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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.
Author's personal copy
12.4
Hydrogen Isotope Signatures in the Lipids of Phytoplankton
JP Sachs, University of Washington, Seattle, WA, USA
ã 2014 Elsevier Ltd. All rights reserved.
12.4.1
Introduction
12.4.2
The Effect of dDwater on dDlipid
12.4.3
The Effect of Biosynthesis on dDlipid
12.4.4
The Effect of Species on dDlipid
12.4.5
The Effect of Salinity on dDlipid
12.4.6
The Effect of Temperature on dDlipid
12.4.7
The Effect of Growth Rate on dDlipid
12.4.7.1
Substrate-Limited Growth Rate Effects
12.4.7.2
Light-Limited Growth Rate Effects
12.4.8
Summary and Conclusions
Acknowledgments
References
12.4.1
Introduction
Key Points
• Experimental evidence indicates that the following factors influence
algal lipid dD values. The magnitude of the effect based on published
studies 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 those
interested in applying algal lipid dD values in paleoclimate or paleoenvironmental studies:
• Use lipids unique to a family or species.
• Be mindful that changes in salinity will cause changes in lipid dD
values that will be additive to those associated with changes in
precipitation and evaporation.
• Expect lipid dD values to decrease if temperature or growth rate
increases.
The ability to rapidly measure deuterium-to-hydrogen
ratios in small quantities of individual lipids is less than
15 years old and has already led to significant advances in
Earth science. Pioneered by John Hayes and Alex Sessions in
the 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 to
contribute substantively to our understanding of lipid biosynthesis in bacteria (Sessions et al., 2002), algae (Zhang and
Sachs, 2007) and higher plants (Sessions, 2006), catabolism
in bacteria (Valentine et al., 2004), trophic relationships in
marine food webs (Chikaraishi, 2006), climate during the
Quaternary (Hu et al., 2003; Pahnke et al., 2007; Schefusz
et al., 2005, 2011) and the Eocene (Pagani et al., 2006), sediment transport in the ocean (Englebrecht and Sachs, 2005), the
source and fate of petroleum hydrocarbons (Pond et al., 2002),
forensic science, pharmacology, and archaeology (Benson
et al., 2006), to name a few. And despite a burgeoning interest
in applying lipid D/H ratios in these fields, there is much we do
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not understand about the environmental factors that influence
D/H fractionation in eukaryotes and prokaryotes and the biochemical and biophysical processes within cells that cause D/H
fractionation. This article focuses on the hydrogen isotope
signatures in lipids produced by phytoplankton because
those are the signatures most commonly used to evaluate
environmental conditions in the past (see Chapter 12.5).
Because all biosynthetic products in phytoplankton and
cyanobacteria appear to be depleted in deuterium relative to
environmental water, ferredoxin–NADP reductase (FNR) is a
likely source of D/H fractionation. This enzyme on the thylakoid membrane in photosystem I (PS1) catalyzes the reduction of NADPþ to NADPH. As such, it represents the first
opportunity newly liberated protons from the oxidation of
water in PS2 can experience mass-based fractionation.
NADPH thus produced is used to reduce CO2 in the Calvin
cycle, causing any fractionation by FNR to be reflected downstream in all biomolecules. Since hydrogen derived from
NADPH is likely to be depleted in deuterium by #250% or
more (Luo et al., 1991; Schmidt et al., 2003), it is expected
that lipids and other biosynthetic products will be depleted in
deuterium compared to environmental water. Any change in
growth conditions has the potential to alter D/H fractionation in algal cells by affecting the relative rates at which
PS2 (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 hydrogen
exchange with cellular H2O will diminish the initial
D-depletion (Hayes, 2001; Yakir, 1992; Yakir and Deniro,
1990). Changes in nutrients, light, temperature, or salinity
will alter demand for Hþ by PS1, H# by PGA, and the
exchange of hydrogen with cellular H2O, providing ample
opportunities for D/H fractionation.
Well established at this point is the fact that D/H ratios in
algal lipids are highly correlated with the D/H ratios of the
water in which the phytoplankton grew (Englebrecht and
Sachs, 2005; Sachs and Schwab, 2011; Sauer et al., 2001;
Schouten et al., 2006; Schwab and Sachs, 2011; Zhang and
Sachs, 2007; Figure 1). Also well established is the fact that all
lipids in autotrophic phytoplankton are depleted in deuterium
http://dx.doi.org/10.1016/B978-0-08-095975-7.01007-X
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Hydrogen Isotope Signatures in the Lipids of Phytoplankton
C27
200
B. braunii, A race
alkadienes
–100
–300
–100
300
0
–100
–200
y = 0.726x - 213.768
R2 = 0.999
100
E. huxleyi
alkenones
100
0
–200
(a)
C31
Alkadiene δD (‰)
Alkadiene δD (‰)
100
C29
y = 0.648x - 210.707
R2 = 1.000
–300
–100
500
100
(b)
Water δD (‰)
300
500
Water δD (‰)
Dinosterol δD (‰)
–270
–290
Chesapeake Bay
dinosterol
–310
–330
y = 1.87x -258
R2 = 0.95
–350
–50
(c)
–40
–30
–20
–10
0
Water δD (‰)
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 and
Sachs, 2007.
by approximately 100–400% relative to the water in which
they grew, owing to kinetic isotope effects (KIEs) during lipid
synthesis (Sessions et al., 1999). Furthermore, differing by
100% or more can be different lipids produced by the same
alga and the same lipid produced by different genera (Sessions
et al., 1999; Zhang and Sachs, 2007). The biosynthetic steps in
which the D/H fractionation occurs are just starting to be
worked out (Sessions, 2006; Sessions et al., 2002; Zhang and
Sachs, 2007) but remain largely unknown. Of equal or greater
importance for the application of hydrogen isotope techniques
in paleoceanography, geochemistry, and ecology are the environmental influences on the magnitude of that fractionation.
Here again, the influence on D/H ratios in algal lipids of the
most basic environmental parameters such as temperature,
salinity, light levels, and nutrient concentrations remains
largely unknown.
Though some progress has been made in characterizing and
understanding the hydrogen isotope signatures of lipids in
phytoplankton, there remains much more that we do not
know 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 isotopic
composition of environmental water and lipids, both within
a single species and between different species. Next, we discuss
the influence of water salinity on lipid D/H ratios, followed by
a discussion of the effects of temperature, nutrients, light levels,
and growth rates. We conclude with some thoughts on the
most pressing and promising questions to be addressed with
hydrogen isotopes in algal lipids.
