophic bacteria using the

Temperature ("C) Temperature ("C)

EU-C18FAEU-C18FAME

EU-C17:1ene EU-C16FAMEVA-C16FAME

a Pal

miti

c ac

id–w

ater

a Lip

id–w

ater

Eudorina Volvox

y =-0.0033x + 0.9887y =-0.0039x + 0.9608

y =-0.0022x + 0.9798y =-0.0038x + 0.9947y =-0.0037x + 0.9885y =-0.0035x + 0.9792

y =-0.0039x + 1.0138

10(a) (b)

0.85

0.88

0.91

0.94

0.97

0.85

0.88

0.91

0.94

0.97

15 20 25 30 10 15 20 25 30

Figure 14 D/H fractionation was higher (i.e., lower a values) at 25 &C than at 15 &C in lipids from cultured green algae E. unicocca (closedsymbols) and V. aureus (open symbols). (a) C16:0 (palmitic) acid in both species. (b) nC17:1 alkene in Eudorina (squares) and several fatty acids andnatural fatty acid methyl esters in both species. The slope of the lines suggests an isotope enrichment 3–4% per degree Celsius. Adapted from Zhang Z,Sachs JP, and Marchetti A (2009) Hydrogen isotope fractionation in freshwater and marine algae: II. Temperature and nitrogen limited growth rateeffects. Organic Geochemistry 40: 428–439.

y= −0.0012x+ 0.79 R2= 0.19

y= −0.0008x+ 0.80 R2= 0.94

y= −0.0004x+ 0.82 R2= 0.43

y= −0.0028x+ 0.80

y= −0.0043x+ 0.84

y= −0.0028x+ 0.83

y= −0.0049x+ 0.89 R2= 0.97

0.73

0.74

0.75

0.76aal

keno

ne–w

ater

0.77

0.78

0.79

0.8

0.81

0.82

7 12 17

Temperature (°C)22 27

Eh S= 25

Eh S= 29

Eh S= 35

Go S= 25

Go S= 29

Go S= 35

Wol Go

E. huxleyi

G. oceanica

Figure 15 D/H fractionation in C37 alkenones from the cultured marine coccolithophorids Emiliania huxleyi and Gephyrocapsa oceanica from thestudies of Schouten et al. (2006) (colored symbols) and Wolhowe et al. (2009) (black symbols). Closed symbols are from E. huxleyi. Open symbols arefrom G. oceanica. Schouten et al. (2006) performed experiments at three salinities (S ¼ 25, 29, and 35). Taken together, the data imply a decrease inalkenone dD values of 0.4–5% per degree Celsius increase in water temperature.




ribulose monophosphate pathway for carbon assimilation.However, after passing some optimum temperature that is spe-cific to each enzyme, denaturalization can occur, making theenzyme lose its effectiveness as a catalyst. In this way, the set ofenzymes involved in the synthesis of a particular lipid, theirsensitivity to temperature, and any D/H fractionation theyimpart will influence the dD value of the lipid.

Isotope fractionation in biochemical processes arises fromunequal zero-point energies of bonds between heavy and lightisotopes, resulting in different activation energies (Bigeleisenand Wolfsberg, 1959), which in turn are influenced by tem-perature. Fractionation factors are thus a function of both thetemperature and the activation energy of the individualenzyme-catalyzed reactions that comprise the lipid biosyn-thetic pathway.

Yet the greater D/H fractionation we observed at higher tem-perature is at odds with theory that indicates that equilibriumfractionation should decrease as temperature increases (Clarkand Fritz, 1997). In any complex organism, changing growthtemperature could lead to a whole suite of metabolic changes,any one of which could result in the observed isotopic differ-ences. For example, different enzymes (isoenzymes or isozymes)may be used to synthesize lipids at different temperatures, eachwith its own fractionation (Jahnke et al., 1999). In one study bySteele and Fry (2000), two isoenzymes of xyloglucan endotrans-glycosylase (XET) isolated from Arabidopsis (i.e., cauliflowerflorets) were found to be temperature-dependent, one with anoptimum temperature of $12 &C and exhibiting 55% of itsmaximal catalytic rate at #5 &C, total XET activity of mixed iso-enzymes with a temperature optimum of $30 &C (Steele andFry, 2000). Assuming those two enzymes exhibit different iso-tope effects, the temperature at which the organism exists wouldcause different D/H ratios in the lipid product.

Other possible causes for a temperature influence on D/Hfractionation in lipid synthesis include different mechanismsfor synthesizing NADPH at different temperatures, such asphotosynthesis versus the pentose phosphate pathway

(Kruger and von Schaewen, 2003), and hydrogen tunnelingdue to strengthened substrate–enzyme complex vibration atelevated temperatures (Kohen et al., 1999).

