Do betaine lipids replace phosphatidylcholine as fatty acid editing hubs in microalgae?

Abstract

Acyl editing refers to a deacylation and reacylation cycle on a lipid, which allows for fatty acid desaturation and modification prior to being removed and incorporated into other pools. Acyl editing is an important determinant of glycerolipid synthesis and has been well-characterized in land plants, thus this review begins with an overview of acyl editing in plants. Much less is known about acyl editing in algae, including the extent to which acyl editing impacts lipid synthesis and on which lipid substrate(s) it occurs. This review compares what is known about acyl editing on its major hub phosphatidylcholine (PC) in land plants with the evidence for acyl editing of betaine lipids such as diacylglyceryltrimethylhomoserine (DGTS), the structural analog that replaces PC in several species of microalgae. In land plants, PC is also known to be a major source of fatty acids and diacylglycerol (DAG) for synthesis of the neutral lipid triacylglycerol (TAG). We review the evidence that DGTS contributes substantially to TAG accumulation in algae as a source of fatty acids, but not as a precursor to DAG. We conclude with evidence of acyl editing on other membrane lipid substrates in plants and algae apart from PC or DGTS, and discuss future analyses to elucidate the role of DGTS and other betaine lipids in acyl editing in microalgae.

Full text

Do betaine lipids replace
phosphatidylcholine as fatty
acid editing hubs in microalgae?
Danielle Yvonne Hoffmann *and Yair Shachar-Hill
Department of Plant Biology, Michigan State University, East Lansing, MI, United States
Acyl editing refers to a deacylation and reacylation cycle on a lipid, which allows for
fatty acid desaturation and modification prior to being removed and incorporated
into other pools. Acyl editing is an important determinant of glycerolipid synthesis
and has been well-characterized in land plants, thus this review begins with an
overview of acyl editing in plants. Much less is known about acyl editing in algae,
including the extent to which acyl editing impacts lipid synthesis and on which lipid
substrate(s) it occurs. This review compares what is known about acyl editing on its
major hub phosphatidylcholine (PC) in land plants with the evidence for acyl
editing of betaine lipids such as diacylglyceryltrimethylhomoserine (DGTS), the
structural analog that replaces PC in several species of microalgae. In land plants,
PC is also known to be a major source of fatty acids and diacylglycerol (DAG) for
synthesis of the neutral lipid triacylglycerol (TAG). We review the evidence that
DGTS contributes substantially to TAG accumulation in algae as a source of fatty
acids, but not as a precursor to DAG. We conclude with evidence of acyl editing on
other membrane lipid substrates in plants and algae apart from PC or DGTS, and
discuss future analyses to elucidate the role of DGTS and other betaine lipids in acyl
editing in microalgae.
KEYWORDS
microalgae, lipid, acyl editing, phosphatidylcholine, triacylglyceride (TAG)
Abbreviations: BTA1, Betaine Lipid Synthase 1; CPT, CDP-choline:1,2-Diacylglycerol Cholinephosphotransferase;
CrLAT1, C. reinhardtii Lysolipid Acyltransferase 1; DAG, Diacylglycerol; DGAT, Diacylglycerol Acyltransferase;
DGDG, Digalactosyldiacylglycerol; DGTA, Diacylglycerylhydroxymethyltrimethyl-b-Alanine; DGTS, Diacylglyceryl-
N,N,N-trimethylhomoserine; ER, Endoplasmic Reticulum; FA, Fatty Acid; FFA, Free Fatty Acid; FAD, Fatty Acid
Desaturase; G3P, Glycerol 3-Phosphate; GPAT, Glycerol-3-Phosphate Acyltransferase; GPC, Glycerolphosphocholine;
GPCAT, Glycerolphosphocholine Acyltransferase; LC-MS, Liquid Chromatography Mass Spectrometry; LPAAT,
Lysophosphatidic Acid Acyltransferase; LPCAT, Lysophosphatidylcholine Acyltransferase; LPCT,
Lysophosphatidylcholine Transacylase; LPEAT, Lysophosphatidylethanolamine Acyltransferase; LPLAT,
Lysophospholipid Acyltransferase; MGDG, Monogalactosyldiacylglycerol; PA, Phosphatidic Acid; PC,
Phosphatidylcholine; PDAT, Phospholipid : Diacylglycerol Acyltransferase; PDCT, Phosphatidylcholine :
Diacylglycerol Cholinephosphotransferase; PE, Phosphatidylethanolamine; PG, Phosphatidylglycerol; PGD1, Plastid
Galactoglycerolipid Degradation 1; PLIP1, Plastid Lipase 1; PUFA, Polyunsaturated Fatty Acid; SQDG,
Sulfoquinovosyldiacylglycerol; TAG, Triacylglycerol; TGD, Trigalactosyldiacylglycerol.
Frontiers in Plant Science frontiersin.org01
OPEN ACCESS
EDITED BY
Ben Lucker,
Prosel Biosciences, United States
REVIEWED BY
Agnieszka Zienkiewicz,
Nicolaus Copernicus University in Torun
´,
Poland
Juliette Jouhet,
UMR5168 Laboratoire de Physiologie
Cellulaire Vegetale (LPCV), France
*CORRESPONDENCE
Danielle Yvonne Hoffmann
youngd34@msu.edu
SPECIALTY SECTION
This article was submitted to
Marine and Freshwater Plants,
a section of the journal
Frontiers in Plant Science
RECEIVED 22 October 2022
ACCEPTED 04 January 2023
PUBLISHED 19 January 2023
CITATION
Hoffmann DY and Shachar-Hill Y (2023) Do
betaine lipids replace phosphatidylcholine
as fatty acid editing hubs in microalgae?
Front. Plant Sci. 14:1077347.
doi: 10.3389/fpls.2023.1077347
COPYRIGHT
© 2023 Hoffmann and Shachar-Hill. This is
an open-access article distributed under the
terms of the Creative Commons Attribution
License (CC BY). The use, distribution or
reproduction in other forums is permitted,
provided the original author(s) and the
copyright owner(s) are credited and that
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reproduction is permitted which does not
comply with these terms.
TYPE Review
PUBLISHED 19 January 2023
DOI 10.3389/fpls.2023.1077347

Introduction
Research in algal lipid metabolism has undergone a resurgence in
recent years (Merchant et al., 2012;Du and Benning, 2016;Goncalves
et al., 2016), particularly due to interest in the neutral lipid
triacylglycerol (TAG), which is a source of nutritionally valuable
fatty acids and a key feedstock for biodiesel fuel production. Both the
quantity and composition of TAG are important and are governed by
fluxes through intermediate glycerolipids pools such as membrane
lipids (Goncalves et al., 2013;Legeret et al., 2016;Young and Shachar-
Hill, 2021). The process of acyl editing, which is used to mean the
addition and removal of an acyl group on a lipid with the potential for
tailoring and modification and subsequent incorporation into another
lipid pool, plays a central role in fluxes between membrane lipids and
TAG. Much of what is currently known about acyl editing has been
determined in land plants, which utilize the extraplastidial lipid
phosphatidylcholine (PC) as the main hub of acyl editing and fatty
acid modification, and which also contributes to TAG synthesis.
However, a range of algae lack PC and rather contain ether-linked
betaine lipids (Kato et al., 1996;Künzler and Eichenberger, 1997),
thus raising the question: To what extent do betaine lipids replace the
functions of PC in algae? A better understanding of acyl editing is
crucial to efforts in algal lipid engineering; improving the yield and
composition of algal TAG without this knowledge is likely to remain a
hit-or-miss affair.