12.4.2
The Effect of dDwater on dDlipid
Experiments with freshwater (Zhang and Sachs, 2007) and
marine (Englebrecht and Sachs, 2005) phytoplankton grown
in batch culture indicate that lipid dD values (dDlipid) are
highly correlated with water dD values (dDwater). Zhang and
Sachs (2007) grew five species of freshwater green algae in
batch culture, including Eudorina unicocca, Volvox aureus, and
three strains of Botryococcus braunii (two A race, one B race),
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Hydrogen Isotope Signatures in the Lipids of Phytoplankton
under controlled conditions in media containing five different
concentrations of deuterium. The hydrogen isotopic ratios of
lipids in the algae, including alkadienes, botryococcenes, heptadecenes, fatty acids, and phytadiene, were correlated with
water dD values with R2 values in excess of 0.99 for all lipids
in all species (Figure 2).
Similarly, Englebrecht and Sachs (2005) grew the marine
coccolithophorid Emiliania huxleyi in batch cultures with five
different deuterium enrichments (Figure 3). The hydrogen
isotope composition of C37 and C38 alkenones was linearly
correlated with dDwater with R2 values greater than 0.99.
A similar batch culture experiment with E. huxleyi performed
by Paul (2002) and reported in Schouten et al. (2006) produced 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 phytoplankton, representing both marine and freshwater realms,
and from both the acetogenic and the isoprenoid biosynthetic
pathways, have dDlipid values that are linearly correlated with
environmental water dD values, it is reasonable to expect that
this will be the case for all algal lipids under most circumstances. Field studies across the environmental gradient from
seawater to river water and across large biogeographic zones
corroborate this supposition.
Experiments performed in the Chesapeake Bay (CB) estuary
(Maryland and Virginia, USA) indicate that the dD values of four
different alkenones (both C37 and C38 di- and triunsaturated
varieties) were correlated with the dD value of water across a
salinity gradient of 19 with R2 values of 0.7–0.8 (Figure 4; Schwab
and Sachs, 2009, 2011). In addition, the dD values of the dinoflagellate sterol dinosterol (4a,23,24-trimethyl-5a-cholest-22Een-3b-ol) were correlated with the dD values of water across a
salinity gradient of 19 PSU with an R2 value of 0.9 (Figure 5; Sachs
and Schwab, 2011). The consistently high correlation between
dDwater and dDlipid in an estuarine setting in which the assemblage
of alkenone- and dinosterol-producing phytoplankton is likely to
300
C30bot
100
dD of botryococcenes (‰)
y = 0.751x - 277.948
(C30 bot)
C31bot
C32 + C33 bot
0
C34bot(4)
C34bot(5) + ?
-100
C34bot iso
y = 0.730x - 297.445
(C31 bot)
-200
-300
y = 0.726x - 329.067 (C34 bot [4])
dD of free fatty acids (corrected, ‰)
200
(Martinque)
0
100
200
300
400
C28:1
C20:1
y = 0.746x - 165.893
(C28:1)
-100
100
200
0
y = 0.807x - 167.963
(C28:1)
-100
y = 0.791x - 169.872
(C16)
100
200
300
400
500
dD of water at harvest (‰)
0
y = 0.557x - 282.096
-100
-200
-300
y = 0.715x - 162.069
(C16)
0
(c)
dD of phytadienes (‰)
dD of naturally occuring FAME (‰)
C18:1
-200
C20:1
100
100
C16
0
C28:1
0
y = 0.770x - 148.222
(C20:1)
100
C18:1
(b)
300
200
200
500
dD of water at harvest (‰)
(a)
y = 0.810x - 153.424
(C20:1)
C16
-200
y = 0.647x - 359.821 (C34 bot iso?)
-400
81
300
400
dD of water at harvest (‰)
500
0
(d)
100
200
300
400
500
dD of water at harvest (‰)
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 the
dD value of the water. Reproduced from Zhang Z and Sachs JP (2007) Hydrogen isotope fractionation in freshwater algae: I. Variations among lipids
and species. Organic Geochemistry 38: 582–608.
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Hydrogen Isotope Signatures in the Lipids of Phytoplankton
dDC37:2 alkenone (vs. VSMOW)
300
200
Englebrecht and Sachs (2005)
Paul (2002)
100
0
-100
dD37:2 = 0.724 ! dDH2O - 226
R2 = 0.991 (n = 13)
-200
-300
-10
90
190
290
390
490
590
dDH2O (vs. VSMOW)
Figure 3 dD values of alkenones versus dD values of growth water from batch culture experiments with the marine coccolithophorid E. huxleyi. Data
are from Englebrecht and Sachs (2005) and Paul (2002), as reported in Schouten et al. (2006).
-140
Alkenone dD
-160
-180
-200
c37:3 y = 0.84x - 156, R2 = 0.80
c37:2 y = 0.79x - 173, R2 = 0.72
-220
-240
-40
C38:3 y = 0.75x - 179, R2 = 0.75
C38:2 y = 0.98x - 183, R2 = 0.80
-30
-20
-10
Water dD (‰)
Figure 4 dD values of four C37 alkenones in suspended particles were highly correlated with dD values of water along a $175 km section of
the Chesapeake Bay estuary. Reproduced from Schwab VF and Sachs JP (2011) Hydrogen isotopes in individual alkenones from the Chesapeake
Bay estuary. Geochimica et Cosmochimica Acta 75: 7552–7565.
vary attests to the fact that isotopic changes of water are strongly
imprinted on lipids in phytoplankton.
Two other studies looked at the relationship between
dDwater and the dD value of lipids common to many algal
species in surface sediments from lakes across large biogeoclimatic zones in western Europe (Sachse et al., 2004) and the
eastern United States (Huang et al., 2002, 2004). In the
European lake survey, Sachse et al. (2004) found a high correlation between the dD value of the C17 normal alkane (nC17)
and the dD value of climatological average rainfall. In the study
of eastern US lakes, Huang et al. (2002) found a high correlation between the dD values of palmitic acid (nC16:0), nC17, and
phytol in surface sediments and lake water dD values across a
diversity of climates and limnologies (Figure 6). However,
these lipids are produced by many different plankton and
higher plants, so their sources in sediments are likely to be
mixed. A final example of the near-universality of the imprint
of water dD values on algal lipids comes from the fact that even
total lipid extracts (TLEs) from a wide diversity of freshwater
and saline lakes have dD values that are highly correlated with
dDwater (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 the
relationships depicted in Figures 1–6 can be attributed to the
biosynthetic pathway from which particular lipids are produced,
an observation first made by Estep and Hoering (1980)
(Estep and Hoering, 1980) and expanded upon by Sessions
et al. (1999). Lipids produced by the two isoprenoid pathways, 1-deoxyxylulose 5-phosphate/2-C-methyl-D-erythritol 4phosphate (DOXP/MEP) and mevalonic acid (MVA), have
branched carbon chains (e.g., sterols, phytol, botryococcenes,
and phytadienes) and are depleted in deuterium by approximately 100–200% relative to lipids synthesized by the acetogenic pathway which have a straight chain of carbon atoms
(e.g., fatty acids, alkenones, alkadienes, and the leaf wax
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Hydrogen Isotope Signatures in the Lipids of Phytoplankton
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with some desaturases. The same was inferred for some decarboxylase and methylase enzymes involved with lipid synthesis.