12.4.7 The Effect of Growth Rate on dDlipid

12.4.7.1 Substrate-Limited Growth Rate Effects

The only published study of which I am aware that has con-trolled growth rate in order to investigate its influence on D/Hfractionation in lipids is Zhang et al. (2009). Other studiesbearing on this question have inferred how different growthrates, brought about by either temperature or salinity varia-tions in batch cultures in the case of Schouten et al. (2006), ordifferent stages of a batch culture by Wolhowe et al. (2009),may have caused changes in D/H fractionation in alkenones.Schouten et al. (2006) calculated growth rates from their tem-perature and salinity experiments and found an increase inD/H fractionation in alkenones as growth rate increased.Wolhowe et al. (2009) looked at the influence of growthstage in batch cultures of E. huxleyi and observed 20–30%greater D/H fractionation in alkenones from cells in the sta-tionary phase relative to the exponential phase of growth(Wolhowe et al., 2009). But these results are not readily com-parable to growth rate experiments, and it is unclear howapplicable they are to the environment. We conducted twocontinuous culture experiments, or chemostats, with themarine diatom T. pseudonana, at constant growth rates con-trolled by N limitation. Our results indicate that there is asubstantial increase in D/H fractionation with increasinggrowth rate in a sterol, an isoprenoid (e.g., branched) lipidsynthesized via the MVA pathway, and a smaller or negligibleincrease in D/H fractionation in fatty acids synthesized via theacetogenic pathway.

Two cultures of T. pseudonana were cultivated at growthrates of 0.4 and 1.9 day#1. With only two data points, weestimate that D/H fractionation increased by 25% per day as

Temperature ("C)

Alk

enon

e–w

ater

D/H

frac

tiona

tion

MeC37:3 in sedimentsMeC37:2 in sediments


MeC38:2 in sedimentsand filters

0.860

0.850

0.840

0.830

0.820

0.810

0.80015 16 17 18 19 20

and filters

and filters

and filters

Figure 16 Hydrogen isotope fractionation between four alkenones and environmental water (aalkenone/water) as a function of measured surface watertemperature in the Chesapeake Bay estuary, May 2006. Suspended particle values are in black. Surface sediment values are in gold. No influence oftemperature on D/H fractionation in alkenones is observed in these data. Reproduced from Schwab VF and Sachs JP (2011) Hydrogen isotopes inindividual alkenones from the Chesapeake Bay estuary. Geochimica et Cosmochimica Acta 75: 7552–7565.




growth rate increased in the sterol 24-methyl-cholesta-5,24(28)-dien-3b-ol but remained constant in the C16:1, C14:0,C16:0, and C18:0 fatty acids (Figure 17). At the same time,there was no change in a for the four fatty acids synthesizedfrom the acetogenic pathway from cultures grown at 0.4 and1.9 day#1 despite the nearly fivefold increase in growth rate(Zhang et al., 2009).

In order for fatty acid dD values to remain constant atdifferent growth rates, the isotopic composition of the hydro-gen source(s) for fatty acid synthesis (1) must either be unaf-fected by growth rate changes, (2) must have stable D/H ratios,such as might occur with a large reservoir of hydrogen, or (3)must compensate in a way that D-depleted hydrogen is addedin some steps and D-enriched hydrogen is added in others. Asreviewed in Zhang and Sachs (2007), fatty acid synthesis canbe divided into three steps: the activation of acetyl-CoA tomalonyl-CoA, chain initiation from a unit of acetyl-CoA plusa unit of malonyl-CoA and subsequent elongation withmalonyl-CoA by a fatty acid synthase (FAS) complex, anddesaturation by desaturase enzyme (Duan et al., 2002).

At the odd-numbered carbon positions, carbon comes fromthe carboxyl group (#C¼O–) of acetate, and hydrogen isderived entirely from NADPH. At the even-numbered carbonpositions, one hydrogen atom is derived from acetate and theother from water during the enoyl-ACP reductase step (Duanet al., 2002; Schmidt et al., 2003). Presently, the precise mech-anism of enzyme-mediated exchange of carbon-bound hydro-gen with intracellular water is not known. If substantialhydrogen exchange occurs between fatty acids and their inter-mediates with a large pool of intracellular water, it couldexplain the insensitivity to growth rate of fatty acid dD values.

Kreuzer-Martin et al. (2006) reported that C14:0 and C16:0

fatty acids exhibited larger deuterium depletions in the logphase of growth than in the stationary phase of growth in thebacterium Escherichia coli and attributed the greater isotopicdepletion to a larger contribution of hydrogen from intracel-lular water during log-phase growth. As a heterotroph, E. colimaintains metabolic water that is more D-depleted than itsextracellular water (Kreuzer-Martin et al., 2006). As a photo-autotroph, T. pseudonana maintains intracellular (or meta-bolic) water enriched in deuterium because isotopicallydepleted hydrogen is continuously removed for NADPH pro-duction (Schmidt et al., 2003), leaving D-enriched metabolicwater. Furthermore, there is no evidence to suggest that theenzymes involved in fatty acid synthesis differ between log andstationary growth phases (Heath and Rock, 1996; Rock andJackowski, 2002). If Kreuzer-Martin et al. (2006)’s hypothesisapplies, fatty acids in T. pseudonana nitrate-replete (NR) cellsought to be less D-depleted than those in nitrate-limited (NL)cells, a prediction not supported by our data. Thus, eitherhydrogen exchange occurs between intracellular water andfatty acids or their intermediates, and/or intracellular waterdD values are insensitive to growth rates.

