The Plant Journal (2015) 82, 481–503
doi: 10.1111/tpj.12823
SI CHLAMYDOMONAS
Algae after dark: mechanisms to cope with anoxic/hypoxic
conditions
Wenqiang Yang1,*, Claudia Catalanotti1, Tyler M. Wittkopp1,2, Matthew C. Posewitz3 and Arthur R. Grossman1
Department of Plant Biology, Carnegie Institution for Science, Stanford, CA 94305, USA,
2
Department of Biology, Stanford University, Stanford, CA 94305, USA, and
3
Department of Chemistry and Geochemistry, Colorado School of Mines, Golden, CO 80401, USA
1
Received 18 November 2014; revised 28 February 2015; accepted 3 March 2015; published online 9 March 2015.
*For correspondence (e-mail [email protected])
SUMMARY
Chlamydomonas reinhardtii is a unicellular, soil-dwelling (and aquatic) green alga that has significant
metabolic flexibility for balancing redox equivalents and generating ATP when it experiences hypoxic/
anoxic conditions. The diversity of pathways available to ferment sugars is often revealed in mutants in
which the activities of specific branches of fermentative metabolism have been eliminated; compensatory
pathways that have little activity in parental strains under standard laboratory fermentative conditions are
often activated. The ways in which these pathways are regulated and integrated have not been extensively
explored. In this review, we primarily discuss the intricacies of dark anoxic metabolism in Chlamydomonas,
but also discuss aspects of dark oxic metabolism, the utilization of acetate, and the relatively uncharacterized but critical interactions that link chloroplastic and mitochondrial metabolic networks.
Keywords: Chlamydomonas reinhardtii, dark growth, oxic conditions, anoxic conditions, fermentation,
acetate metabolism.
INTRODUCTION
Chlamydomonas reinhardtii (referred to as Chlamydomonas throughout) is a soil-dwelling photosynthetic organism with certain metabolic features that are similar to those
associated with vascular plants (photosynthesis), and others that were lost during vascular plant evolution (e.g. flagella biogenesis). This alga has been exploited as an
attractive reference system for several decades. As a result
of sequencing of the Chlamydomonas nuclear genome
(Merchant et al., 2007), the development of sophisticated
molecular techniques applicable to this alga (Harris, 2001;
Grossman et al., 2007; Purton, 2007; Gonzalez-Ballester
et al., 2011), and its ability to grow photoautotrophically,
mixotrophically and heterotrophically, Chlamydomonas is
ideal for dissecting a range of biological, cellular, molecular
and physiological processes, including flagella/cilia function and assembly (Dutcher, 1995; Cao et al., 2013), the biogenesis and activity of chloroplasts (Rochaix, 2001;
Duanmu et al., 2013; Heinnickel et al., 2013), acclimation of
cells to changing nutrient conditions (macro- and micronutrients) (Merchant et al., 2006; Moseley et al., 2009; Page
et al., 2009; Gonzalez© 2015 The Authors
The Plant Journal © 2015 John Wiley & Sons Ltd
Ballester et al., 2010; Pootakham et al., 2010; Aksoy et al.,
2013), phototaxis and photoperception (Nagel et al., 2002;
Wagner et al., 2008), the characteristics of the carbon-concentrating mechanism (Fang et al., 2012; Meyer and Griffiths, 2013), and lipid biosynthesis for the potential
production of biofuels (Li et al., 2012; Johnson and Alric,
2013). Moreover, Chlamydomonas synthesizes molecular
hydrogen (H2) when experiencing anoxia, which is likely a
frequent occurrence during the evening in environments
where there is limited aeration and active microbial respiration (Melis and Happe, 2001, 2004; Ghirardi et al., 2009;
Grossman et al., 2011; Catalanotti et al., 2013; Yang et al.,
2014a). Finally, Chlamydomonas is a powerful model for
dissecting aspects of dark, oxic metabolism (Salinas et al.,
2014), for which little information is available.