For example, decarboxylation of the C30:1 fatty acid to produce
the C29 alkadiene in B. braunii (A race) resulted in a D-depletion
of the product by as much as 40%, and the C31–C34 botryococcenes in cultured B. braunii (B race) were all depleted in
D relative to the C30 homologue from which they are synthesized (Zhang and Sachs, 2007). Elongation of carbon chains
may cause some D/H fractionation but is likely to be minor
compared to the previously discussed processes (Chikaraishi
et al., 2004; Zhang and Sachs, 2007).
y = 1.87x - 258
R2 = 0.95
Dinosterol dD (‰)
-290
-310
y = 1.35x - 284
R2 = 0.86
-330
-350
-45
Sediment
Particles
-35
-25
-15
Water dD (‰)
12.4.4
-5
Figure 5 dD values of the dinoflagellate sterol dinosterol were linearly
correlated with water dD values in the Chesapeake Bay estuary when
measured 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 Bay
estuary. Geochimica et Cosmochimica Acta 75: 7552–7565.
Palmitic acid dD (‰, VSMOW)
-100
-140
d DPA = -167.0 + 0.939 ! dDH2O
R = 0.894
-180
-220
-260
-300
-100
-80
83
-60
-40 -20
0
Water dD (‰, VSMOW)
20
40
Figure 6 dD value of palmitic acid from surface sediments of a transect
of lakes in the eastern United States (Huang et al., 2002).
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 pathway
contribute to the observed net D-depletion in lipids. These
include 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 step
in which hydrogen derived from NADPH is added will cause
isotopic depletion of the product (Hayes, 2001; Luo et al., 1991;
Schmidt et al., 2003). This was demonstrated by Chikaraishi
et al. (2009) by determining the dD values of phytol and its
precursors in cucumber cotyledons (Chikaraishi et al., 2009).
Desaturation of fatty acids and alkenones results in unsaturated
products that are substantially depleted in D relative to precursors (Chikaraishi et al., 2004; D’andrea et al., 2007; Schwab and
Sachs, 2009, 2011) (cf. Figure 4) implying a large KIE associated
The Effect of Species on dDlipid
It is the sum total of the aforementioned (and other) KIEs that
dictate the dD value of algal lipids. Because there are a series of
enzyme-mediated reactions leading to the synthesis of any
lipid, the physiological state of a cell will almost certainly
influence the extent to which each individual KIE is expressed
in the final product. The type and relative abundance of different biochemicals in different species of algal cells are also
expected to result in different isotopic depletions of a particular
lipid relative to environmental water. For example, under similar batch culture growth conditions, two species of Chlorophyceae (E. unicocca and V. aureus) and three species of
Trebouxiophyceae (B. braunii) produced palmitic acid (nC16:0
fatty acid) that differed by 160% relative to water (Zhang and
Sachs, 2007; Figure 10).
In addition to the magnitude of the KIE expressed in each
step of lipid synthesis differing between species, it is possible
that the enzymes used for a particular reaction can differ
between species, as is the case for fatty acid desaturase enzymes
in eukaryotic phytoplankton and cyanobacteria (Chi et al.,
2008). Ubiquitous lipids such as palmitic acid (C16:0), with a
multitude of aquatic and terrestrial sources, may therefore not
be 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 was
first observed by Schouten et al. (2006) in alkenones from
cultured coccolithophorids (E. huxleyi and G. oceanica) over
the salinity range 25–35 PSU. Our field data from Christmas
Island and the CB support that finding.
A decrease in D/H fractionation (increase in a) as salinities
increased occurred in several algal and cyanobacterial lipids in
ponds on Christmas Island with salinities of 17–149 PSU in June–
July 2005 (Sachse and Sachs, 2008; Figure 11). Fractionation
factors (a) for TLEs were between 0.797 and 0.908 and covaried
with salinity according to the relationship a ¼ 0.0007*S þ 0.79
(R2 ¼ 0.74, n ¼ 32), indicating a 0.7% decrease in D/H fractionation 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 the
nC17 alkane from algae (Sachse et al., 2004), diploptene from
bacteria (Elvert et al., 2001; Wakeham, 1990), and phytene
from algae and/or bacteria (Boudou et al., 1986; Hefter et al.,
1993; Schouten et al., 2001) were similar to one another
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Hydrogen Isotope Signatures in the Lipids of Phytoplankton
-180.00
-190.00
d D TLE
-200.00
-210.00
-220.00
-230.00
y = 0.9389x - 151.44
R2 = 0.8189
-240.00
-250.00
-100.00
-80.00
-60.00
-40.00
d D Water
dD (‰)
Methylococcus
Daucus
Spartina
Zostera
Fucus
Ascophyllum
-100
Isochrysis
0
Alexandrium
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 of
the lake water. Salinities of all lakes were less than 60 ppt (Nelson and Sachs, unpublished).
-200
-300
-400
n-alkyl C skeletons
n-alkanes
n-alcohols
Fatty acids
Isoprenoid C skeletons
Sterols
Sesqui- and triterpenoids
Phytol and phytene
Figure 8 Acetogenic lipids (open symbols) are usually enriched in
D relative to isoprenoid lipids in a variety of plants, algae, and bacteria.
Reproduced from Sessions AL, Burgoyne TW, Schimmelmann A, and
Hayes JM (1999) Fractionation of hydrogen isotopes in lipid
biosynthesis. Organic Geochemistry 30: 1193–1200.
(0.00076–0.0011) and to the subset of TLEs from which the biomarkers were purified, 0.00083 a units per PSU salinity
(Figure 11). The higher y-intercept (or a at zero salinity) for
nC17 relative to phytene and diploptene in Figure 11 is consistent
with the widely noted D-depletion in isoprenoid lipids relative to
acetogenic lipids (Hayes, 2001; Sessions et al., 1999; Zhang and
Sachs, 2007). A y-intercept for the TLEs that is intermediate
between acetogenic nC17 and isoprenoid phytene and diploptene
is 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 was
0.00099 in suspended particles and significant at the 95% confidence level (Figure 12). This translates into a decrease in D/H
fractionation of 0.99% per PSU increase in salinity. This sensitivity of D/H fractionation to salinity is similar to those for
phytene (1.1% per PSU), nC17 alkane (0.80% per PSU), and
TLEs (0.70% per PSU) from Christmas Island. Regardless of the
biosynthetic pathway, environment, or source of the lipid, the
D/H fractionation associated with salinity appears to be relatively constant in the diverse settings of the CB estuary and the
hypersaline ponds on Christmas Island.