D/H fractionation in sterols from T. pseudonana variedwidely with nitrogen nutritional status. e values for24-methyl-cholesta-5,24(28)-dien-3b-ol were 37% lower inrapidly growing NR cells than in slow-growing NL cells(Zhang et al., 2009). Sterols, like other isoprenoid lipids, orig-inate from a branched isoprene C5 unit, isopentenyl diphos-phate (IPP), or its isomer, dimethylallyl diphosphate(DMAPP). The ‘eukaryotic’ acetate–MVA pathway starts fromthree acetyl-CoAs and requires six enzymes, two NADPHs, andthree ATPs to produce IPP, which is subsequently converted toDMAPP by IPP isomerase. In contrast, the non-MVA, ‘prokary-otic’ DOXP/MEP pathway of IPP synthesis begins with pyru-vate and glycerinaldehyde-3-phosphate and involves sevenenzymes, three ATP equivalents, and three NADPHs to pro-duce both IPP and DMAPP (Lichtenthaler, 2004).

In higher plants and many algae, such as the Bacillariophytato which T. pseudonana belongs, the two IPP-producingbiochemical pathways operate simultaneously, with the MVApathway restricted to the cytoplasm and the DOXP/MEP path-way restricted to the chloroplast (Armbrust, 2004; Bick andLange, 2003; Hemmerlin et al., 2003; Lichtenthaler, 1999;Schwender et al., 2001; Figure 18). Although sterol synthesisin most plants and algae occurs via the MVA pathway, somealgae, including certain chlorophytes, can use the DOXP/MEPpathway to synthesize sterols and other isoprenoid lipids suchas phytol and carotenoids (Lichtenthaler, 1999; Sato et al.,2003; Schwender et al., 2001; Figure 18).

Other evidence also suggests that under certain conditions,isoprenoid intermediates can be transferred between the cyto-solic MVA pathway and the plastidic DOXP/MEP pathway.Laule et al. (2003) provided indirect support for the presenceof such an export mechanism in the vascular plant Arabidopsisthaliana (Laule et al., 2003). Hemmerlin et al. (2003) reportedthat sterols could be synthesized via the DOXP/MEP pathwaywhen the MVA pathway in Bright Yellow-2 cells was inhibitedand that isoprenoids normally produced in the plastid by theDOXP/MEP pathway could be produced in the cytosol via theMVA pathway when the former was inhibited. This suggested

Growth rate (day–1)

alip

id–w

ater

C16:1 C16:0 C18:0 SterolC14:0

y =-0.025x + 0.715

0.89

0.84

0.79

0.74

0.69

0.640 0.5 1 1.5 2

Figure 17 a vs. growth rate in T. pseudonana. D/H fractionationincreased (a decreased) as specific growth rate increased in the sterol24-methyl-cholesta-5,24(28)-dien-3b-ol (circles). But a remainedconstant in fatty acids at two growth rates. This suggests that D/Hfractionation during lipid synthesis via the MVA biosynthetic pathway issensitive to changes in growth rate but not via the acetogenic pathway.1s error bars are smaller than the symbol in most cases. Adapted fromZhang Z, Sachs JP, and Marchetti A (2009) Hydrogen isotopefractionation in freshwater and marine algae: II. Temperature andnitrogen limited growth rate effects. Organic Geochemistry 40: 428–439.




that significant exchange of isoprenoid intermediates occurredacross the plastid envelope (Hemmerlin et al., 2003). Further-more, Bick and Lange (2003) proposed that plastid mem-branes possess a unidirectional proton/IPP coupled transportsystem for the export of IPP from the plastid to the cytosol(Bick and Lange, 2003).

We thus hypothesize that in fast-growing (NR) cells, IPPand DMAPP monomers from both the MVA and DOXP/MEPpathways are mixed to produce sterols (Figure 18). Forinstance, if the supply of IPP from the cytosolic MVA route isinsufficient for sterol synthesis, additional IPP may come fromthe DOXP/MEP pathway. The metabolic state of the cells coulddictate the relative contribution from the two pathways.Because the last reduction step during IPP synthesis via theDOXP/MEP pathway is characterized by a particularly largehydrogen isotope effect (Zhang and Sachs, 2007), and thusIPPs produced via that pathway are substantially D-depletedrelative to IPPs produced via the MVA pathway, a higher pro-portion of IPP derived from the DOXP/MEP pathway wouldresult in the production of more deuterium-depleted sterols(Figure 18). This could explain the 37% depletion in deute-rium in the fast-growing NR T. pseudonana cells relative to theslow-growing NL cells.