DARK METABOLISM IN PHOTOSYNTHETIC ORGANISMS
General aspects
Photosynthetic microorganisms generate energy exclusively through dark metabolism for almost half of the day
481
482 Wenqiang Yang et al.
(Perez-Garcia et al., 2011). The availability of O2 during the
dark phase of the diel cycle heavily influences the differential activation of distinct metabolic processes. Many algae
not only have extensive fermentation networks available to
generate ATP when O2 is not available, but are also able to
respire intracellular energy stores (e.g. starch), as well as
assimilate extracellular organic substrates (e.g. acetate and
glucose) for growth/ATP generation when O2 becomes
available. It is only by developing an understanding of the
metabolic circuits associated with dark, oxic and hypoxic
metabolism and their integration over the diel cycle (with
metabolism that dominates in the light) that we will obtain
a comprehensive understanding of net carbon cycling and
the overall energy budgets of photosynthetic organisms in
the environment. Such studies may also provide valuable
information regarding specific roles of enzymes predicted
to be associated with dark metabolism and the diversity of
metabolic networks available to sustain ATP production in
the dark. To appreciate the variety of ways in which carbon
is cycled over the course of the day and the metabolic consequences of this cycling, it is critical to understand fluctuations in aquatic and terrestrial O2 levels, the nature of
catabolism in the dark, how much fixed carbon is directed
toward respiratory and fermentation processes daily, and
the impact of catabolic processes on fixed carbon storage.
Additionally, dark, anoxic metabolism in photosynthetic
microbes has important ecological consequences, as many
algae and cyanobacteria excrete reduced energy carriers
(e.g. organic acids/alcohols and H2) during the night when
the environment becomes hypoxic or anoxic (Mus et al.,
2007; Ananyev et al., 2008; Dubini et al., 2009; Carrieri et al.,
2010). These excreted reducing equivalents and carbon substrates fuel the growth of an often diverse group of co-existing heterotrophic microbes. It is likely that the types and
amounts of products secreted by specific photosynthetic
microorganisms markedly influence the types and densities
of the biota present in a variety of aquatic and soil ecosystems (Hoehler et al., 2002; Spear et al., 2005).
In this review, we present current advances in our understanding of fermentation, and also describe older pioneering studies, that demonstrate the fascinating mechanisms
used by algae, and particularly Chlamydomonas, to function metabolically in the dark. We also briefly discuss
aspects of metabolism in the light, trafficking of reductants
between chloroplasts and mitochondria, and chlororespiration, as this information establishes a metabolic framework
through which to assess dark, oxic and anoxic metabolism.
Mitochondrial mutants defective for heterotrophic growth
Chlamydomonas is capable of growing in the dark under
oxic conditions, while at the same time maintaining photosynthetically competent thylakoid membranes, through
assimilation and metabolism of acetate. Many Chlamydomonas mutants deficient for dark heterotrophic growth
have lesions in genes encoding proteins that function in
mitochondria (see below), but the lesions may also affect
proteins located outside of the mitochondria. Several
commonly used laboratory ‘wild-type’ strains, including
CC-4425 (D66+), cw15 and CC–4619 (dw15), exhibit some
growth impairment in the dark (Table 1); this finding probably reflects lesions that have accumulated during longterm growth of the cultures in continuous light, which may
obscure features of these organisms that have evolved for
fitness in the natural environment.
The mitochondrial electron transport chain (mETC) is the
site of oxidative phosphorylation. It uses reductant generated from glycolysis, the pyruvate dehydrogenase complex
and the tricarboxylic acid (TCA) cycle to establish an electrochemical transmembrane gradient that drives ATP synthesis. Most Chlamydomonas mutants with compromised
mitochondrial function are unable to use acetate as a carbon source for heterotrophic growth. ‘Dark-dying’ mutants
include those that either lack or have defects in specific
components associated with complexes I–IV of the mETC,
or that affect the proper assembly of these complexes.
The Chlamydomonas mitochondrial proteome includes
approximately 350 proteins (Atteia et al., 2009), while the
mitochondrial genome contains only 12 genes, seven of
which encode proteins that function in the mETC (Gray and
Boer, 1988; Michaelis et al., 1990). Therefore, the majority of
proteins contributing to mitochondrial function, including
respiratory activity, are nucleus-encoded and imported into
the organelle by the Transporter Inner Membrane and Transporter Outer Membrane (TIM-TOM) for mitochondria protein
transport complex (Neupert, 1997). A number of Chlamydomonas mutants that are defective for dark growth and are
disrupted for mitochondrial genes have been identified
(known as dum, i.e. dark uniparental minus, indicating nonMendelian inheritance from the mt parent), although most
Chlamydomonas mutants with dark-growth deficiencies
have lesions in nuclear genes that encode mitochondrialocalized proteins that are not associated with a specific,
experimentally determined function (Table 1) (Salinas et al.,
2014).