The higher slope of the linear regression of adino/water onto
salinity (Figure 4(b)) in sedimentary dinosterol relative to
particulate dinosterol can be explained by high dinoflagellate
production during summer when river flows and runoff are
lowest. The particulate dinosterol and water samples were
taken during high flow conditions in springtime when surface
water dD values and salinities in the CB are expected to be
relatively low compared to the summer. In other words, the
water overlying sediments in May is likely to be characterized
by lower dDwater and salinity values than would typically occur
in summer when dinoflagellate production peaks. Annual fluxweighted dDdino values are thus expected to be higher than
dDdino values measured in May in response to the combined
influence of higher dDwater values and diminished D/H fractionation in higher salinity water during summer.
Unlike dinosterol, D/H fractionation in alkenones from
suspended particles in the CB was unchanged as salinity
increased from 10 to 29 PSU (Schwab and Sachs, 2011; Figure 13). We have no satisfactory explanation for the different
behavior of alkenones relative to alkenones in cultured coccolithophorids, dinosterol in the CB, or a variety of lipids in
saline ponds on Christmas Island. One possibility is that a
change in the species of alkenone producers occurs along the
length of the estuary such that a change in the magnitude of
D/H fractionation imparted to alkenones by the different
species cancels out the effect of salinity on D/H fractionation.
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Hydrogen Isotope Signatures in the Lipids of Phytoplankton
85
300
• 170‰ difference
in D/H fractionation
for different lipids in
B. braunii (B race)
Acetogenic
200
Lipid dD (‰)
100
0
Palmitic acid
170‰
-100
C16:0 FAME
-200
C34 botryococcene
Isoprenoid
C30 botryococcene
-300
Phytadiene
-400
0
100
200
300
400
500
Water dD (‰)
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 freshwater
algae: I. Variations among lipids and species. Organic Geochemistry 38: 582–608.
dD of C16 fatty acids (corrected, ‰)
400
160‰ difference
in palmitic acid
between families of
green algae
300
200
100
160‰
0
V. Aureus (15 "C)
E. unicocca
V. Aureus (25 "C)
B. Braunii (A race)
B. Braunii (B race)
-100
-200
0
100
200
300
400
dD of water at harvest (‰)
500
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 Z
and Sachs JP (2007) Hydrogen isotope fractionation in freshwater algae: I. Variations among lipids and species. Organic Geochemistry 38: 582–608.
The magnitude of the species effect in batch cultures of two
coccolithophorids (E. huxleyi and Gephyrocapsa oceanica) was
about 30% in Schouten et al. (2006), which, coincidentally, is
just the right magnitude to counter a 1% decrease in D/H
fractionation per unit increase in salinity in the CB if the
assemblage of alkenone producers was dominated by E. huxleyi
at low salinity, and G. oceanica at high salinity. Another possible explanation for the lack of change in a for alkenones in
the CB estuary is that greater osmoregulation capacity in
coastal haptophytes may result in a diminished sensitivity of
alkenone–water D/H fractionation to salinity changes.
A linear decrease in D/H fractionation as salinity increases is
clear from the Christmas Island saline pond lipids, the CB
dinosterol, and Schouten et al. (2006)’s coccolithophorid culture experiments. Unexplained is why Schouten observed a
threefold greater sensitivity of D/H fractionation to changes in
salinity in alkenones from batch culture (3.3% per PSU) than
either of our field studies indicate for a wide diversity of lipids
(0.7–1.1% per PSU). The most likely explanation for this discrepancy is that different growth rates for the coccolithophorids
at different salinities in the batch cultures imparted additional
D/H fractionation in the alkenones. This effect is discussed in
detail later in Section 12.4.7.
The mechanism or mechanisms by which increasing
salinity causes decreasing D/H fractionation between water
and lipids in algae and cyanobacteria remains unknown.
Controlled experiments with continuous cultures are needed
so that all growth and environmental conditions can be held
constant while salinity is varied. Such experiments are underway in our laboratory at the University of Washington, and
presumably in other laboratories. In the meantime, we offered
three hypotheses to explain why increased salinity resulted in
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Hydrogen Isotope Signatures in the Lipids of Phytoplankton
0.95
Xmas nC17
Xmas nC17:1
Xmas nC18
Xmas nC18:1
Xmas phytene
Xmas diploptene
Xmas TLE
CB dinosterol
0.9
a lipid–water
0.85
y = 0.81 + 0.0008x R2 = 0.79
y = 0.76 + 0.0012x R2 = 0.95
0.8
y = 0.77 + 0.00074x R2 = 0.48
y = 0.81 + 0.00069x R2 = 0.85
0.75
y=0.65 + 0.0011x R2 = 0.91
y = 0.72 + 0.00082x R2 = 1
0.7
y = 0.78 + 0.00076x R2 = 0.66
0.65
y = 0.68 + 0.00099x R2 = 0.57
0
50
100
150
Salinity
Figure 11 alipid–water versus salinity relationships for total lipid extracts and lipid biomarkers in Christmas Island saline ponds (open symbols) and
dinosterol from the Chesapeake Bay estuary (orange squares). Dinosterol is an isoprenoid lipid from dinoflagellate algae, and nC17 (circles) and nC17:1
are acetogenic lipids from algae and cyanobacteria. Diploptene (triangles) is an isoprenoid from bacteria, and phytene (diamonds) is an isoprenoid lipid
from bacteria and algae. Data from Sachse and Sachs (2008) and Sachs and Schwab (2011).
0.740
water pool from which lipids are synthesized (Kreuzer-Martin
et al., 2006; Sachse and Sachs, 2008).
Sediment
a = 0.00174 ± 0.000321 *
salinity + 0.685 ± 0.005, R2
= 0.77
0.730
a
0.720
0.710
0.700
Suspended particles
a = 0.000990 ± 0.000229 *
salinity + 0.685 ± 0.004,
R2 = 0.57
0.690
0.680
5
10
15
20
Salinity
25
30
35
Figure 12 adinosterol/water values in suspended particles (filled circles,
N ¼ 16) and surface sediment (open circles, N ¼ 11) plotted against
salinity. Error bars for adino/water represent propagated errors reported for
dDdino and dDwater measurements. Linear regressions for suspended
particles (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 Bay
estuary. Geochimica et Cosmochimica Acta 75: 7552–7565.
decreased D/H fractionation between algal lipids and extracellular water in Sachs and Schwab (2011) and Sachse and Sachs
(2008). They are as follows:
Hypothesis 1 Higher salinities cause decreased water transport across the cell membrane, resulting in greater recycling
of internal water, and an increasingly D-enriched internal
Water transport across the cell membrane via aquaporins
and Naþ channels is restricted upon exposure to high salinities
to prevent Naþ toxicity (Boursiac et al., 2005; Pomati et al.,
2004), while at the same time, the volume of water in cyanobacterial cells has been shown to decline by up to 25% during
salt stress (Allakhverdiev et al., 2000a,b). Lower transport rates
of water across the cell wall and a decreased pool of internal
water would act in concert to continuously enrich the dD values
of cellular water as NADPH is continuously replenished with
D-depleted hydrogen. Estimates of the dD values of hydrogen
for NADPþ protonation range from #250 to #600% (Luo
et al., 1991; Schmidt et al., 2003). Lipids (and all other biosynthetic products) would reflect the increasingly elevated dD
values characterizing the internal water. Consequently, it is possible that D/H fractionation between dinosterol and CB water
decreases from the headwaters to the mouth of the bay as a
result of decreased water transport across dinoflagellate cell
membranes, diminution of their internal water pool, and an
increase in the dD value of internal water as water is recycled
rather than replenished from outside the cell, and D-depleted
hydrogen is withdrawn for NADPH production.