Consistent with our findings, Sessions et al. (1999) reportedthat sterols from dormant plants, which may be analogous to

our slow-growing NL T. pseudonana cells, were deuterium-enriched by 50–100% relative to actively growing plants,whereas fatty acids were substantially less enriched in deute-rium, by 0–30%. Cells in actively growing plants may incorpo-rate some IPPs into sterols from the DOXP/MEP pathwayresulting in more deuterium depletion than in sterols synthe-sized in dormant plants if the MVA pathway alone producessterols. As we found, D/H fractionation in fatty acids observedby Sessions et al. (1999) was not greatly affected by growth rates.

Sessions (2006) attributed the larger D-depletion in lipidsfrom actively growing plants to their faster metabolism. Thismight explain why sterols in faster growing cells are moredepleted in deuterium but does not explain why fatty acidD/H ratios were minimally affected by growth rate in ourexperiments. Because isoprenoid lipids alone expressedincreased D/H fractionation at higher growth rates, it suggeststhat the exchange of isoprenoid precursors (IPPs) between thecytosol and the plastid occurs in rapidly growing cells, but notthe exchange of acetogenic precursors.

Verification of these findings with additional T. pseudonanachemostats andwith cultures of other phytoplankton are requiredbefore firm conclusions can be made regarding the influence ofgrowth rate onD/H fractionation indifferent lipids from the samecell. But taken together, the batch culture experiments ofSchouten et al. (2006) and Wolhowe et al. (2009) and the

Acetyl-CoA

N-limited (NL)/N-replete (NR)

Pyruvate + glyceraldehyde-3-phosphate Acetyl-CoA

HMG-CoA

Mevalonic acid

Mevalonate-5-phosphate

Mevalonate-5-pyrophosphate

-CO2

IPP DMAPP

Lanosterol

24-Methyl-cholesta-5,24(28)-dien-3b-ol

1-Deoxy-D-xylulose 5-phosphate (DOXP)

2-C-methyl-D-erythritol 4-phosphate (MEP)

IPP DMAPP

NAD(P)H

D-depleting step

D-depleting step

Relatively more D-deplete

NLYield: 0.30 pg cell-1

D/H: e =-143.6‰

NL (w/o cross talk)Yield: 0.059 pg cell-1

D/H: e =-294.7‰

NRYield: 0.07 pg cell-1

D/H: e =-142.0‰

NR (w/ cross talk)Yield: 0.074 pg cell-1

D/H: e =-332.0‰

-CO2

b-carotene

Zeaxanthin, etc.

Acetogenic pathway Isoprenoid biosynthesis (I)DOXP/MEP

Isoprenoid biosynthesis (II)MVA pathway

Plastid

Relatively less D-deplete

Now more D-depleted

37‰ more negative

Cytosol

NADPHNADP

+

NAD+

C16 fatty acid

Fatty acidbiosynthesis

Cross talk

Figure 18 Schematic diagram showing the cross talk of IPPs between the plastidic DOXP/MEP pathway and the cytosolic MVA pathway in the marinediatom Thalassiosira pseudonana. The two IPP-producing biochemical pathways operate in parallel, with the MVA pathway housed in the cytoplasm(synthesizing isoprenoids such as b-carotene) and the DOXP/MEP pathway housed in the chloroplast (synthesizing isoprenoids such as sterols). Fattyacid concentrations were fourfold higher in NL (0.30 pg per cell) than in NR (0.07 pg per cell) cells owing to their role as energy-storage compounds.Sterol concentrations were similar in both NL (0.059 pg per cell) and NR (0.074 pg per cell), perhaps owing to their role as components of cellmembranes. Under faster growing conditions, the IPPs from the plastidic DOXP/MEP pathway that produces highly D-depleted products mightcross into the cytosol, providing additional IPPs to the MVA-pathway products that are less D-depleted. As a result, sterols synthesized in the NRculture had dD values that were 37% lower (#332% vs. #295% in the NL culture). Acetogenic compounds such as fatty acids are synthesizedexclusively in the plastid. As a result, they had similar dD values (#144% in NR vs. #142% in NL) despite a fourfold higher concentration in the NLculture. Reproduced from Zhang Z, Sachs JP, and Marchetti A (2009) Hydrogen isotope fractionation in freshwater and marine algae: II.Temperature and nitrogen limited growth rate effects. Organic Geochemistry 40: 428–439.




chemostat culture experiments of Zhang et al. (2009) support thenotion that hydrogen isotope fractionation in some algal lipidsincreaseswith increasing growth rate, but perhapsnot in all lipids.