The first respiratory-deficient Chlamydomonas strains,
which were isolated by Wiseman et al. (1977), were generated by nitrosoguanidine mutagenesis followed by selection for cells unable to grow in the dark. Several of these
nuclear mutants exhibited altered mitochondrial cytochrome c oxidase activity (Wiseman et al., 1977).
Subsequently, many mutants with defects in complex I (dum5, dum17, dum20, dum23, dum25), complex III
(dum1, dum11, dum15, dum22, dum24) or complex IV
(dum18, dum19) of the mitochondrial respiratory system
were identified after treatment of cells with the mutagenic
dyes acriflavine and ethidium bromide (Matagne et al.,
1989; Dorthu et al., 1992; Colin et al., 1995; Duby and
Matagne, 1999; Remacle et al., 2001a,b; Cardol et al., 2002,
© 2015 The Authors
The Plant Journal © 2015 John Wiley & Sons Ltd, The Plant Journal, (2015), 82, 481–503
Dark hypoxic growth of algae 483
Table 1 Mutants that are unable to grow (or grow slowly) under dark oxic conditions, and mutants in genes encoding enzymes that function
under dark anoxic conditions
Mutant name
Protein encoded by mutated gene
Dark growth deficiency
CC–4425 (D66)
Unknown
CC–4619 (dw15)
Unknown
cw15
Unknown
fdx5
Ferredoxin 5
ack1
Acetate kinase 1
dk series mutants Alterations in mitochondrial inner
membranes and deficiencies in
cytochrome oxidase activity
dum series
‘Dark uniparental minus’, mutations
in respiratory complexes I, III and IV
nda1
Type II NAD(P)H dehydrogenase
atp2
CF1 b subunit
icl1
Isocitrate lyase 1
y1
Protochlorophyllide oxidoreductase
Dark anoxia
pfl1
Pyruvate formate lyase 1
amiPFL1
Pyruvate formate lyase 1
pfr1
Pyruvate:ferredoxin oxidoreductase 1
adh1
Alcohol/aldehyde dehydrogenase 1
ack1
Acetate kinase 1
ack2
Acetate kinase 2
pat2
Phosphate acetyltransferase 2
ack1 ack2
Double mutant
pat2 ack2
Double mutant
hydEF
Hydrogenase maturation factor EF
Hydrogenase maturation factor G
hydG
hydA1
Hydrogenase 1
hydA2
Hydrogenase 2
hydA1 hydA2
Double mutant
amiTHB8
2–on–2 hemoglobin
Method of creating mutation
Publication
Unknown
Unknown
Unknown
Random insertional mutagenesis
Random insertional mutagenesis
Nitrosoguanidine mutagenesis
W. Yang, unpublished
W. Yang, unpublished
W. Yang, unpublished
W. Yang, unpublished results
W. Yang, unpublished results
Wiseman et al., 1977
Acriflavine-induced mutagenesis
Reviewed by Salinas et al., 2014
RNAi
RNAi
Random insertional mutagenesis
UV mutagenesis
Lecler et al., 2012
Lapaille et al., 2010
Plancke et al., 2014
Sager, 1955
Random insertional mutagenesis
MicroRNA
TILLING
Random insertional mutagenesis
Random insertional mutagenesis
Random insertional mutagenesis
Random insertional mutagenesis
Cross between ack1 and ack2
Cross between pat2 and ack2
Random insertional mutagenesis
Random insertional mutagenesis
Random insertional mutagenesis
Random insertional mutagenesis
Cross between hydA1 and hydA2
MicroRNA
Philipps et al., 2011; Catalanotti et al., 2012
Burgess et al., 2012
C. Catalanotti, unpublished results
Magneschi et al., 2012
Yang et al., 2014b
Yang et al., 2014b
Yang et al., 2014b
Yang et al., 2014b
Yang et al., 2014b
Posewitz et al., 2004a
M. C. Posewitz, unpublished results
Meuser et al., 2012
Meuser et al., 2012
Meuser et al., 2012
Hemschemeier et al., 2013b
2008). Chlamydomonas is unique among photosynthetic
organisms in that its mitochondrial DNA may be targeted
for site-directed mutagenesis (via homologous recombination) using biolistic transformation (Remacle et al., 2006).
More recent approaches to identify mitochondrial mutants
have exploited random insertional mutagenesis and RNA
interference to knockout or knockdown expression of
nuclear genes important for mitochondrial function
(Table 1) (Cardol et al., 2006; Remacle et al., 2010; Barbieri
et al., 2011; Salinas et al., 2014).