PC serves as the major acyl hub in plants, where it is the
predominant lipid outside of the chloroplast in most plant cells. PC
plays a central role in several aspects of plant lipid metabolism by
acting as the major substrate for extraplastidial acyl modifications, in
lipid trafficking between subcellular compartments, and in
conducting fluxes toward TAG synthesis (Miquel and Browse, 1992;
Bates et al., 2012;Chen et al., 2015). In many microalgal species,
betaine lipids are present in inverse proportion to PC levels, replacing
it completely in some cases and partially in others. Due to this inverse
relationship and the structural similarity of betaine lipids and PC
(Figure 1), it has been hypothesized that betaine lipids play analogous
roles to PC in algae. The betaine lipid diacylglyceryl-N,N,N-
trimethylhomoserine (DGTS) is widely distributed in nonflowering
groups such as green algae, lichens, mosses, and ferns and is absent in
seed plants and flowering plants (Künzler and Eichenberger, 1997),
suggesting that the capacity to synthesize DGTS was lost in the
spermatophyte ancestor to seed plants. Some bacteria are capable of
synthesizing DGTS in response to phosphate starvation (Geiger et al.,
1999), and DGTS is present in some fungi as well (Künzler and
Eichenberger, 1997). It is believed that a green algae ancestor was
capable of producing DGTS, thus creating a “DGTS branch.”
However, some algae in this branch lack DGTS, indicating
that some organisms either reduced or lost the capacity for
betaine lipid synthesis. The replacement of PC with DGTS may
have evolved as a stress acclimation response to low phosphorus or
low temperature environments.
In this review, we compare the roles of PC in plant lipid
metabolism with what is known about the role of betaine lipids,
particularly DGTS, in algae. The first sections below describe the
metabolic roles of PC in membrane synthesis and oil accumulation in
plants. The subsequent sections present evidence in favor and against
DGTS replacing the different functions of PC in algae and discuss
alternative acyl editing substrates apart from PC or DGTS. The
evidence outlined for PC’s role in plants serves as a pointer to
methods that have or can be used in the future to address these
questions in algae. Rational engineering of algal lipid metabolism such
as TAG accumulation and composition will require knowledge of
their acyl hubs, as evidenced by prior attempts to engineer the
accumulation of particular fatty acids in oilseeds, in which flux
through PC represented a major bottleneck (Bates and Browse, 2011).
Overview of acyl editing in plants
The process of acyl editing is an important determinant of
eukaryotic membrane lipid synthesis as well as TAG accumulation
and has been well-characterized in land plants. Therefore, we begin
this review with an overview of acyl editing and related functions in
plants. The existence of a deacylation and reacylation cycle on PC, the
most abundant membrane lipid in animals and the major
extraplastidic one in plants, was originally inferred from
14
C-tracer
experiments. Supplying rat lung tissue with
14
C-acetate and
14
C-
glycerol resulted in a ratio of radioactivity in fatty acids compared to
glycerol that was much higher in phospholipids than in TAG,
implying that turnover of fatty acids occurred on PC (Lands, 1958).
It was hypothesized that this occurred via the action of a
phospholipase generating lysophosphatidylcholine (lyso-PC, PC
with one fatty acid removed), which was then reacylated to form
PC (Lands, 1960). In microsomes of developing safflower cotyledons,
14
C-labeling revealed rapid exchange of fatty acids between
diacylglycerol (DAG) and PC (Stobart and Stymne, 1985), and
evidence was found that this acyl exchange was catalyzed by the
forward and reverse reactions of acyl-CoA:lysophosphatidylcholine
acyltransferase (LPCAT) (Stymne and Stobart, 1984). Thus, acyl
exchange on PC, termed “the Lands’cycle,”could proceed via two
possible mechanisms: the action of a phospholipase followed by
reacylation by LPCAT, or by both forward and reverse reactions
catalyzed by LPCAT (Figure 2).
A study utilizing
14
C-acetate and
14
C-glycerol labeling in pea
leaves found that the majority of newly made fatty acids were rapidly
incorporated into PC rather than proceeding via the de novo synthesis
FIGURE 1
Structures of PC and DGTS. Figure reproduced with permission from
Klug and Benning, 2001. Copyright (2001) National Academy of
Sciences, U.S.A. DGTS, diacylglyceryl-N,N,N-trimethylhomoserine;
PC, phosphatidylcholine.
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pathway (Bates et al., 2007). Here, “de novo synthesis”refers to the
Kennedy pathway in which glycerol-3-phosphate (G3P) is acylated to
form lyso-phosphatidic acid, which is then acylated to form
phosphatidic acid (PA), whose phosphate group is removed to form
DAG. PC is then synthesized from DAG via cytidine-5′-
diphosphocholine:diacylglycerol cholinephosphotransferase (CPT).
This labeling study in pea leaves found that over 90% of
14
C-
labeled PC molecules contained one
14
C-labeled fatty acid and one
unlabeled fatty acid, and 62% of the label was found at the sn-2
position and 38% at the sn-1 position (Bates et al., 2007). PC acyl
editing was found to be the major route of newly made fatty acid flux
in plant leaves (Bates et al., 2007), a process that allows
polyunsaturated fatty acids (PUFAs) to be produced on PC and
then distributed to other lipids.
Labeling with
14
C-acetate and
14
C-glycerol during TAG synthesis
in developing soybean embryos also revealed the direct incorporation
of newly made fatty acids into PC similar to that observed in pea
leaves (Bates et al., 2009). In addition, 86% of newly synthesized fatty
acids in PC were found at the sn-2 position, suggesting that acyl
editing occurs primarily at this location (Bates et al., 2009). It was
concluded that the major flux of newly made fatty acids proceeded
through PC via acyl editing in this oilseed, highlighting the
importance of acyl editing in lipid synthesis in TAG accumulation
as well as membrane synthesis.
Phosphatidylcholine serves as
the substrate of fatty acid modification
and key intermediate in TAG synthesis
in plants
In plants, PC is the major site of extraplastidial desaturation to
produce PUFAs, as studies have demonstrated that oleic acid (18:1D9)
is desaturated to form linoleic acid (18:2D9,12) in a lipid-linked
manner on PC (Sperling and Heinz, 1993;Sperling et al., 1993;
Figure 3). This work found that desaturation could occur at the sn-
1or sn-2 position of PC based on extraction of PC from in vivo cell
cultures followed by treatment with a stereospecific lipase (Sperling
et al., 1993), but given that in PC polyunsaturated C18-fatty acids are
more abundant at the sn-2 position,thisisthemajorsiteof
desaturation on PC in planta. Desaturation of 18:1 to 18:2 on PC is
catalyzed by the desaturase Fatty Acid Desaturase 2 (FAD2), while
desaturation of 18:2 to 18:3 on PC is catalyzed by FAD3 (Figure 3).
These desaturase genes were discovered by isolating Arabidopsis
mutants deficient in the fatty acid resulting from their activity
(Miquel and Browse, 1992;Browse et al., 1993), and these genes
were cloned and demonstrated to complement the mutant phenotype
(Arondel et al., 1992;Okuley et al., 1994). Thus, it is believed that PC
is synthesized from 16:0/18:1 or 18:1/18:1 DAG, with 16:0 primarily
at the sn-1 position while 18:1 can occur at both the sn-1 and sn-2
positions, after which 18:1 is desaturated to form 18:2 followed by
18:3 (Figure 3;Browse and Somerville, 1991). Given that there is a
lack of evidence for DAG or other lipids in the endoplasmic reticulum
(ER) serving as substrates for desaturation, PC is believed to be the
major site of fatty acid desaturation in this membrane system.
Thus, PC is the site of extraplastidial desaturation, and acyl flux
through PC is a major route for PUFA incorporation into TAG. The
fatty acids of PC can reach TAG by three major routes as shown in
Figure 4: 1) Fatty acids may be removed from PC by a phospholipase
or acyltransferase and enter the acyl-CoA pool, where they are
then available for glycerolipid synthesis. 2) PC may be
converted into DAG, the precursor for TAG synthesis, by the
reverse action of cytidine-5′-diphosphocholine:diacylglycerol
cholinephosphotransferase (CPT), by phosphatidylcholine:
diacylglycerol cholinephosphotransferase (PDCT), or by the action
of phospholipase C or phospholipase D followed by phosphatidic acid
phosphatase to remove the phosphate headgroup. 3) A fatty acid may
be directly transferred from PC onto the sn-3 position of DAG to
produce TAG by the action of phospholipid:diacylglycerol
acyltransferase (PDAT) (Dahlqvist et al., 2000;Stahl et al., 2004).