12.4.7.2 Light-Limited Growth Rate Effects

Light levels also influence growth rates of phytoplankton, butto our knowledge, there are no published studies on the influ-ence of light levels on D/H fractionation in phytoplankton. Inhigher plant leaf waxes, Yang et al. (2009) found that D/Hfractionation was about 40% less under continuous light thanunder a 12 h light–dark cycle (Yang et al., 2009). And in unpub-lished batch culture studies with E. huxleyi and G. oceanica,Benthien et al. (2009) reported a large effect of light intensityon D/H fractionation in alkenones (Benthien et al., 2009). Thatstudy concluded that D/H fractionation in alkenones was highlysensitive to light intensity, which in turn controlled growth rate.Until that work is published, it is impossible to evaluate the rolethat light may play in determining hydrogen isotope ratios inalgal lipids. Future studies should make this a high priority,making use of continuous cultures (chemostats) for controlledexperimentation. Both light and nutrient limitation can controlgrowth rates, but the effect of light onD/H fractionation in lipidsis likely to be different than that of nutrients. Changes in the fluxof photons directly affect the supply of Hþ generated by PS2,rather than its demand by PS1. Steady state growth experimentswith chemostats in which growth rates are separately controlledby light and nutrient levels ought to elucidate the role that eachplays in controlling D/H fractionation in phytoplankton lipids.

12.4.8 Summary and Conclusions

The use of hydrogen isotope ratios in lipids to decipher bio-geochemical and climatic processes is expanding rapidly. Therelative success of these efforts depends on an understanding ofthe environmental conditions that influence D/H fraction-ation. Although much remains unknown about the mecha-nisms of D/H fractionation in phytoplankton and theprimary environmental conditions that influence that fraction-ation, culture and field experiments conducted to date indicatethat D/H fractionation in algal lipids is sensitive to tempera-ture, salinity, and nitrate-limited growth rate. Specifically, theyindicate that D/H fractionation (1) increases with increasingtemperature, (2) decreases with increasing salinity, and (3)increases with increasing N-controlled growth rate inisoprenoid sterols from T. pseudonana, but may be insensitiveto growth rate in acetogenic fatty acids from T. pseudonana.

Acknowledgments

Many of the insights expressed would not have been possiblewithout the tenacity and creativity of my students and postdocsduring the last 10 years. In particular, I would like to acknowl-edge the work of Zhaohui Zhang, Valerie Schwab, RienkSmittenberg, Katharina Pahnke, Dirk Sachse, Amy Englebrecht,and Dan Nelson. This material is based upon work supportedby the National Science Foundation under Grant No. OCE-1027079.

References

Allakhverdiev SI, Sakamoto A, Nishiyama Y, Inaba M, and Murata N (2000a) Ionic andosmotic effects of NaCl-induced inactivation of photosystems I and II inSynechococcus sp. Plant Physiology 123: 1047–1056.

Allakhverdiev SI, Sakamoto A, Nishiyama Y, and Murata N (2000b) Inactivation ofphotosystems I and II in response to osmotic stress in Synechococcus. Contributionof water channels. Plant Physiology 122: 1201–1208.

Armbrust EV (2004) The genome of the diatom Thalassiosira pseudonana : Ecology,evolution, and metabolism. Science 306: 79–86.

Benson S, Lennard C, Maynard P, and Roux C (2006) Forensic applications of isotoperatio mass spectrometry – A review. Forensic Science International 157: 1–22.

Benthien A, van der Meer MTJ, Schouten S, Zondervan I, and Bijma J (2009) The effectof growth rate and light intensity on the hydrogen isotopic composition of long-chain alkenones produced by Emiliania huxleyi. In: ISOCOMPOUND ’09, Potsdam,Germany.

Bick J and Lange BM (2003) Metabolic cross talk between cytosolic and plastidialpathways of isoprenoid biosynthesis: Unidirectional transport of intermediatesacross the chloroplast envelope membrane. Archives of Biochemistry andBiophysics 415(2): 146–154.

Bigeleisen J and Wolfsberg M (1959) Theoretical and experimental aspects of isotopeeffects in chemical kinetics. Advances in Chemical Physics 1: 15–76.

Borowitzka LJ and Brown AD (1974) The salt relations of marine and halophilic speciesof the unicellular green alga, Dunaliella. Archives of Microbiology 96: 37–52.

Borowitzka LJ, Kessly DS, and Brown AD (1977) The salt relations of Dunaliella.Archives of Microbiology 113: 131–138.

Boudou JP, Trichet J, Robinson N, and Brassell SC (1986) Profile of aliphatichydrocarbons in a recent polynesian microbial mat. International Journal ofEnvironmental Analytical Chemistry 26: 137–155.

Boursiac Y, Chen S, Luu DT, Sorieul M, van den Dries N, and Maurel C (2005) Earlyeffects of salinity on water transport in Arabidopsis roots. Molecular and cellularfeatures of aquaporin expression. Plant Physiology 139: 790–805.

Brown AD (1978) Compatible solutes and extreme water stress in eukaryoticmicroorganisms. Advances in Microbial Physiology 7: 181–242.

Buhring SI, Smittenberg RH, Sachse D, et al. (2009) A hypersaline microbial mat fromthe Pacific Atoll Kiritimati: Insights into composition and carbon fixation usingbiomarker analyses and a 13C-labeling approach. Geobiology 7: 308–323.