While many mitochondrial mutants are disrupted for
respiratory function and compromised for dark heterotrophic growth, some exhibit less severe phenotypes. Some
mutants defective for mitochondria- and nucleus-encoded
subunits of complex I grow slowly in the dark (Remacle
et al., 2001a) and consume O2, generating a transmembrane proton gradient via a rotenone-resistant type II
NAD(P)H dehydrogenase (NDA1) coupled with electron
transport to the alternative oxidase (Remacle et al.,
2001a). nda1 RNAi lines showed abnormal growth phenotypes when the knockdown lines were grown heterotrophically (Lecler et al., 2012). Some mutants affected in
complex III (cob, encoding apocytochome b) and
complex IV (cox1, encoding subunit 1 of cytochrome oxidase) retained some respiratory activity via the non-phosphorylating alternative (salicylhydroxyamic acid-sensitive)
pathway, which transfers electrons from reduced ubiquinone to O2, but these strains only grew photoautotrophically (Remacle et al., 2001b). Finally, all 17 subunits of
Chlamydomonas complex V (mitochondrial ATP synthase) are nucleus-encoded. Knockdown of ATP2 (encoding the CF1 b subunit) resulted in decreased respiratory
O2 consumption and obligate photoautotrophy as a consequence of the loss of mitochondrial ATP synthesis (Lapaille et al., 2010). Interestingly, a decrease in ATP2 RNA
also affected photosynthetic activity, causing a shift into
state II [mobile light harvesting complex moves off of
photosystem II (PSII)], which may be part of a physiological compensating response that favors cyclic electron
flow (CEF) and increased ATP production in the light by
the photosynthetic electron transport system (Lapaille
et al., 2010) and/or reduces the effect of a loss of mitochondrial respiration as an electron valve that oxidizes
chloroplast-generated reductant when the redox status of
the plastid is increased (e.g. when there is excess excitation, such as under high light conditions).
© 2015 The Authors
The Plant Journal © 2015 John Wiley & Sons Ltd, The Plant Journal, (2015), 82, 481–503
Dark hypoxic growth of algae 497
ditions. There are several other energetic and redox considerations that distinguish light from dark growth in photosynthetic organisms. NADPH plays a critical role in
driving anabolic processes, including the synthesis of lipids, amino acids and nucleotides, and is directly produced
by the activity of FDX:NADP+ oxidoreductase. During dark
metabolism, many reactions including those of the TCA
cycle and glycolysis generate NADH; the oxidative pentose
phosphate pathway produces NADPH. Interconversion
between NADH and NADPH may be achieved by the pyridine nucleotide transhydrogenase (Agledal et al., 2010;
Holm et al., 2010). This enzyme regulates the NAD(H)/
NADP(H) ratio through a reversible hydride transfer that
occurs in either an energy-dependent or energy-independent manner (Olausson et al., 1992; Pedersen et al., 2008);
the NAD(H)/NADP(H) ratio helps to control the extent to
which the cells perform catabolic and anabolic processes.
Some bacteria, including E. coli, rely heavily on pyridine
nucleotide transhydrogenase activity to modulate metabolism (Sauer et al., 2004; Fuhrer and Sauer, 2009).
Other electron carriers, such as the FDXs and thioredoxins, are small redox carriers that supply electrons to a range
of cellular processes, as previously discussed. Furthermore,
while some of the FDX proteins may be efficiently reduced
by NADH or NADPH, FDXs with a very negative redox
potential may only be able to acquire electrons through PSI,
which suggests tailoring of redox components in the light
and the dark. As mentioned above, we isolated a mutant of
Chlamydomonas that does not grow in the dark (but does
grow in the light) and is null for FDX5 (W. Yang, unpublished results). This result supports the concept that the
FDX family in Chlamydomonas represents a group of proteins with a specialized function as electron carriers, but
their functions may only be possible in the light (when PSI
through FDX1 supplies much of the reductant) or dark
(where NADH supplies most of the reductant). More information is required with respect to the redox potential of the
various FDXs and the affinities with which they interact with
their specific target proteins. Other redox carriers such as
thioredoxins may also be critical for ‘dark’ metabolism.
PERSPECTIVES
Chlamydomonas is a metabolically versatile organism that
performs photosynthetic CO2 fixation, aerobic respiration
and anaerobic fermentation. This alga is a model for
examining many aspects of photosynthetic metabolism,
and has been the subject of numerous metabolic studies.