Hypothesis 2 Elevated salinities cause growth rates to decline,
resulting in diminished D/H fractionation between lipids and
extracellular water.
Many studies on phytoplankton and algae indicate that
growth rates generally decrease upon exposure to elevated salinities (Cifuentes et al., 2001; Clavero et al., 2000; Herbst and
Bradley, 1989). Though different algae are adapted to a range of
optimal salinities, virtually all experience a decline in growth
rate once a threshold salinity is passed. Laboratory culture
studies on the marine diatom Thalassiosira pseudonana (Zhang
et al., 2009) and the marine coccolithophorids E. huxleyi and
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Hydrogen Isotope Signatures in the Lipids of Phytoplankton
87
0.860
0.850
0.840
0.830
0.820
0.810
0.800
8
10
12
14
16
18
20
22
24
26
28
30
Salinity
MeC37:3 in sediments
and filters
MeC38:3 in sediments
and filters
MeC37:2 in sediments
and filters
MeC38:2 in sediments
and filters
Figure 13 Hydrogen isotope fractionation between four alkenones and environmental water (aalkenone/water) as a function of measured surface
water salinity in the Chesapeake Bay estuary, May 2006. Suspended particle values are in black. Surface sediment values are in gold. No influence
of salinity on D/H fractionation in alkenones is observed in this data. Reproduced from Schwab VF and Sachs JP (2011) Hydrogen isotopes in
individual alkenones from the Chesapeake Bay estuary. Geochimica et Cosmochimica Acta 75: 7552–7565.
G. oceanica (Schouten et al., 2006) indicate substantial decreases
in D/H fractionation expressed in a variety of lipids as growth
rates are reduced, either through purposeful nitrogen limitation
(Zhang et al., 2009) or some combination of salinity and temperature (Schouten et al., 2006). If growth rates of dinosterolproducing dinoflagellates declined along the length of the CB, it
is possible that D/H fractionation between dinosterol and CB
water decreased in concert.
Hypothesis 3 Upon exposure to high salinities, rapid production of osmolytes draws D-depleted hydrogen from an internal
pool of water, leaving it, and subsequent lipids synthesized
from it, enriched in deuterium.
When subjected to osmotic stress, cells from all domains of
life accumulate organic solutes to counteract the external
osmotic pressure (Borowitzka and Brown, 1974; Borowitzka
et al., 1977; Brown, 1978; Galinski, 1995; Grant, 2004; Mackay
et al., 1984; Reed and Stewart, 1985; Reed et al., 1986; Roberts,
2004, 2005; Ventosa et al., 1998). Also called osmolytes or
compatible solutes (because they provide osmotic balance without interfering with the metabolic functions of the cell; Ventosa
et al., 1998), these small organic molecules span a wide diversity
of structures and can be zwitterionic, uncharged, or anionic
(Roberts, 2005). Furthermore, they can represent a very substantial fraction of cellular hydrogen, reaching up to 10–20% of the
dry weight of cells in some hypersaline bacteria (Ventosa et al.,
1998). If hydrogen is shuttled off from internal pools of water
and NADPH to rapidly synthesize osmolytes in response to
osmotic stress, residual pools of those substances would be left
enriched in deuterium, and all subsequent biosynthetic products
would reflect this enrichment. Thus, if dinoflagellates produce
increasingly greater concentrations of osmolytes as salinities
increase seaward in the CB, their internal pools of water and
NADPH might become increasingly D-enriched, as would the
dinosterol synthesized from that water, resulting in less D/H
fractionation relative to CB water.
Both hypotheses 1 and 3 call on a change in the internal
water dD value as salinity changes and would thus predict that
all biosynthetic products would experience a decrease in D/H
fractionation relative to external water since water is the source
of hydrogen for all biochemicals in phytoplankton. Hypothesis 2 calls upon a salinity-induced reduction in growth rate as
the source of lowered D/H fractionation in dinosterol (and
other lipids) relative to water, but does not necessarily predict
that all biosynthetic products be increased in dD. These are
testable hypotheses that future work ought to address.
12.4.6
The Effect of Temperature on dDlipid
Hydrogen isotope fractionation in algal lipids appears to
increase as temperature increases. Zhang et al. (2009) showed
that acetogenic lipids from two species of freshwater green
algae (E. unicocca and V. aureus) grown in batch culture at
15 & C were enriched in deuterium by 20–40% relative to
those grown at 25 & C (Zhang et al., 2009; Figure 14). The lipids
included palmitic acid (C16:0) in both species (Figure 14(a)),
the naturally occurring methyl esters of palmitic acid in both
species, stearic acid (C18:0) and its naturally occurring methyl
ester in E. unicocca, and heptadecene (C17:1ene) in E. unicocca
(Figure 14(b)). Yet with only two temperatures investigated, it
would be imprudent to generalize these results.
Further support for an increase in D/H fractionation in
algal lipids as temperature increases comes from two independent laboratory culture studies with marine coccolithophorids
(Figure 15). Schouten et al. (2006) cultured E. huxleyi in batch
cultures at three temperatures and three salinities and found an
increase in D/H fractionation (decrease in alpha) in alkenones
of about 0.4–1% per degree Celsius increase in temperature
(Figure 15). The sensitivity of D/H fractionation to temperature
appears to have been even larger in alkenones from G. oceanica,
in which dD values decreased by 3–4% per degree Celsius
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Hydrogen Isotope Signatures in the Lipids of Phytoplankton
Eudorina
EU-C17:1ene
VA-C16FAME
0.97
y = -0.0039x + 1.0138
0.94
0.91
0.88
EU-C16FAME
y = -0.0022x + 0.9798
y = -0.0038x + 0.9947
y = -0.0037x + 0.9885
y = -0.0035x + 0.9792
0.94
aLipid–water
aPalmitic acid–water
0.97
EU-C18FA
EU-C18FAME
Volvox
0.91
0.88
y = -0.0033x + 0.9887
y = -0.0039x + 0.9608
0.85
10
15
20
25
Temperature ("C)
(a)
0.85
10
30
15
20
25
Temperature ("C)
(b)
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 (closed
symbols) and V. aureus (open symbols). (a) C16:0 (palmitic) acid in both species. (b) nC17:1 alkene in Eudorina (squares) and several fatty acids and
natural 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 rate
effects. Organic Geochemistry 40: 428–439.