Each of these routes of acyl flux from PC can contribute to increase
the PUFA content in TAG.
In addition to desaturation, PC also acts as the substrate for
several unusual fatty acid modifications (Figure 3). Mutations in plant
fatty acid desaturases can give rise to alternate enzymatic activities
such as hydroxylation, epoxygenation, and triple-bond formation. For
example, many variants of the FAD2 desaturase, which desaturates
oleic acid (18:1) to linoleic acid (18:2) on PC, have been discovered. In
castor bean, a hydroxylase acts on the D12 position of oleic acid on the
sn-2 position of PC to produce ricinoleic acid, and this hydroxylase is
FIGURE 3
Desaturation and production of unusual fatty acids on PC.
Desaturation and unusual fatty acid synthesis occurs on the sn-2
position of PC. Here ‘18:X’refers to a fatty acid with 18 carbons and an
unspecified number of double bonds. FAD, Fatty Acid Desaturase;
PC, phosphatidylcholine.
FIGURE 2
The Lands Cycle. Cycle of deacylation of PC via either a phospholipase
or LPCAT to produce lyso-PC, which is then reacylated by LPCAT to
regenerate PC. LPCAT, acyl-CoA:lysophosphatidylcholine
acyltransferase; PC, phosphatidylcholine.
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a homolog of FAD2 (Bafor et al., 1991;Van De Loo et al., 1995). An
acetylenase in Crepis alpina is a variant of FAD2 that introduces a
triple bond at the D12 position of linoleic acid on PC to form
crepenynic acid, and this unusual fatty acid then accumulates in
TAG (Banas et al., 1997). In addition, a D12-epoxygenase from Crepis
palaestina catalyzes the formation of vernolic acid from linoleic acid,
presumably on PC (Lee et al., 1998). Thus, PC is the key substrate for
synthesizing PUFAs as well as uncommon fatty acids.
Although PC is the site of synthesis of unusual fatty acids, some
plants contain high amounts of unusual fatty acids in TAG while PC
contains low levels of these fatty acids. For example, in developing
endosperms of castor bean, the hydroxylated fatty acid ricinoleic acid
accumulates to only 5% in PC while it accumulates to ~85% in TAG
despite its synthesis on PC (Stahl et al., 1995). Thus, acyl flux through
the sn-2 position of PC must occur with high efficiency and selectivity.
However, when the castor bean hydroxylase is transgenically
expressed in Arabidopsis, hydroxy-fatty acids only accumulate to
~17% in seed TAG (Broun and Somerville, 1997). Isotopic labeling
analyses using
14
C-glycerol revealed a ~50% reduction in label
incorporation in total lipids, primarily due to a reduction in the use
of de novo DAG for PC synthesis (Bates and Browse, 2011). Thus, flux
through PC can represent a bottleneck for the accumulation of
particular fatty acids in TAG (Bates and Browse, 2011).
Another example of PC acting as a control point for fluxes of
unusual fatty acids is the unusual fatty acid petroselinic acid (18:
lcisD6), which is present in low levels in PC (15-20%) and high levels
in TAG (70-75%) in both carrot and coriander seed endosperm
(Cahoon and Ohlrogge, 1994). However,
14
C-acetate time course
labeling experiments revealed that at early time points, radiolabel was
the most concentrated in PC and entered it at the highest rates, while
at later time points radiolabel accumulated most strongly in TAG, and
primarily (80-85%) in petroselinic acid (Cahoon and Ohlrogge, 1994).
These results suggest that there is significant flux of petroselinic acid
from PC into TAG. Similar results were found in Thunbergia alata,
whose unusual monoenoic fatty acid 16:1D6 comprises >80% of TAG
fatty acids and which appears first in PC (Schultz and Ohlrogge,
2000). Therefore, in order to engineer high levels of select fatty acids
in transgenic oilseeds or algae, the major fluxes of TAG synthesis,
including the sites of modification and distribution, must
be elucidated.
Mechanisms of acyl editing and acyl
flux through PC into TAG
PC conversion into DAG for TAG synthesis has been shown to be
an important biochemical pathway in oilseeds. For instance,
14
C-
acetate and
14
C-glycerol labeling of developing soybean embryos
demonstrated that over 95% of TAG synthesis was found to be
derived from DAG generated from PC, or “PC-derived DAG”
(Bates et al., 2009;Figure 5). This TAG synthesis pathway was also
found to be significant in Arabidopsis seeds, with
14
C-glycerol labeling
revealing that PC-derived DAG is utilized for as much as 93% of TAG
synthesis (Bates and Browse, 2011). Thus, in several oilseeds de novo-
synthesized DAG is converted to PC, which is converted back to
DAG, which produces TAG (Figure 5). However, developing castor-
bean endosperm appears to utilize de novo-synthesized DAG to
produce TAG (Bafor et al., 1991), so this pathway is not
ubiquitously utilized by all plants.
PC-derived DAG can be synthesized by the reverse action of CPT,
which produces PC (Slack et al., 1983). PC-DAG interconversion can
FIGURE 5
PC-derived DAG pathway to TAG synthesis. Black text and arrows
represent the de novo synthesis pathway to PC and TAG, blue text and
arrows represent the PC-derived DAG pathway to produce TAG. The
enzyme PDCT is shown in bold because it is believed to catalyze the
major PC-DAG conversion. CPT, cytidine-5′-diphosphocholine:
diacylglycerol cholinephosphotransferase; DAG, diacylglycerol; DGAT,
diacylglycerol acyltransferase; G3P, glycerol 3-phosphate; PC,
phosphatidylcholine; PDAT, phospholipid:diacylglycerol
acyltransferase; PDCT, phosphatidylcholine:diacylglycerol
cholinephosphotransferase; TAG, triacylglycerol.
FIGURE 4
Routes of acyl flux through PC into TAG. Orange arrows, PC acyl editing reactions. Blue arrows, headgroup exchange for PC-DAG conversion. Green
arrows, transfer of an acyl group from PC onto TAG via the acyltransferase PDAT. CPT, cytidine-5′-diphosphocholine:diacylglycerol
cholinephosphotransferase; DAG, diacylglycerol; DGAT, diacylglycerol acyltransferase; G3P, glycerol 3-phosphate; LPC, lyso-phosphatidylcholine;
LPCAT, acyl-CoA:lysophosphatidylcholine acyltransferase; PC, phosphatidylcholine; PDAT, phospholipid:diacylglycerol acyltransferase; PDCT,
phosphatidylcholine:diacylglycerol cholinephosphotransferase; PLA, phospholipase; TAG, triacylglycerol.
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also be catalyzed by PDCT, which reversibly exchanges the
phosphocholine headgroup between PC and DAG. DAG may also
be produced from PC via the action of phospholipase C, or via
phospholipase D followed by phosphatidic acid phosphatase.
However, characterization of the Arabidopsis rod1 mutant, which
encodes PDCT, revealed that at least 40% of the PUFAs in TAG are
derived from PC-derived DAG produced by this enzyme (Lu et al.,
2009). Thus, PDCT catalyzes the major PC-DAG interconversion in
developing Arabidopsis seeds (Figure 5).
In addition to headgroup removal, PC can also contribute to TAG
synthesis via the well-known PDAT enzyme, which has been
characterized in yeast, oilseeds, and other plant tissues (Dahlqvist
et al., 2000;Stahl et al., 2004). PDAT catalyzes an acyl-CoA
independent pathway of TAG formation by transferring an acyl
group from the sn-2 position of PC onto DAG (Figure 4). PDAT is
known to conduct significant flux into TAG in yeast and oilseeds,
although knockout of PDAT in Arabidopsis resulted in no statistically
significant difference in seed lipid content, likely due to compensation
by diacylglycerol acyltransferase (DGAT) activity (Mhaske et al.,
2005). In microalgae, pdat insertional mutants appeared to
accumulate 25% less TAG during nitrogen deprivation compared to
the parental strain (Boyle et al., 2012), suggesting that it may be a
determinant of TAG accumulation in algae. However, artificial
miRNA-silenced PDAT knockdowns in Chlamydomonas reinhardtii
did not show significant reduction of TAG under N-deprivation
(Yoon et al., 2012), and further characterization of C. reinhardtii
pdat knockout mutants suggested that PDAT primarily regulates
TAG biosynthesis under favorable conditions rather than persistent
stress conditions (Lee et al., 2022). Thus, PDAT may play a
diminished role in TAG synthesis in algae compared to yeast
and plants.