Many pathways and enzymes associated with fermentation metabolism in this organism are only now being
defined, and almost nothing is known about mechanisms
by which these pathways are regulated, or the ways in
which fermentation products are partitioned among the
various cell compartments. The generation of lesions that
block some of these pathways is providing new insights
into compensatory responses that allow sustained ATP
production while eliminating reducing equivalents
through generation of reduced carbon compounds that
are excreted from cells. Initial characterizations of Chlamydomonas have demonstrated that this alga has flexible, mixed-acid fermentation pathways, with features
common to bacterial-, plant- and yeast-type fermentation.
Most enzymes for fermentative metabolism in the algae,
inferred from genomic and metabolic studies, have not
been biochemically characterized. Expression patterns of
genes encoding these enzymes, the biochemical properties of these enzymes (including potential interactions
with each other), and the diversity of fermentation pathways plus the extent to which they are used under various conditions, require further examination in a broader
spectrum of algal systems. Additionally, the diversity of
external and internal end-products accumulated by various algae during fermentation is still mostly unknown.
Such information is critical for developing a clear understanding of metabolic diversity both within and among
the various algal groups, and the ways in which fermentation pathways have been shaped by environmental conditions. Furthermore, there are many technologies,
including flux balance analysis, mass flux analysis, timeresolved fluorescence measurements and the use of O2
microsensors that may help to evaluate the redox conditions of cells and correlate those conditions with the activities of both oxic and anoxic metabolisms. An
understanding of the various pathways critical for dark
metabolism and the ways in which these pathways are
controlled constitutes a domain of metabolism that must
be fully described if we are to understand the energy budget of photosynthetic microbes in the environment and
potential ways to manipulate carbon cycling. Finally, fermentation metabolism in algae appears to represent a significant ecological component of carbon flux in soils (and
sediments) that strongly affects its content of organic
acids, alcohols and H2, which in turn affects the biotic
composition of the ecosystem.
ACKNOWLEDGEMENTS
The work performed in our laboratories and described here was
supported by grants from the US the Department of Energy (numbers DE-FG02-12ER16338 and DE-FG02-12ER16339). Aspects of the
work were also funded by US National Science Foundation grants
to A.R.G. (MCB0824469 and MCB-0951094).
REFERENCES
Agledal, L., Niere, M. and Ziegler, M. (2010) The phosphate makes a difference: cellular functions of NADP+. Redox Rep. 15, 2–10.
Aksoy, M., Pootakham, W., Pollock, S.V., Moseley, J.L., Gonzalez-Ballester,
D. and Grossman, A.R. (2013) Tiered regulation of sulfur deprivation
responses in Chlamydomonas reinhardtii and identification of an associated regulatory factor. Plant Physiol. 162, 195–211.
Allen, F.L. and Horwitz, L. (1957) Oxygen evolution and photoreduction in
adapted Scenedesmus. Arch. Biochem. Biophys. 66, 45–63.
© 2015 The Authors
The Plant Journal © 2015 John Wiley & Sons Ltd, The Plant Journal, (2015), 82, 481–503
Dark hypoxic growth of algae 485
produced by photophosphorylation. Under such conditions, ATP may be acquired from mitochondria by trafficking electrons out of chloroplasts through the action of the
dihydroxyacetone 3–phosphate (DHAP)/3–phosphoglycer€ mer, 1995; Boschetti and Schmid,
ate (3–PGA) shuttle (Kro
1998). DHAP export coupled with its oxidation to 3–PGA
generates NADH (or NADPH) for respiratory energy production. The rate of shuttling depends on the rate at which
chloroplastic 3–PGA is reduced and cytosolic DHAP is oxidized (Heineke et al., 1991).
The malate/oxaloacetate (OAA) shuttle is also central to
inter-organelle communication. Chloroplast NADP+-dependent malate dehydrogenase (MDH) is activated in the light,
and, like the DHAP/3–PGA shuttle, the malate/OAA shuttle
helps to adjust the cellular NADPH/NADP+ ratio and coordinate the availability of reducing equivalents with the synthesis of ATP (Anderson and House, 1979; Scheibe, 1987;
Weber et al., 1995). When the redox state of chloroplasts is
increased (high NADPH/NADP+), MDH uses NADPH to
reduce OAA to malate, which is then transported from chloroplasts to mitochondria where it is converted back to OAA
and NADH; the latter may then be re-oxidized through respiratory activity (Scheibe, 1987). This shuttle also affects the
NADPH/ATP ratio, which may be important for optimizing
carbon fixation in the light (Scheibe, 1987). While a malate/
OAA transporter on the chloroplastic envelope of Chlamydomonas has not been identified, the low CO2-inducible
protein LCI20 is a candidate for this function (Terashima
et al., 2011; Johnson and Alric, 2013). The malate/OAA shuttle may also potentially work in the opposite direction,
transporting reducing equivalents from mitochondria to
chloroplasts when the stromal NADPH/NADP+ ratio is low,
or under conditions in which mitochondria are overreduced. Hence, this shuttle has an inter-organellar function
in the management of cellular reductant/energy demands in
both the light and the dark.