a alkenone–water
0.82
E. huxleyi
0.81
Eh S = 25
y = −0.0012x + 0.79
R 2 = 0.19
0.8
Eh S = 29
y = −0.0008x + 0.80
R 2 = 0.94
Eh S = 35
y = −0.0004x + 0.82
R 2 = 0.43
Go S = 25
y = −0.0028x + 0.80
Go S = 29
y = −0.0043x + 0.84
0.75
Go S = 35
y = −0.0028x + 0.83
0.74
Wol Go
0.79
0.78
0.77
0.76
0.73
y = −0.0049x + 0.89
R 2 = 0.97
G. oceanica
7
12
17
22
27
Temperature (°C)
Figure 15 D/H fractionation in C37 alkenones from the cultured marine coccolithophorids Emiliania huxleyi and Gephyrocapsa oceanica from the
studies of Schouten et al. (2006) (colored symbols) and Wolhowe et al. (2009) (black symbols). Closed symbols are from E. huxleyi. Open symbols are
from G. oceanica. Schouten et al. (2006) performed experiments at three salinities (S ¼ 25, 29, and 35). Taken together, the data imply a decrease in
alkenone dD values of 0.4–5% per degree Celsius increase in water temperature.
increase in temperature (Figure 15). This is the same temperature sensitivity we observed in lipids from E. unicocca and
V. aureus cultures (see Figure 14). Although Schouten et al.
(2006)’s G. oceanica alkenone data are from just two temperatures, they are compelling because three cultures were grown at
each of those temperatures, one at each of three salinities, and
because an independent set of three G. oceanica batch cultures by
Wolhowe et al. (2009) yields a similarly high temperature
sensitivity. D/H fractionation in alkenones increased by 5%
(i.e., alpha decreased by 0.0049) per degree Celsius in that
study (Figure 15) (Wolhowe et al., 2009).
Contradicting these two studies are the results from the CB
where no change was observed in apparent D/H fractionation in
four different alkenones over the surface water temperature range
of 15& –20 & C (Schwab and Sachs, 2011; Figure 16). But as discussed in the section on salinity, where it was shown that no
change in D/H fractionation between alkenones and water was
evident in the CB (Figure 13), other factors may be affecting the
hydrogen isotope trends in alkenones along the Bay, such as a
change in the assemblage of alkenone producers along the salinity gradient.
Zhang et al. (2009) discuss three mechanisms by which
temperature may influence D/H fractionation during lipid synthesis: its influence on enzyme activities, KIEs, and hydrogen
tunneling. Lipid biosynthesis is catalyzed by a variety of enzymes
whose conformation facilitates reaction with substrates. When
the enzyme is in the native (active) state, the reaction rate
increases with temperature. Temperature changes may induce
the (partial) replacement of an enzyme by an isoenzyme with
better 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 factors (5% difference) in methanotrophic bacteria using the
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Alkenone–water D/H fractionation
Hydrogen Isotope Signatures in the Lipids of Phytoplankton
89
0.860
0.850
0.840
0.830
0.820
0.810
0.800
15
16
17
18
Temperature ("C)
19
20
MeC37:3 in sediments
and filters
MeC38:3 in sediments
and filters
MeC37:2 in sediments
and filters
MeC38:2 in sediments
and filters
Figure 16 Hydrogen isotope fractionation between four alkenones and environmental water (aalkenone/water) as a function of measured surface water
temperature in the Chesapeake Bay estuary, May 2006. Suspended particle values are in black. Surface sediment values are in gold. No influence of
temperature on D/H fractionation in alkenones is observed in these data. Reproduced from Schwab VF and Sachs JP (2011) Hydrogen isotopes in
individual alkenones from the Chesapeake Bay estuary. Geochimica et Cosmochimica Acta 75: 7552–7565.
ribulose monophosphate pathway for carbon assimilation.
However, after passing some optimum temperature that is specific to each enzyme, denaturalization can occur, making the
enzyme lose its effectiveness as a catalyst. In this way, the set of
enzymes involved in the synthesis of a particular lipid, their
sensitivity to temperature, and any D/H fractionation they
impart will influence the dD value of the lipid.
Isotope fractionation in biochemical processes arises from
unequal zero-point energies of bonds between heavy and light
isotopes, resulting in different activation energies (Bigeleisen
and Wolfsberg, 1959), which in turn are influenced by temperature. Fractionation factors are thus a function of both the
temperature and the activation energy of the individual
enzyme-catalyzed reactions that comprise the lipid biosynthetic pathway.
Yet the greater D/H fractionation we observed at higher temperature is at odds with theory that indicates that equilibrium
fractionation should decrease as temperature increases (Clark
and Fritz, 1997). In any complex organism, changing growth
temperature could lead to a whole suite of metabolic changes,
any one of which could result in the observed isotopic differences. For example, different enzymes (isoenzymes or isozymes)
may be used to synthesize lipids at different temperatures, each
with its own fractionation (Jahnke et al., 1999). In one study by
Steele and Fry (2000), two isoenzymes of xyloglucan endotransglycosylase (XET) isolated from Arabidopsis (i.e., cauliflower
florets) were found to be temperature-dependent, one with an
optimum temperature of $12 & C and exhibiting 55% of its
maximal catalytic rate at #5 & C, total XET activity of mixed isoenzymes with a temperature optimum of $ 30 & C (Steele and
Fry, 2000). Assuming those two enzymes exhibit different isotope effects, the temperature at which the organism exists would
cause different D/H ratios in the lipid product.
Other possible causes for a temperature influence on D/H
fractionation in lipid synthesis include different mechanisms
for synthesizing NADPH at different temperatures, such as
photosynthesis versus the pentose phosphate pathway
(Kruger and von Schaewen, 2003), and hydrogen tunneling
due to strengthened substrate–enzyme complex vibration at
elevated temperatures (Kohen et al., 1999).
12.4.7
12.4.7.1
The Effect of Growth Rate on dDlipid
Substrate-Limited Growth Rate Effects
The only published study of which I am aware that has controlled growth rate in order to investigate its influence on D/H
fractionation in lipids is Zhang et al. (2009). Other studies
bearing on this question have inferred how different growth
rates, brought about by either temperature or salinity variations in batch cultures in the case of Schouten et al. (2006), or
different 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 temperature and salinity experiments and found an increase in
D/H fractionation in alkenones as growth rate increased.