Two studies probed the enzymes underlying the acyl editing
mechanism in Arabidopsis seeds by generating double mutants in
both LPCAT genes of Arabidopsis (lpcat1/lpcat2)(Bates et al., 2012;
Wang et al., 2012), which deacylate and reacylate lyso-PC and PC in
the acyl editing cycle (Figure 2). Both studies observed a reduction of
PUFA content in seeds by ~10% in the lpcat1/lpcat2 double mutant
(Bates et al., 2012;Wang et al., 2012). In addition, both studies used
14
C-labeling to confirm that in the double mutants, nascent fatty acids
were incorporated in the de novo synthesis route of DAG followed by
PC rather than by direct incorporation into PC characteristic of acyl
editing (Bates et al., 2012;Wang et al., 2012). Both studies also found
that the lpcat1/lpcat2 double mutants accumulated lyso-PC,
indicating they were impaired in their ability to reacylate lyso-PC
to PC. Thus, these studies provided strong evidence that the LPCAT
genes are crucial components of the acyl editing cycle in Arabidopsis.
In addition, Bates et al. generated a triple mutant lpcat1/lpcat2/rod1
that was also deficient in PDCT, which interconverts DAG and PC.
The triple mutant’s PUFA content in seed TAG was reduced by about
two-thirds (Bates et al., 2012), suggesting that PDCT and the LPCAT
genes are responsible for the majority of fatty acid flux in and out of
PC, via PC-DAG interconversion by PDCT and acyl editing
by LPCAT.
Early evidence in microsomes of developing safflower cotyledons
suggested that acyl exchange on PC was catalyzed by the forward and
reverse reactions of LPCAT (Stymne and Stobart, 1984), and this was
later confirmed by expressing Arabidopsis LPCAT2 in yeast and
measuring LPCAT activity by the rate of incorporation of
14
C-18:1
fatty acid into PC (Lager et al., 2013). These assays revealed that
LPCAT1 and LPCAT2 can catalyze both the acylation and
deacylation of PC (Lager et al., 2013). While both LPCATs can
acylate and deacylate at the sn-1 position, the sn-2 position is
strongly preferred (Lager et al., 2013). Both LPCAT enzymes
showed low activity toward 16:0 fatty acid, and thus are probably
not involved in acylating or deacylating 16:0 on PC (Lager et al.,
2013). Thus, LPCAT is reversible in vitro, although it is still possible
that acyl editing could proceed via the action of a phospholipase
followed by the forward reaction of LPCAT to regenerate PC.
In addition to LPCAT reversibility, Lager et al. used microsomes
of developing safflower seeds to characterize other reactions involved
in PC acyl editing. By performing isotopic labeling experiments with
[
14
C]18:1-lyso-PC and [
14
C]choline, they demonstrated that lyso-PC:
lyso-PC transacylation occurred, and named this activity
lysphosphatidylcholine transacylase (LPCT) (Lager et al., 2015).
Thus, LPCT activity utilizes two lyso-PC molecules and produces
PC and glycerophosphocholine (GPC; Figure 6A). By incubating
microsomal preparations of various developing oilseeds in [
14
C]
GPC and 18:1-CoA and measuring the radioactive PC product,
acyl-CoA:glycerophosphocholine acyltransferase (GPCAT) activity
was also demonstrated (Lager et al., 2015), which catalyzes the
acylation of GPC to lyso-PC (Figure 6B). The enzyme responsible
for LPCT activity has yet to be identified, but the GPCAT gene has
been identified and cloned in Arabidopsis, and GPCAT homologs
have been found across other eukaryotic clades including green algae
(Głab et al., 2016).
Many studies on developing seeds are performed in vitro, and one
study sought to compare in vitro versus in vivo growth on acyl lipid
metabolism in Camelina sativa leaves (Klin
ska et al., 2021).
Significant differences were observed between the two growth
conditions, with in vivo leaves containing over twice the total fatty
acid content of in vitro leaves, in vivo conditions resulting in a higher
degree of desaturation in fatty acids compared to in vitro, and the
dominant lipid in in vitro conditions being PC rather than
monogalactosyldiacylglycerol (MGDG) (Klin
ska et al., 2021).
Interestingly, higher activity of acyl-CoA:lysophospholipid
A
B
FIGURE 6
Additional PC acyl editing reactions. (A) Lyso-PC:lyso-PC transacylation
via LPCT activity. (B) Acylation of GPC by GPCAT activity. GPC,
glycerophosphocholine; GPCAT, acyl-CoA:glycerophosphocholine
acyltransferase; LPCT, lysphosphatidylcholine transacylase;
PC, phosphatidylcholine.
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acyltransferases (LPLATs) was observed in vitro,aswellasan
increased rate of phospholipid remodeling (Klinska et al., 2021).
Collectively, these results indicate that acyl lipid metabolism and
acyl editing differ substantially between in vitro and in vivo
conditions, and results from one condition should not be
extrapolated to the other.
Acyl editing is also known to occur in response to stress, such as
temperature stress, salt stress, and nutrient deprivation. For instance,
lipidomic profiling during chilling stress in barley roots revealed that
as particular molecular species of PC decreased, those molecular
species increased in PA (Vilchez et al., 2021). Transcriptional analysis
of genes involved in acyl editing revealed that the barley LPCAT
gene was upregulated within five hours of cold recovery, and
phospholipase D activity rose during cold treatment and after five
hours of cold recovery (Vilchez et al., 2021). Thus, it is likely that PC
is hydrolyzed to produce PA during cold stress and recovery, and that
acyl editing of PC plays an important role in chilling stress
and recovery.
Betaine lipids may replace PC as a fatty
acid editing hub in algae
In flowering plants, PC is the major extraplastidial membrane
lipid, but some algae and nonflowering plants contain DGTS and/or
other betaine lipids in place of PC as the major extraplastidial lipid.
DGTS is a betaine lipid that is structurally highly similar to PC, but it
lacks phosphate in its headgroup and rather contains an ether bond
linking the headgroup to the glycerol backbone (Figure 1). Due to its
structural similarity to PC, DGTS is widely believed to replace the
function of PC in extrachloroplastic membranes, and several studies
in algae and diatoms containing DGTS have found evidence that
DGTS may substitute for several of the functions of PC.
The green microalga C. reinhardtii is the most studied alga in lipid
and other areas of metabolism. C. reinhardtii lacks PC and rather
contains DGTS, and an inverse relationship between the quantities of
these two lipids occurs in many species of microalgae (Dembitsky,
1996). When
14
C-oleic acid is supplied exogenously to C. reinhardtii
cells, label is rapidly incorporated into DGTS and it contains the
majority of the radiolabel, similar to that which is observed in PC in
flowering plants (Schlapfer and Eichenberger, 1983;Giroud
and Eichenberger, 1989). The golden-brown microalga
Ochromonas danica contains the betaine lipids DGTS and
diacylglycerylhydroxymethyltrimethyl-b-alanine (DGTA), and small
amounts of PC. When this alga was incubated with
14
C-oleic acid, the
majority of the radiolabel was incorporated into DGTS, suggesting
that DGTS is the primary acceptor of exogenous oleic acid (Vogel and
Eichenberger, 1992). Similar findings were observed in the brown
algae Fucus vesiculosus and Ascophyllum nodosum, which contain
DGTA and minor quantities of PC. In these algae, labeling with
14
C-
acetate revealed DGTA to be the lipid with the highest incorporation
of radioactivity (Jones and Harwood, 1993). Thus, in algae containing
ether-linked betaine lipids in place of PC, supplying exogenous
labeled fatty acids or acetate results in the early and strong
incorporation of label into betaine lipid, analogous to the rapid
incorporation of radiolabel into PC observed in land plants.