Burgoyne TW and Hayes JM (1998) Quantitative production of H-2 by pyrolysis of gaschromatographic effluents. Analytical Chemistry 70: 5136–5141.

Chi X, Yang Q, Zhao F, et al. (2008) Comparative analysis of fatty acid desaturasesin cyanobacterial genomes. Comparative and Functional Genomics 2008: 25 pp.http://dx.doi.org/10.1155/2008/284508.

Chikaraishi Y (2006) Carbon and hydrogen isotopic composition of sterols in naturalmarine brown and red macroalgae and associated shellfish. Organic Geochemistry37: 428–436.

Chikaraishi Y, Suzuki Y, and Naraoka H (2004) Hydrogen isotopic fractionations duringdesaturation and elongation associated with polyunsaturated fatty acid biosynthesisin marine macroalgae. Phytochemistry 65: 2293–2300.

Chikaraishi Y, Tanaka R, Tanaka A, and Ohkouchi N (2009) Fractionation of hydrogenisotopes during phytol biosynthesis. Organic Geochemistry 40: 569–573.

Cifuentes AS, Gonzalez MA, Inostroza I, and Aguilera A (2001) Reappraisal ofphysiological attributes of nine strains of Dunaliella (Chlorophyceae): Growth andpigment content across a salinity gradient. Journal of Phycology 37: 334–344.

Clark I and Fritz P (1997) Environmental Isotopes in Hydrogeology. New York, BocaRaton, FL: Lewis Publishers.

Clavero E, Hernandez MM, Grimalt JO, and Garcia-Pichel F (2000) Salinity toleranceof diatoms from thalassic hypersaline environments. Journal of Phycology 36:1021–1034.

D’andrea WJ, Liu ZH, Alexandre MD, Wattley S, Herbert TD, and Huang YS (2007) Anefficient method for isolating individual long-chain alkenones for compound-specifichydrogen isotope analysis. Analytical Chemistry 79: 3430–3435.

Duan J-R, Billault I, Mabon F, and Robins R (2002) Natural deuterium distribution infatty acids isolated from peanut seed oil: A site-specific study by quantitative 2HNMR spectroscopy. ChemBioChem 3: 752–759.

Elvert M, Whiticar MJ, and Suess E (2001) Diploptene in varved sediments of SaanichInlet: Indicator of increasing bacterial activity under anaerobic conditions during theHolocene. Marine Geology 174: 371–383.

Englebrecht AC and Sachs JP (2005) Determination of sediment provenance at driftsites using hydrogen isotopes and unsaturation ratios in alkenones. Geochimica etCosmochimica Acta 69: 4253–4265.

Estep MF and Hoering TC (1980) Biogeochemistry of the stable hydrogen isotopes.Geochimica et Cosmochimica Acta 64: 1197–1206.




Galinski EA (1995) Osmoadaptation in bacteria. Advances in Microbial Physiology37: 272–328.

Grant WD (2004) Life at low water activity. Philosophical Transactions of the RoyalSociety of London: Series B, Biological Sciences 359: 1249–1267.

Hayes JM (2001) Fractionation of carbon and hydrogen isotopes in biosyntheticprocesses. Reviews in Mineralogy and Geochemistry 43: 225–277.

Heath RJ and Rock CO (1996) Roles of the FabA and FabZ b-hydroxyacyl-acyl carrierprotein dehydratases in Escherichia coli fatty acid biosynthesis. Journal ofBiological Chemistry 271: 27795–27801.

Hefter J, Thiel V, Jenisch A, Galling U, Kempe S, and Michaelis W (1993) Biomarkerindications for microbial contribution to Recent and Late Jurassic carbonatedeposits. Facies 29: 93–105.

Hemmerlin A, Hoeffler JF, Meyer O, et al. (2003) Cross-talk between the cytosolicmevalonate and the plastidial methylerythritol phosphate pathways in tobacco brightyellow-2 cells. Journal of Biological Chemistry 278: 26666–26676.

Herbst DB and Bradley TJ (1989) Salinity and nutrient limitations on growth of benthicalgae from two alkaline salt lakes of the western Great Basin (USA). Journal ofPhycology 25: 673–678.

Hu FS, Kaufman D, Yoneji S, et al. (2003) Cyclic variation and solar forcing of Holoceneclimate in the Alaskan subarctic. Science 301: 1890–1893.

Huang Y, Shuman B, Wang Y, and Webb T (2002) Hydrogen isotope ratios of palmiticacid in lacustrine sediments record late Quaternary climate variations. Geology30: 1103–1106.

Huang Y, Shuman B, Wang Y, and Webb T (2004) Hydrogen isotope ratios of individuallipids in lake sediments as novel tracers of climatic and environmental change:A surface sediment test. Journal of Paleolimnology 31: 363–375.

Jahnke LL, Summons RE, Hope JM, and Des Marais DJ (1999) Carbon isotopicfractionation in lipids from methanotrophic bacteria II: The effects of physiology andenvironmental parameters on the biosynthesis and isotopic signatures ofbiomarkers. Geochimica et Cosmochimica Acta 63: 79–93.