Wolhowe et al. (2009) looked at the influence of growth
stage in batch cultures of E. huxleyi and observed 20–30%
greater D/H fractionation in alkenones from cells in the stationary phase relative to the exponential phase of growth
(Wolhowe et al., 2009). But these results are not readily comparable to growth rate experiments, and it is unclear how
applicable they are to the environment. We conducted two
continuous culture experiments, or chemostats, with the
marine diatom T. pseudonana, at constant growth rates controlled by N limitation. Our results indicate that there is a
substantial increase in D/H fractionation with increasing
growth rate in a sterol, an isoprenoid (e.g., branched) lipid
synthesized via the MVA pathway, and a smaller or negligible
increase in D/H fractionation in fatty acids synthesized via the
acetogenic pathway.
Two cultures of T. pseudonana were cultivated at growth
rates of 0.4 and 1.9 day#1. With only two data points, we
estimate that D/H fractionation increased by 25% per day as
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Hydrogen Isotope Signatures in the Lipids of Phytoplankton
C16:1
C14:0
C16:0
C18:0
Sterol
0.89
a lipid–water
0.84
0.79
0.74
0.69
y = -0.025x + 0.715
0.64
0
0.5
1
1.5
Growth rate (day –1)
2
Figure 17 a vs. growth rate in T. pseudonana. D/H fractionation
increased (a decreased) as specific growth rate increased in the sterol
24-methyl-cholesta-5,24(28)-dien-3b-ol (circles). But a remained
constant in fatty acids at two growth rates. This suggests that D/H
fractionation during lipid synthesis via the MVA biosynthetic pathway is
sensitive to changes in growth rate but not via the acetogenic pathway.
1s error bars are smaller than the symbol in most cases. Adapted 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.
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 synthesized
from the acetogenic pathway from cultures grown at 0.4 and
1.9 day#1 despite the nearly fivefold increase in growth rate
(Zhang et al., 2009).
In order for fatty acid dD values to remain constant at
different growth rates, the isotopic composition of the hydrogen source(s) for fatty acid synthesis (1) must either be unaffected 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 added
in some steps and D-enriched hydrogen is added in others. As
reviewed in Zhang and Sachs (2007), fatty acid synthesis can
be divided into three steps: the activation of acetyl-CoA to
malonyl-CoA, chain initiation from a unit of acetyl-CoA plus
a unit of malonyl-CoA and subsequent elongation with
malonyl-CoA by a fatty acid synthase (FAS) complex, and
desaturation by desaturase enzyme (Duan et al., 2002).
At the odd-numbered carbon positions, carbon comes from
the carboxyl group (#C¼O–) of acetate, and hydrogen is
derived entirely from NADPH. At the even-numbered carbon
positions, one hydrogen atom is derived from acetate and the
other from water during the enoyl-ACP reductase step (Duan
et al., 2002; Schmidt et al., 2003). Presently, the precise mechanism of enzyme-mediated exchange of carbon-bound hydrogen with intracellular water is not known. If substantial
hydrogen exchange occurs between fatty acids and their intermediates with a large pool of intracellular water, it could
explain 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 log
phase of growth than in the stationary phase of growth in the
bacterium Escherichia coli and attributed the greater isotopic
depletion to a larger contribution of hydrogen from intracellular water during log-phase growth. As a heterotroph, E. coli
maintains metabolic water that is more D-depleted than its
extracellular water (Kreuzer-Martin et al., 2006). As a photoautotroph, T. pseudonana maintains intracellular (or metabolic) water enriched in deuterium because isotopically
depleted hydrogen is continuously removed for NADPH production (Schmidt et al., 2003), leaving D-enriched metabolic
water. Furthermore, there is no evidence to suggest that the
enzymes involved in fatty acid synthesis differ between log and
stationary growth phases (Heath and Rock, 1996; Rock and
Jackowski, 2002). If Kreuzer-Martin et al. (2006)’s hypothesis
applies, fatty acids in T. pseudonana nitrate-replete (NR) cells
ought to be less D-depleted than those in nitrate-limited (NL)
cells, a prediction not supported by our data. Thus, either
hydrogen exchange occurs between intracellular water and
fatty acids or their intermediates, and/or intracellular water
dD values are insensitive to growth rates.
D/H fractionation in sterols from T. pseudonana varied
widely with nitrogen nutritional status. e values for
24-methyl-cholesta-5,24(28)-dien-3b-ol were 37% lower in
rapidly growing NR cells than in slow-growing NL cells
(Zhang et al., 2009). Sterols, like other isoprenoid lipids, originate from a branched isoprene C5 unit, isopentenyl diphosphate (IPP), or its isomer, dimethylallyl diphosphate
(DMAPP). The ‘eukaryotic’ acetate–MVA pathway starts from
three acetyl-CoAs and requires six enzymes, two NADPHs, and
three ATPs to produce IPP, which is subsequently converted to
DMAPP by IPP isomerase. In contrast, the non-MVA, ‘prokaryotic’ DOXP/MEP pathway of IPP synthesis begins with pyruvate and glycerinaldehyde-3-phosphate and involves seven
enzymes, three ATP equivalents, and three NADPHs to produce both IPP and DMAPP (Lichtenthaler, 2004).
In higher plants and many algae, such as the Bacillariophyta
to which T. pseudonana belongs, the two IPP-producing
biochemical pathways operate simultaneously, with the MVA
pathway restricted to the cytoplasm and the DOXP/MEP pathway restricted to the chloroplast (Armbrust, 2004; Bick and
Lange, 2003; Hemmerlin et al., 2003; Lichtenthaler, 1999;
Schwender et al., 2001; Figure 18). Although sterol synthesis
in most plants and algae occurs via the MVA pathway, some
algae, including certain chlorophytes, can use the DOXP/MEP
pathway to synthesize sterols and other isoprenoid lipids such
as 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 cytosolic MVA pathway and the plastidic DOXP/MEP pathway.
Laule et al. (2003) provided indirect support for the presence
of such an export mechanism in the vascular plant Arabidopsis
thaliana (Laule et al., 2003). Hemmerlin et al. (2003) reported
that sterols could be synthesized via the DOXP/MEP pathway
when the MVA pathway in Bright Yellow-2 cells was inhibited
and that isoprenoids normally produced in the plastid by the
DOXP/MEP pathway could be produced in the cytosol via the
MVA pathway when the former was inhibited. This suggested
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Hydrogen Isotope Signatures in the Lipids of Phytoplankton
91
N-limited (NL)/N-replete (NR)
Pyruvate + glyceraldehyde-3-phosphate
Acetyl-CoA
Acetyl-CoA
-CO2
1-Deoxy-D-xylulose 5-phosphate (DOXP)
Fatty acid
biosynthesis
HMG-CoA
NADPH
+
NADP
D-depleting step
2-C-methyl-D-erythritol 4-phosphate (MEP)
NAD(P)H
Mevalonate-5-phosphate
+
NAD
D-depleting step
Relatively more D-deplete
C16 fatty acid
IPP
DMAPP
Cross talk
NL
Yield: 0.30 pg cell-1
D/H: e = -143.6‰
NR
Yield: 0.07 pg cell-1
D/H: e = -142.0‰
Acetogenic pathway
Mevalonic acid
Mevalonate-5-pyrophosphate
Relatively less -CO2
D-deplete
IPP
DMAPP
Now more D-depleted
b-carotene
Lanosterol
37‰ more negative
Zeaxanthin, etc.