In addition to being the first major lipid into which exogenous
radiolabel is incorporated, DGTS also appears to be the site of
extraplastidial desaturation of C18 fatty acids akin to PC. When
14
C-oleic acid was supplied exogenously to C. reinhardtii, radiolabel
first appeared in molecular species of DGTS containing 18:1, and then
shifted to species containing 18:2, followed by species containing
18:3D5,9,12, suggesting that C18 fatty acid desaturation occurs on
DGTS (Schlapfer and Eichenberger, 1983;Giroud and Eichenberger,
1989). Similar results were obtained in a pulse-chase labeling
experiment in the golden-brown microalga O. danica,inwhich
labeling with
14
C-oleic acid resulted in radiolabel being primarily
concentrated in DGTS in its 18:1 and 18:2 fatty acids (Vogel and
Eichenberger, 1992). During the chase, radiolabel decreased very
rapidly in 18:1 while it decreased more slowly in 18:2 fatty acids,
and radiolabel increased strongly in 18:3 and 18:4 fatty acids (Vogel
and Eichenberger, 1992). This suggested that 18:1 is desaturated on
DGTS to produce 18:3 and 18:4 fatty acids. Thus, in algae containing
DGTS rather than PC, DGTS appears to take on PC’s role as a
substrate of extraplastidial fatty acid desaturation.
Labeling experiments in oilseed plants using
14
C-acetate have
demonstrated that radiolabeled fatty acids are rapidly incorporated
into PC prior to their incorporation into DAG and TAG (Bates et al.,
2009). Pulse-chase experiments in C. reinhardtii cells supplied with
14
C-radiolabeled fatty acids in nutrient replete medium and then
transferred into unlabeled, nitrogen-deprived medium to induce TAG
accumulation revealed that radiolabel was rapidly lost from DGTS
while it increased in TAG during the chase period (Xu et al., 2016).
Similarly, when the brown alga F. vesiculosus was incubated with
14
C-
acetate, radiolabel was initially most concentrated in the betaine lipid
DGTA, and over time decreased in DGTA while increasing strongly
in neutral lipids, primarily TAG (Jones and Harwood, 1993). Thus,
pulse-chase experiments with radiolabeled substrates indicate that
fatty acids pass through betaine lipids into TAG in algae in a similar
manner as PC in land plants.
In addition to pulse-chase experiments, several lipidomics-based
analyses point to fatty acid flux through betaine lipids into TAG. One
study utilizing subcellular lipidomics to investigate the origins of TAG
formation in nitrogen-deprived C. reinhardtii indicated that the
membrane lipids MGDG, digalactosyldiacylglycerol (DGDG), and
DGTS all contribute acyl chains to TAG accumulation (Yang et al.,
2018). In particular, they noted that as certain molecular species of
DGTS accumulated, there was a corresponding increase in the levels
of those acyl chains in TAG (Yang et al., 2018). In support of this
finding, a glycerolipidomics study of the BAFJ5 starchless mutant in
C. reinhardtii subjected to high light and nitrogen deprivation
concluded that 18:3D9,12,15/16:0 from DGDG and 16:0/18:3D5,9,12
from DGTS were major contributors to 18:3 accumulation in TAG
(Yang et al., 2020). The diatom Phaeodactylum strain Pt4 (UTEX 646)
contains DGTA as the major extraplastidial lipid and minor amounts
of PC (Abida et al., 2015), and analysis of its lipid molecular species
composition during nitrogen deprivation indicated that the betaine
lipid was the largest membrane lipid contributor of fatty acids to TAG
accumulation (Popko et al., 2016). The major TAG species that
accumulated were matched to a reduction in those corresponding
species in MGDG and the betaine lipid, but only the decrease in the
latter was large enough to account for the increase observed in TAG
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(Popko et al., 2016). Thus, several lines of evidence indicate that
betaine lipids serve as a major source of fatty acids for TAG synthesis.
The relationship between DGTS composition and levels and TAG
accumulation was further evident in knockdown transformants in the
betaine lipid synthase gene (BTA1) in C. reinhardtii.Artificial
microRNA knockdown of the BTA1 gene expression level led to a
significant decrease in DGTS and MGDG contents while TAG
increased 2-3 fold (Lee et al., 2017). The authors postulate that the
observed decrease in MGDG was due to the indirect effect of ER stress
due to the substantial decrease of DGTS. Their results indicated that
the synthesis of DGTS was inhibited by the reduced expression of
BTA1, thus resulting in an increased amount of DAG that could not be
converted into DGTS, which was then available for TAG synthesis (Lee
et al., 2017). This was evidenced by a 40% reduction in DGTS content
in the BTA1 gene knockdown lines compared to wild type and empty
vector controls, and a 2-3 fold increase in the amounts of DAG and
TAG (Lee et al., 2017). On the other hand, the reduction in MGDG
was interpreted to be due to induced breakdown due to ER stress
rather than inhibition of its synthesis, thus releasing fatty acids which
could then contribute to TAG synthesis (Lee et al., 2017). Therefore,
the effects of downregulating BTA1 expression suggested that DGTS
plays a significant role in the accumulation of TAG in C. reinhardtii.
Due to structural differences between DGTS and PC, the
mechanism by which DGTS contributes to TAG accumulation
likely differs from that of PC. DGTS contains an ether bond linking
its headgroup to the glycerol backbone (Figure 1), making it unlikely
for DGTS’s headgroup to be chemically or enzymatically removed to
form DAG as occurs in PC. Enzymes capable of breaking this ether
bond have not yet been identified, but are presumed to exist in order
for cells to catabolize DGTS. Moreover, the vast majority of molecular
species of DGTS contain a C18 fatty acid on the sn-2 position (Giroud
et al., 1988) while ~90% of TAG molecular species contain a C16 fatty
acid on the sn-2 position (Fan et al., 2011), thus supporting the idea
that DGTS is not a direct precursor to DAG for TAG assembly.
Therefore, fatty acid flux from DGTS into TAG probably occurs via
the action of lipases or acyltransferases. In support of this hypothesis,
Vogel and Eichenberger, 1992 observed turnover of radiolabel in the
18:2 fatty acid of DGTS that was substantially faster than the turnover
of its polar headgroup. Similarly,
13
C-labeling of C. reinhardtii lipids
followed by an unlabeled chase into nitrogen deprivation revealed
very little turnover of DGTS’s glycerol backbone and homoserine
betaine headgroup compared to its fatty acids (Young et al., 2022). In
addition, several studies have found that the quantity of DGTS stays
constant in C. reinhardtii cells during nitrogen deprivation (Fan et al.,
2011;Yang et al., 2018), despite high rates of TAG accumulation and
the occurrence of fatty acid flux from DGTS into TAG. Therefore, it is
probable that fatty acids from DGTS reach TAG by deacylation rather
than removal of the betaine headgroup to form a DAG molecule that
is then converted into TAG.
On the other hand, there is evidence that MGDG rather than
DGTS is converted into DAG for TAG synthesis in C. reinhardtii.
Lipidomic analysis of heat-stressed C. reinhardtii revealed a strong
decrease in the major MGDG molecular species and an accumulation
of this species in DAG and TAG (Legeret et al., 2016). This finding
was corroborated by their use of a crfad7 mutant, which altered the
major MGDG molecular species and confirmed its decrease under
heat stress and accumulation in DAG and TAG (Legeret et al., 2016).
In accordance with this, Young et al., 2022 found evidence of MGDG
recycling into DAG and TAG in C. reinhardtii during nitrogen
deprivation, as isotopic label decreased in the major MGDG species
and increased in the corresponding species of DAG and TAG. In
addition, the stereochemical positions of the characteristic fatty acids
of the major MGDG species were matched to the identical positions
in DAG and TAG (Young et al., 2022). Data mining of the C.
reinhardtii lipid body proteome (Moellering and Benning, 2010) led
to the identification of a putative galactosyl hydrolase gene (CrGH),
and insertional mutants in this gene have reduced TAG content (Gu
et al., 2021). Thus, the route of acyl flux through PC via headgroup
removal appears to be replaced in microalgae by removal of the
galactosyl headgroup of MGDG rather than headgroup removal
of DGTS.