Kohen A, Cannio R, Bartolucci S, and Klinman JP (1999) Enzyme dynamics andhydrogen tunnelling in a thermophilic alcohol dehydrogenase. Nature399: 496–499.

Kreuzer-Martin HW, Lott MJ, Ehleringer JR, and Hegg EL (2006) Metabolic processesaccount for the majority of the intracellular water in log-phase Escherichia coli cellsas revealed by hydrogen isotopes. Biochemistry 45: 13622–13630.

Kruger NJ and von Schaewen A (2003) The oxidative pentose phosphate pathway:Structure and organisation. Current Opinion in Plant Biology 6: 236–246.

Laule O, Furholz A, Chang H-S, et al. (2003) Crosstalk between cytosolic andplastidial pathways of isoprenoid biosynthesis in Arabidopsis thaliana. Proceedingsof the National Academy of Sciences of the United States of America100: 6866–6871.

Lichtenthaler HK (1999) The 1-deoxy-D-xylulose-5-phosphate pathway of isoprenoidbiosynthesis in plants. Annual Review of Plant Physiology and Plant MolecularBiology 50: 47–65.

Lichtenthaler H (2004) The 1-deoxy-D-xylulose-5-phosphate pathway of isoprenoidbiosynthesis in plants. Annual Review of Plant Physiology and Plant MolecularBiology 50: 47–65.

Luo Y-H, Steinberg L, Suda S, Kumazawa S, and Mitsui A (1991) Extremely low D/Hratios of photoproduced hydrogen by cyanobacteria. Plant and Cell Physiology32: 897–900.

Mackay MA, Norton RS, and Borowitzka LJ (1984) Organic osmoregulatory solutes incyanobacteria. Journal of General Microbiology 130: 2177–2191.

Pagani M, Pedentchouk N, Huber M, et al. (2006) Arctic hydrology during globalwarming at the Palaeocene/Eocene thermal maximum. Nature 442: 671–675.

Pahnke K, Sachs JP, Keigwin LD, Timmermann A, and Xie S-P (2007) Eastern tropicalPacific hydrologic changes during the past 27,000 years from D/H ratios inalkenones. Paleoceanography 22. http://dx.doi.org/10.1029/2007PA001468.

Paul HA (2002) Application of Novel Stable Isotope Methods to ReconstructPaleoenvironments: Compound Specific Hydrogen Isotopes and Pore-WaterOxygen Isotopes. Zurich: Swiss Federal Institute of Technology p. 149.

Pomati F, Burns B, and Neilan B (2004) Use of ion-channel modulating agents to studycyanobacterial Na(þ) – K(þ) fluxes. Biological Procedures Online 6: 137–143.

Pond KL, Huang YS, Wang Y, and Kulpa CF (2002) Hydrogen isotopic composition ofindividual n-alkanes as an intrinsic tracer for bioremediation and sourceidentification of petroleum contamination. Environmental Science and Technology36: 724–728.

Reed RH, Borowitzka LJ, Mackay MA, et al. (1986) Organic solute accumulation inosmotically stressed cyanobacteria. FEMS Microbiology Letters 39: 51–56.

Reed RH and Stewart WDP (1985) Osmotic adjustment and organic solute accumulationin unicellular cyanobacteria from freshwater and marine habitats. Marine Biology88: 1–9.

Roberts MF (2004) Osmoadaptation and osmoregulation in archaea: Update 2004.Frontiers in Bioscience 9: 1999–2019.

Roberts MF (2005) Organic compatible solutes of halotolerant and halophilicmicroorganisms. Saline Systems 1: 1–30.

Rock CO and Jackowski S (2002) Forty years of bacterial fatty acid synthesis.Biochemical and Biophysical Research Communications 292: 1155–1166.

Sachs JP and Schwab VF (2011) Hydrogen isotopes in dinosterol from the ChesapeakeBay estuary. Geochimica et Cosmochimica Acta 75: 444–459.

Sachse D, Radke J, and Gleixner G (2004) Hydrogen isotope ratios of recent lacustrinesedimentary n-alkanes record modern climate variability. Geochimica etCosmochimica Acta 68: 4877–4889.

Sachse D and Sachs JP (2008) Inverse relationship between D/H fractionation incyanobacterial lipids and salinity in Christmas Island saline ponds. Geochimicaet Cosmochimica Acta 72: 793–806.

Sato Y, Ito Y, Okada S, Murakami M, and Abe H (2003) Biosynthesis of thetriterpenoids, botryococcenes and tetramethylsqualene in the B race ofBotryococcus braunii via the non-mevalonate pathway. Tetrahedron Letters44: 7035–7037.

Sauer PE, Eglinton TI, Hayes JM, Schimmelmann A, and Sessions AL (2001) Compound-specific D/H ratios of lipid biomarkers from sediments as a proxy for environmentaland climatic conditions. Geochimica et Cosmochimica Acta 65: 213–222.