24-Methyl-cholesta-5,24(28)-dien-3b-ol
NL (w/o cross talk)
Yield: 0.059 pg cell-1
D/H: e = -294.7‰
NR (w/ cross talk)
Yield: 0.074 pg cell-1
D/H: e = -332.0‰
Isoprenoid biosynthesis (I)
DOXP/MEP
Plastid
Isoprenoid biosynthesis (II)
MVA pathway
Cytosol
Figure 18 Schematic diagram showing the cross talk of IPPs between the plastidic DOXP/MEP pathway and the cytosolic MVA pathway in the marine
diatom 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). Fatty
acid 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 cell
membranes. Under faster growing conditions, the IPPs from the plastidic DOXP/MEP pathway that produces highly D-depleted products might
cross into the cytosol, providing additional IPPs to the MVA-pathway products that are less D-depleted. As a result, sterols synthesized in the NR
culture had dD values that were 37% lower (#332% vs. #295% in the NL culture). Acetogenic compounds such as fatty acids are synthesized
exclusively 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 NL
culture. 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.
that significant exchange of isoprenoid intermediates occurred
across the plastid envelope (Hemmerlin et al., 2003). Furthermore, Bick and Lange (2003) proposed that plastid membranes possess a unidirectional proton/IPP coupled transport
system for the export of IPP from the plastid to the cytosol
(Bick and Lange, 2003).
We thus hypothesize that in fast-growing (NR) cells, IPP
and DMAPP monomers from both the MVA and DOXP/MEP
pathways are mixed to produce sterols (Figure 18). For
instance, if the supply of IPP from the cytosolic MVA route is
insufficient for sterol synthesis, additional IPP may come from
the DOXP/MEP pathway. The metabolic state of the cells could
dictate the relative contribution from the two pathways.
Because the last reduction step during IPP synthesis via the
DOXP/MEP pathway is characterized by a particularly large
hydrogen isotope effect (Zhang and Sachs, 2007), and thus
IPPs produced via that pathway are substantially D-depleted
relative to IPPs produced via the MVA pathway, a higher proportion of IPP derived from the DOXP/MEP pathway would
result in the production of more deuterium-depleted sterols
(Figure 18). This could explain the 37% depletion in deuterium in the fast-growing NR T. pseudonana cells relative to the
slow-growing NL cells.
Consistent with our findings, Sessions et al. (1999) reported
that sterols from dormant plants, which may be analogous to
our slow-growing NL T. pseudonana cells, were deuteriumenriched by 50–100% relative to actively growing plants,
whereas fatty acids were substantially less enriched in deuterium, by 0–30%. Cells in actively growing plants may incorporate some IPPs into sterols from the DOXP/MEP pathway
resulting in more deuterium depletion than in sterols synthesized in dormant plants if the MVA pathway alone produces
sterols. As we found, D/H fractionation in fatty acids observed
by Sessions et al. (1999) was not greatly affected by growth rates.
Sessions (2006) attributed the larger D-depletion in lipids
from actively growing plants to their faster metabolism. This
might explain why sterols in faster growing cells are more
depleted in deuterium but does not explain why fatty acid
D/H ratios were minimally affected by growth rate in our
experiments. Because isoprenoid lipids alone expressed
increased D/H fractionation at higher growth rates, it suggests
that the exchange of isoprenoid precursors (IPPs) between the
cytosol and the plastid occurs in rapidly growing cells, but not
the exchange of acetogenic precursors.
Verification of these findings with additional T. pseudonana
chemostats and with cultures of other phytoplankton are required
before firm conclusions can be made regarding the influence of
growth rate on D/H fractionation in different lipids from the same
cell. But taken together, the batch culture experiments of
Schouten et al. (2006) and Wolhowe et al. (2009) and the
Treatise on Geochemistry, Second Edition, (2014), vol. 12, pp. 79-94
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92
Hydrogen Isotope Signatures in the Lipids of Phytoplankton
chemostat culture experiments of Zhang et al. (2009) support the
notion that hydrogen isotope fractionation in some algal lipids
increases with increasing growth rate, but perhaps not in all lipids.
12.4.7.2
Light-Limited Growth Rate Effects
Light levels also influence growth rates of phytoplankton, but
to our knowledge, there are no published studies on the influence of light levels on D/H fractionation in phytoplankton. In
higher plant leaf waxes, Yang et al. (2009) found that D/H
fractionation was about 40% less under continuous light than
under a 12 h light–dark cycle (Yang et al., 2009). And in unpublished batch culture studies with E. huxleyi and G. oceanica,
Benthien et al. (2009) reported a large effect of light intensity
on D/H fractionation in alkenones (Benthien et al., 2009). That
study concluded that D/H fractionation in alkenones was highly
sensitive to light intensity, which in turn controlled growth rate.
Until that work is published, it is impossible to evaluate the role
that light may play in determining hydrogen isotope ratios in
algal lipids. Future studies should make this a high priority,
making use of continuous cultures (chemostats) for controlled
experimentation. Both light and nutrient limitation can control
growth rates, but the effect of light on D/H fractionation in lipids
is likely to be different than that of nutrients. Changes in the flux
of photons directly affect the supply of Hþ generated by PS2,
rather than its demand by PS1. Steady state growth experiments
with chemostats in which growth rates are separately controlled
by light and nutrient levels ought to elucidate the role that each
plays in controlling D/H fractionation in phytoplankton lipids.
12.4.8
Summary and Conclusions
The use of hydrogen isotope ratios in lipids to decipher biogeochemical and climatic processes is expanding rapidly. The
relative success of these efforts depends on an understanding of
the environmental conditions that influence D/H fractionation. Although much remains unknown about the mechanisms of D/H fractionation in phytoplankton and the
primary environmental conditions that influence that fractionation, culture and field experiments conducted to date indicate
that D/H fractionation in algal lipids is sensitive to temperature, salinity, and nitrate-limited growth rate. Specifically, they
indicate that D/H fractionation (1) increases with increasing
temperature, (2) decreases with increasing salinity, and (3)
increases with increasing N-controlled growth rate in
isoprenoid sterols from T. pseudonana, but may be insensitive
to growth rate in acetogenic fatty acids from T. pseudonana.
Acknowledgments
Many of the insights expressed would not have been possible
without the tenacity and creativity of my students and postdocs
during the last 10 years. In particular, I would like to acknowledge the work of Zhaohui Zhang, Valerie Schwab, Rienk
Smittenberg, Katharina Pahnke, Dirk Sachse, Amy Englebrecht,
and Dan Nelson. This material is based upon work supported
by the National Science Foundation under Grant No. OCE1027079.
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