DGTS appears to play a role in phosphorus-sparing in microalgae.
Under optimal growth conditions, the green alga Chlorella kessleri does
not contain DGTS, and contains PC as the dominant extraplastidic
lipid. Within 48 hours of transfer to phosphorus-deficient conditions,
this alga almost completely replaces the phospholipids PC and
phosphatidylethanolamine (PE) with an equivalent amount of DGTS
(Oishi et al., 2022). Interestingly, the fatty acid composition of DGTS
was nearly identical to that of PC and PE in phosphorus-replete
conditions (Oishi et al., 2022). C. kessleri was found to contain a
homolog of the C. reinhardtii BTA1 gene, and the expression of this
gene appears to be induced under phosphorus-deprived conditions in
order to lower the alga’s phosphorus demand (Oishi et al., 2022).
Similar findings were reported in the heterokont Nannochloropsis
oceanica, which contains both DGTS and PC under nutrient replete
growth conditions. During phosphorus deprivation, PC is reduced by
95% in N. oceanica, while DGTS levels strongly increased, becoming
the most abundant membrane lipid in the cells (Meng et al., 2019).
Upon resupply of phosphorus to the cells, PC levels increased and
DGTS fell to pre-deprivation levels (Meng et al., 2019). This study also
found that DGTS appeared to replace PC as the substrate for C18 fatty
acid desaturation and acyl remodeling under phosphorus deprivation
(Meng et al., 2019). Thus, levels of DGTS and PC are inversely
correlated, and DGTS appears to substitute for PC under
phosphorus-deprived conditions due to it being a non phosphorus-
containing lipid.
DGTS may also aid microalgae in adaptation to lower
temperatures. When the haptophyte microalga Pavlova lutheri was
subjected to a low temperature, an increase in the relative amount of
PUFAs and betaine lipid was observed (Tatsuzawa and Takizawa,
1995). At 15°C, the relative percentage of betaine lipid in P. lutheri
increased four-fold compared to cells grown at 25°C (Tatsuzawa and
Takizawa, 1995). In the heterokont N. oceanica, mutants lacking
either DGTS synthesis (bta1) or PC synthesis were generated in order
to determine which lipid plays a role in adaptation to lower
temperatures (Murakami et al., 2018). Only the bta1 mutant which
lacked DGTS synthesis displayed significantly impaired cell growth at
low temperatures, indicating that DGTS is required for maintaining
optimal growth at low temperatures (Murakami et al., 2018). In both
P. lutheri and N. oceanica, DGTS contains a high a proportion of 20:5
fatty acid, therefore it is postulated that enhanced DGTS under low
temperatures aids in maintaining membrane fluidity by having a high
PUFA content (Tatsuzawa and Takizawa, 1995;Murakami
et al., 2018).
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Evidence for acyl editing substrates in
plants and microalgae apart from PC
and DGTS
Indications of an MGDG acyl editing cycle in C. reinhardtii began
with a study characterizing a galactoglycerolipid lipase gene named
PLASTID GALACTOGLYCEROLIPID DEGRADATION1 (PGD1)
(Li et al., 2012). This gene was discovered in an insertional mutant
screen for lowered TAG content during nitrogen deprivation, and the
insertion was found to be in a putative lipase-encoding gene. In terms
of acyl composition, the pgd1 mutant contained a lower amount of
oleic acid (18:1) in TAG (Li et al., 2012). In vitro lipase assays revealed
MGDG to be the substrate of the PGD1 lipase, and PGD1
preferentially hydrolyzes newly synthesized MGDG (18:1D9/16:0) at
the sn-1 position (Li et al., 2012). Pulse-chase analyses using
14
C-
acetate revealed that the proportion of label remained higher in
MGDG and DGDG in the pgd1 mutant during nitrogen
deprivation, whereas in the wild type the label decreased in
membrane lipids as it increased in TAG (Li et al., 2012). Thus,
PGD1 is a lipase that preferentially releases 18:1D9 from newly-made
MGDG, and the released fatty acid joins the acyl-CoA pool where it is
available to contribute to TAG synthesis (Figure 7).
A follow up study further explored the role of PGD1 in membrane
lipid turnover and remodeling in C. reinhardtii in response to
unfavorable environmental conditions (Du et al., 2018). In this
study, sensitivity of lipid analyses was increased by isolating
chloroplasts, and an increased level of MGDG in pgd1 mutants
compared to wild type was observed (Du et al., 2018). In terms of
MGDG acyl composition, a lower quantity of 16:3 and 18:2 was
observed but a higher amount of 16:4 and 18:3 in pgd1 cells compared
to wild type (Du et al., 2018). This supports the model of PGD1-
mediated turnover of MGDG 18:1D9 into TAG, as pgd1 mutants are
impaired in the flux of 18:1 from MGDG into TAG, thereby allowing
oleate to reside in MGDG longer and thus become desaturated. Thus,
these studies point to an acyl editing cycle on MGDG during
nitrogen-deprived conditions.
Recently, an enzyme that appears to catalyze the reacylation of
lyso-MGDG (MGDG with one fatty acid removed) has been
characterized, and this activity would complete an acyl editing cycle
of MGDG. A homolog of the Arabidopsis thaliana LPCAT1 gene was
discovered in C. reinhardtii (CrLAT1), and this gene contained a
conserved membrane-bound O-acyl transferase (MBOAT) domain
(Iwai et al., 2021). Knockdown CrLAT1 mutants contained an
increased proportion of lyso-MGDG and decreased TAG
accumulation compared to wild type cells (Iwai et al., 2021). In
contrast, overexpression CrLAT1 mutants contained a lower
proportion of lyso-MGDG compared to wild type cells (Iwai et al.,
2021). The most abundant fatty acid in lyso-MGDG was 16:4 with a
minor proportion of other C16 fatty acids, supporting a model in
which CrLAT1 acylates lyso-MGDG with a C18 fatty acid. This acyl
editing mechanism in C. reinhardtii consists of PGD1 hydrolyzing
newly-made 18:1D9/16:0 MGDG, releasing 18:1D9 to the cytosolic
acyl-CoA pool where it is available for TAG synthesis, and CrLAT1
may catalyze the reacylation of lyso-MGDG at the sn-1 position with
a C18 fatty acid (Figure 7).
Evidence of acyl editing on lipid substrates other than PC or
DGTS has also been observed in land plants. In one study, a D6
desaturase from Physcomitrella patens was introduced into
Arabidopsis, which lacks a D6 desaturase. The D6 desaturase is ER-
localized with a preference for the sn-2 position of PC, and indeed in
the transformant ~90% of the D6 fatty acids in PC were located in the
sn-2 position (Hurlock et al., 2018). D6 acyl groups are synthesized in
the ER and must be imported into the chloroplast, and they would
presumably be located at the sn-2 position of chloroplastic lipids if
acyl editing does not occur. However, in each of the chloroplastic
lipids [MGDG, DGDG, sulfoquinovosyldiacylglycerol (SQDG), and
phosphatidylglycerol (PG)], the D6 acyl groups were approximately
equally distributed between the sn-1 and sn-2 positions, indicating
that acyl editing of plastidial lipids does occur (Hurlock et al., 2018).
Interestingly, PA isolated from whole leaf tissues displayed the same
stereochemical distribution of D6 fatty acids as PC, while PA isolated
from chloroplasts displayed the reverse stereochemistry with the
majority of the D6 fatty acids located at the sn-1 position (Hurlock
et al., 2018). Thus, some portion of PA is likely subject to acyl editing
as well. Furthermore, through liquid chromatography mass
spectrometry (LC-MS) a 34:7 MGDG molecular species was
identified that contained one ER-derived fatty acid and one
chloroplast-derived fatty acid, as it contained one 18:4 fatty acid
with a D6 desaturation synthesized in the ER and one 16:3 fatty acid
that is synthesized exclusively in chloroplast via the FAD5 desaturase
(Hurlock et al., 2018). This supports the idea that acyl editing occurs
on MGDG, as one exclusively ER-derived fatty acid and one
exclusively chloroplast-derived fatty acid were found on the same
molecule. However, the transgenic lines appeared to have higher lipid
turnover compared to wild type (Hurlock et al., 2018), therefore it is
possible that increased acyl editing may have been triggered in the
transgenic line.