Schefusz E, Kuhlmann H, Mollenhauer G, Prange M, and Patzold J (2011) Forcing ofwet phases in southeast Africa over the past 17,000 years. Nature 480: 509–512.

Schefusz E, Schouten S, and Schneider RR (2005) Climatic controls on central Africanhydrology during the past 20,000 years. Nature 437: 1003–1006.

Schmidt H-L, Werner RA, and Eisenreich W (2003) Systematics of 2H patterns in naturalcompounds and its importance for the elucidation of biosynthetic pathways.Phytochemistry Reviews 2: 61–85.

Schouten S, Ossebaar J, Schreiber K, et al. (2006) The effect of temperature, salinity andgrowth rate on the stable hydrogen isotopic composition of long chain alkenonesproduced by Emiliania huxleyi and Gephyrocapsa oceanica. Biogeosciences3: 113–119.

Schouten S, Rijpstra WIC, Kok M, et al. (2001) Molecular organic tracers ofbiogeochemical processes in a saline meromictic lake (Ace Lake). Geochimica etCosmochimica Acta 65: 1629–1640.

Schwab VF and Sachs JP (2009) The measurement of D/H ratio in alkenones and theirisotopic heterogeneity. Organic Geochemistry 40: 111–118.

Schwab VF and Sachs JP (2011) Hydrogen isotopes in individual alkenones from theChesapeake Bay estuary. Geochimica et Cosmochimica Acta 75: 7552–7565.

Schwender J, Gemunden C, and Lichtenthaler HK (2001) Chlorophyta exclusively usethe 1-deoxyxylulose 5-phosphate/2- C -methylerythritol 4-phosphate pathway forthe biosynthesis of isoprenoids. Planta 212: 416–423.

Sessions AL (2001) Hydrogen Isotope Ratios of Individual Organic Compounds.Department of Geologiocal Sciences, Indiana University, p. 158.

Sessions AL (2006) Seasonal changes in D/H fractionation accompanying lipidbiosynthesis in Spartina alternflora. Geochimica et Cosmochimica Acta70: 2153–2162.

Sessions AL, Burgoyne TW, and Hayes JM (2001) Determination of the H-3 factor inhydrogen isotope ratio monitoring mass spectrometry. Analytical Chemistry73: 200–207.

Sessions AL, Burgoyne TW, Schimmelmann A, and Hayes JM (1999) Fractionation ofhydrogen isotopes in lipid biosynthesis. Organic Geochemistry 30: 1193–1200.

Sessions AL, Jahnke LL, Schimmelmann A, and Hayes JM (2002) Hydrogen isotopefractionation in lipids of the methane-oxidizing bacterium Methylococcuscapsulatus. Geochimica et Cosmochimica Acta 66: 3955–3969.

Steele NM and Fry SC (2000) Differences in catalytic properties between nativeisoenzymes of xyloglucan endotransglycosylase (XET). Phytochemistry54: 667–680.

Valentine DL, Chidthaisong A, Rice A, Reeburgh WS, and Tyler SC (2004) Carbonand hydrogen isotope fractionation by moderately thermophilic methanogens.Geochimica et Cosmochimica Acta 68: 1571–1590.

Ventosa A, Nieto JJ, and Oren A (1998) Biology of moderately halophilic aerobicbacteria. Microbiology and Molecular Biology Reviews 62: 504–544.

Volkman JK, Barrett SM, Blackburn SI, Mansour MP, Sikes EL, and Gelin F (1998)Microalgal biomarkers: A review of recent research developments. OrganicGeochemistry 29: 1163–1179.

Wakeham SG (1990) Algal and bacterial hydrocarbons in particulate matter andinterfacial sediments of the cariaco trench. Geochimica et Cosmochimica Acta54: 1325–1336.

Wolhowe MD, Prahl FG, Probert I, and Maldonado M (2009) Growth phase dependenthydrogen isotopic fractionation in alkenone-producing haptophytes.Biogeosciences 6: 1681–1694.




Yakir D (1992) Variations in the natural abundance of O-18 and deuterium in plantcarbohydrates. Plant, Cell and Environment 15: 1005–1020.

Yakir D and Deniro MJ (1990) Oxygen and hydrogen isotope fractionation duringcellulose metabolism in Lemna-Gibba L. Plant Physiology 93: 325–332.

Yang H, Pagani M, Briggs DEG, et al. (2009) Carbon and hydrogen isotope fractionationunder continuous light: Implications for paleoenvironmental interpretations of theHigh Arctic during Paleogene warming. Oecologia 160: 461–470.

Zhang Z and Sachs JP (2007) Hydrogen isotope fractionation in freshwater algae:I. Variations among lipids and species. Organic Geochemistry 38: 582–608.

Zhang Z, Sachs JP, and Marchetti A (2009) Hydrogen isotope fractionation in freshwaterand marine algae: II. Temperature and nitrogen limited growth rate effects. OrganicGeochemistry 40: 428–439.




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