One study characterizing the function of a phospholipase A
1
,
PLASTID LIPASE1 (PLIP1), in Arabidopsis found evidence of acyl
editing on the plastidic phospholipid PG. PLIP1 contains a conserved
Lipase 3 domain, similar to the PGD1 lipase of C. reinhardtii (Wang
et al., 2017). In vitro assays of PLIP1 revealed that its preferred
substrate is 18:3/16:1D3t PG, and that PLIP1 acts on the sn-1 position
to release the 18:3 fatty acid (Wang et al., 2018). A pulse-chase
experiment using
14
C-acetate in leaves of PLIP1 overexpressor lines in
Arabidopsis revealed rapid incorporation of label in PG compared to
wild type, with PG containing ~70% of the total label by the end of the
FIGURE 7
Acyl editing cycle of MGDG in C. reinhardtii. Removal of 18:1 from
newly made MGDG at the sn-1 position by PGD1, and reacylation of
lyso-MGDG catalyzed by CrLAT1. CrLAT1, C. reinhardtii Lysolipid
Acyltransferase 1; MGDG, monogalactosyldiacylglycerol; PGD1, Plastid
Galactoglycerolipid Degradation 1.
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pulse phase (Wang et al., 2018). During the chase phase, PG rapidly
lost most of its label while PC rapidly accumulated label (Wang et al.,
2017). A
14
C-acetate pulse-chase experiment in developing
Arabidopsis seeds also demonstrated higher incorporation of label
into PG compared to wild type seeds and increased turnover of PG
during the chase period, while label concomitantly turned over in PC
and accumulated heavily in TAG (Wang et al., 2018). Furthermore,
plip1 insertional mutants displayed ~10% lowered seed fatty acid
content (the majority of which is in TAG) compared to wild type,
while PLIP1 overexpressors had a 40-50% increase in seed fatty acid
content (Wang et al., 2018). Taken together, these results indicate that
PLIP1 transfers a polyunsaturated fatty acid from PG into PC, and
this fatty acid fluxes through PC and contributes to TAG synthesis
(Figure 8). Thus, PLIP1 appears to be part of an acyl editing cycle that
acts on PG and contributes to the flux of PUFAs through PC
into TAG.
In addition, the lipase HEAT INDUCIBLE LIPASE1 (HIL1) in
Arabidopsis was characterized via in vitro lipase activity assays and
found to have a substrate preference for MGDG (Higashi et al., 2018).
Insertional hil1 mutants are impaired in the remodeling of 18:3 into
TAG during heat stress (Higashi et al., 2018), thus indicating that
HIL1 plays a role in the turnover of PUFAs from MGDG to TAG.
Previous studies demonstrated that LPCAT can catalyze both the
acylation and deacylation of PC via the forward and reverse reactions,
respectively (Lager et al., 2013). In vitro assays investigated
the capability of Arabidopsis LPCAT2 and Arabidopsis
lysophosphatidylethanolamine acyltransferase (LPEAT2) to catalyze
the forward and reverse reactions in the phospholipids PE and PA
(Jasieniecka-Gazarkiewicz et al., 2016). This study found that
although both AtLPCAT2 and AtLPEAT2 could catalyze the
reverse reaction (deacylating an intact phospholipid), the activity of
the reverse reaction varied greatly between phospholipids and was the
lowest for PA (Jasieniecka-Gazarkiewicz et al., 2016). However, these
enzymatic assays established the possibility of acyl remodeling of PE
and PA. The substrate specificity and forward and reverse activity of
LPAAT and LPEAT were then assayed in microsomal fractions of
Camelina sativa seeds (Klinska et al., 2020). In terms of substrate
specificity in the forward reaction (i.e. acylating a lyso-phospholipid
to form a phospholipid), 16:0 and unsaturated C18 fatty acids were
the preferred substrates of LPAAT, while unsaturated C18 fatty acids
were the preferred acyl donors in the LPEAT-catalyzed reaction
(Klin
ska et al., 2020). The activity of the reverse reaction
(deacylating a phospholipid) was assayed by incubating
14
C-acyl-
CoAs with PA or PE, and it was found that both LPAAT and LPEAT
could catalyze the backward reaction, although the acyl donor used
affected the degree of activity observed (Klinska et al., 2020).
Although the back-reaction could be performed by LPEAT, PE
displayed a very slow remodeling rate and only contributed about
2% to the fatty acids in mature C. sativa seeds (in which TAG
comprises ~93% of total lipids), despite PE constituting ~13-20% of
all polar lipids (Pollard et al., 2015;Klinska et al., 2020). However, PE
may donate a fatty acid to TAG synthesis via the action of PDAT, and
the resulting lyso-PE could be reacylated with acyl-CoA by LPEAT.
Interestingly, although PA only constitutes a minor fraction of polar
lipids in C. sativa seeds (~2-4%), it appears ~5% of fatty acids in
mature C. sativa seeds are first esterified to PA, before being
transferred to the acyl-CoA pool via LPAATs (Klinska et al., 2020).
This degree of acyl editing on PA is surprising given that desaturation
is not known to occur on PA, although PA constitutes a relatively
small proportion of membrane lipids and ultimately is not a
substantial fatty acid contributor to seed oil content. Thus, PA and
PE are potential substrates of acyl remodeling in plants, although
neither seems to contribute flux as significant as PC.
Discussion
Acyl flux through PC serves several important functions in plants,
including acting as a major site of extraplastidial desaturation, as a
distributor of PUFAs into other lipids via acyl editing, and as an
important hub of acyl flux into the neutral lipid TAG. Interestingly,
the model green microalga C. reinhardtii completely lacks PC, and
many species of microalgae contain highly reduced PC content, thus
raising the question of if and how the functions of PC are replaced in
microalgae. Given their inverse relationship and the structural
similarity of DGTS and other betaine lipids to PC, it seems
reasonable to postulate that betaine lipids substitute for the
functions of PC in microalgae. Consistent with such roles is the
evidence described above of rapid incorporation of exogenous fatty
acids into DGTS, extraplastidial desaturation on DGTS, and
substantial fatty acid flux through DGTS into TAG. However, the
structure of DGTS makes it highly improbable that an analogous
biochemical reaction of PC headgroup removal could occur on
DGTS, and studies have found evidence of slower turnover of the
DGTS backbone and headgroup compared to its fatty acids (Vogel
and Eichenberger, 1992;Young et al., 2022). Additionally, knockout
and characterization of the C. reinhardtii homolog of PDAT have not
shown that DGTS replaces PC as the substrate of this enzyme in algae
(Boyle et al., 2012;Yoon et al., 2012). In fact, phylogenetic analysis of
the PDAT protein revealed the divergent evolution of PDAT in plants
and algae and suggested that PDAT has a diminished and/or different
FIGURE 8
Acyl editing of PG in Arabidopsis contributes to TAG synthesis. The
lipase PLIP1 releases 18:3 from 18:3/16:1D3t PG, which is channeled
into PC and eventually incorporated into TAG. FFA, free fatty acid;
LPC, lysophosphatidylcholine; PC, phosphatidylcholine; PG,
phosphatidylglycerol; PLIP1, Plastid Lipase 1; TAG, triacylglycerol.
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role in green algae (Falarz et al., 2020). Similarly, knockdown of the C.
reinhardtii homolog of Arabidopsis LPCAT revealed altered turnover
of MGDG rather than DGTS (Iwai et al., 2021). Taken together, it
seems likely that DGTS takes on some, but not all, of the roles in algae
that PC has in plants. This is illustrated in Figure 9, which highlights
known and potential pathways of fatty acid editing in algae.
In addition to the functions described above, PC may also play a
role in lipid trafficking. The eukaryotic pathway of galactolipid
synthesis requires transport of some lipid precursor from the ER to
the plastid. The Arabidopsis trigalactosyldiacylglycerol (TGD)
transmembrane protein complex of the chloroplast has been shown
to conduct ER-to-plastid lipid trafficking (Xu et al., 2003;Xu et al.,
2010;Fan et al., 2015), although its transported lipid substrate
remains unknown. Potential candidates for the transported lipid
substrate include PC, lyso-PC, DAG, or PA, but regardless of its
identity the transported lipid is believed to be derived from PC.
Interestingly, recent studies suggest that the PC involved in transport
to the chloroplast for galactolipid synthesis is metabolically distinct
from the PC pool utilized for acyl editing (Karki et al., 2019).
Intriguingly, despite its lack of PC C. reinhardtii contains an
ortholog of the plant TGD transporter called CrTGD2, and isotopic
labeling experiments indicated that Chlamydomonas can import lipid
precursors from the ER (Warakanont et al., 2015). It is postulated that
PA may be the lipid species transported from the ER to the
chloroplast in C. reinhardtii. It remains to be determined what role
DGTS may have in lipid trafficking in algae.
Thus, the extent to which betaine or other lipids replace the roles
of PC in algae is an area ripe for future study. While radiolabeling has
been instrumental in confirming some of the functions of DGTS,
more detailed
13
C isotopic labeling time courses should be performed
in order to trace the fatty acid fluxes of DGTS in algae. The relative
amount of fatty acid modification and export from DGTS into other
lipids such as TAG should be estimated by
13
C and/or other isotopic
labeling to determine its potential contribution as an acyl hub. Prior
13
C-labeling studies have demonstrated that DGTS contributes
substantial fatty acid flux toward TAG synthesis (Young and
Shachar-Hill, 2021), and future studies examining the turnover of
individual lipid components on a finer timescale can be expected to
illuminate the mechanism of DGTS’s contribution to the synthesis of
TAG and other glycerolipids. In addition, we believe that more
detailed analysis of algal homologs of plant enzymes known to
conduct acyl flux through PC including LPCAT, PDAT, and
phospholipase A2 would be valuable. Such analyses should include
in vitro enzymatic assays to ascertain whether DGTS may replace PC
as the preferred substrate of these enzymes. Finally, the use of [
13
C
2
18
O
2
]acetate labeling may shed light onto the role of DGTS and other
lipids as acyl editing hubs, as
18
O fatty acid labeling can help
determine the number of hydrolysis reactions that have taken
place, thus informing the history of a fatty acid and possible routes
of its trafficking (Pollard and Ohlrogge, 1999). Given the numerous
critical functions of PC in plant lipid metabolism, determining
the extent to which betaine or other lipids replace the function of
FIGURE 9
Working model of lipid metabolism in C. reinhardtii during TAG accumulation. Enzymes are shown in italics, for those shown in red further evidence is
needed to test their role(s) and/or substrates in vivo. A dashed arrow indicates that for DGTS, headgroup removal or exchange is unlikely. Biosynthesis
reactions of extraplastidial lipids DGTS, PE, and PI are omitted for simplicity. ACP, acyl carrier protein; CDP-DAG, cytidine diphosphate diacylglycerol;
CDS, CDP-DAG synthase, CrGH, C. reinhardtii galactosyl hydrolase; CrLAT1, C. reinhardtii Lysolipid Acyltransferase 1; DAG, diacylglycerol; DGAT,
diacylglycerol acyltransferase; DGDGS, digalactoslydiacylglycerol synthase; DGTS, diacylglyceryl-N,N,N-trimethylhomoserine; FAS, fatty acid synthase;
FAT, fatty acyl-ACP thioesterase; FFA, free fatty acid; G3P, glycerol 3-phosphate; Gal, galactose; GPAT, glycerol-3-phosphate acyltransferase; HIL1, heat
inducible lipase 1; LACS, long chain acyl-CoA synthetase; LPA, lysophosphatidic acid; LPAAT, lysophosphatidic acid acyltransferase; LPCAT, acyl-CoA:
lysophosphatidylcholine acyltransferase; LPEAT, lysophosphatidylethanolamine acyltransferase; MGDG, monogalactosyldiacylglycerol; MGDGS,
monogalactosyldiacylglycerol synthase; PA, phosphatidic acid; PAP, phosphatidic acid phosphatase; PDAT, phospholipid:diacylglycerol acyltransferase;
PE, phosphatidylethanolamine; PG, phosphatidylglycerol; PGD1, Plastid Galactoglycerolipid Degradation 1; PGPP, phosphatidylglycerol phosphate
phosphatase; PGPS, phosphatidylglycerolphosphate synthase; PLA, phospholipase; PLIP2, 3, plastid lipase 2, 3; SQD1, 2, sulfoquinovosyldiacylglycerol
synthase; SQDG, sulfoquinovosyldiacylglycerol; TAG, triacylglycerol.
Hoffmann and Shachar-Hill 10.3389/fpls.2023.1077347
Frontiers in Plant Science frontiersin.org10

PC in algae is important for understanding and engineering algal
lipid metabolism.
Author contributions
DH and YS-H discussed and conceived of the ideas in this paper.
DH researched, wrote the article, and prepared the figures. YS-H
contributed to the final form of the manuscript. All authors
contributed to the article and approved the submitted version.
Funding
This work was supported by the National Institute of General
Medical Sciences of the National Institutes of Health predoctoral
training award from grant no. T32-GM110523 to DH. Its contents are
solely the responsibility of the authors and do not necessarily
represent the official views of the NIGMS or NIH. This work was
also made possible by a University Distinguished Fellowship awarded
to DH by the Graduate School of Michigan State University. This
research was supported by the Office of Science (BER), U.S.
Department of Energy, Grant no DE-SC0018269.
Conflict of interest
The authors declare that the research was conducted in the
absence of any commercial or financial relationships that could be
construed as a potential conflict of interest.
Publisher’s note
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authors and do not necessarily represent those of their affiliated
organizations, or those of the publisher, the editors and the
reviewers. Any product that may be evaluated in this article, or
claim that may be made by its manufacturer, is not guaranteed or
endorsed by the publisher.
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Glossary
BTA1 Betaine Lipid Synthase 1
CPT CDP-choline:1 2-Diacylglycerol Cholinephosphotransferase
CrLAT1 C. reinhardtii Lysolipid Acyltransferase 1
DAG Diacylglycerol;
DGAT Di a c y l g lyc e ro l Acy l t ransfer a se
DGDG Diga lactosy l d iacyl g lycerol
DGTA Diacylglycerylhydroxymethyltrimethyl-b-Alanine
DGTS Diacylglyceryl-N
N N-trimethylhomoserine
ER Endoplasmic Reticulum
FA Fatty Acid
FFA Free Fatty Acid
FAD Fatty Acid Desaturase
G3P Glycerol 3-Phosphate
GPAT Glycerol-3-Phosphate Acyltransferase
GPC Glycerolphosphocholine
GPCAT Glycerolphosphocholine Acyltransferase
LC-MS Liquid Chromatography Mass Spectrometry
LPAAT Lysophosphatidic Acid Acyltransferase
LPCAT Lysophosphatidylcholine Acyltransferase
LPCT Lysophosphatidylcholine Transacylase
LPEAT Lysophosphatidylethanolamine Acyltransferase
LPLAT Lysophospholipid Acyltransferase
MGDG Monogalactosyldiacylglycerol
PA Phosphatidic Acid
PC Phosphatidylcholine
PDAT Phospholipid : Diacylglycerol Acyltransferase
PDCT Phosphatidylcholine : Diacylglycerol Cholinephosphotransferase
PE Phosphatidylethanolamine
PG Phosphatidylglycerol
PGD1 Plastid Galactoglycerolipid Degradation 1
PLIP1 Plastid Lipase 1
PUFA Polyunsaturated Fatty Acid
SQDG Sulfoquinovosyldiacylglycerol
TAG Triacylglycerol
TGD Trigalactosyldiacylglycerol
Hoffmann and Shachar-Hill 10.3389/fpls.2023.1077347
Frontiers in Plant Science frontiersin.org14