In the dark, mitochondrial respiration supplies most of
the ATP for cell growth, which has also been linked to interactions between mitochondria and chloroplasts. For example, when oxidative phosphorylation is inhibited in
mitochondria in the dark, the chloroplastic plastoquinone
(PQ) pool becomes reduced, and the photosynthetic apparatus transitions from state I (mobile antennae on photosystem II) to state II (mobile antennae on photosystem I)
(Gans and Rebeille, 1990) by a mechanism that involves
phosphorylation of the mobile light-harvesting antenna
(Rochaix, 2007). Mitochondrial export of ATP and reductants in the dark may also prime the chloroplast for efficient
photosynthetic activity upon the onset of light by maintaining a proton gradient across the thylakoid membranes (Joliot and Joliot, 1980); this transmembrane gradient is
thought to be sustained by ATP imported from mitochondria and hydrolyzed in chloroplasts via the ‘reverse’ activity
of the thylakoid ATP synthase (Joliot and Joliot, 1980). A
dark proton gradient across thylakoid membranes may also
function in controlling non-photochemical quenching upon
the onset of light by modulating dark accumulation of xanthophyll cycle constituents (e.g. zeaxanthin, antheraxanthin
€ rkman, 1995; Hoefnagel
and violaxanthin) (Gilmore and Bjo
and Wiskich, 1998). Finally, efficient photosynthetic electron
flow after transfer of cells from the dark to the light requires
availability of electrons for PSI reduction (e.g. a partially
reduced PQ pool) following rapid photo-oxidation of the
PSI reaction center chlorophyll special pair (P700 to P700+).
The occurrence of a partially reduced PQ pool in the dark is
attributed to the transfer of reducing equivalents from mitochondria to chloroplasts, but also the degradation of chloroplastic starch reserves (Bulte et al., 1990; Wieckowski and
Wojtczak, 1997). The reduction state of the PQ pool also
involves chlororespiration, a process that requires NAD(P)H
dehydrogenase (NDA2 in Chlamydomonas), which reduces
PQ, and plastid terminal oxidase 2, which regenerates oxidized PQ through reduction of O2; this occurs in Chlamydomonas and other photosynthetic organisms (Bennoun,
2002; Peltier and Cournac, 2002; Bailey-Serres and Voesenek, 2008; Jans et al., 2008; Houille-Vernes et al., 2011).
Although it was once thought that chlororespiration was
involved in dark ATP production in chloroplasts, there is
now evidence suggesting that chlororespiration may not be
electrogenic (i.e. electron transfer from NAD(P)H to O2 is
not coupled to the translocation of protons across thylakoid
membranes) (Cournac et al., 2000; Johnson and Alric,
2013).
Oxyhydrogen reaction
Although hydrogenases are sensitive to O2 (Ghirardi et al.,
1997), these enzymes are capable of extracting electrons
from H2 in the dark when CO2 is also present in the surrounding atmosphere, under hypoxic conditions (e.g.
approximately 1% O2); this reaction, called the oxyhydrogen reaction (Figure 1) may be observed in both whole
algal cells (Gaffron, 1939, 1940, 1942a,b; Russell and Gibbs,
1968; Maione and Gibbs, 1986) and isolated chloroplasts
(Chen and Gibbs, 1992). It has also been observed in the
cyanobacterium Anabaena sp. 7120 (Frenkel et al., 1949;
Houchins and Burris, 1981a,b).
The oxyhydrogen reaction couples the uptake of H2 and
O2 with the fixation of CO2 through the Calvin–Benson
cycle/reductive pentose phosphate pathway (Gaffron, 1940;
Badin and Calvin, 1950; Gingras et al., 1963; Russell and
Gibbs, 1968). While it is likely that a significant amount of
the O2 uptake associated with the oxyhydrogen reaction is
the result of mitochondrial respiration (Allen and Horwitz,
1957; Horwitz, 1957; Erbes and Gibbs, 1981; Chen and
Gibbs, 1992), other electron transfer processes may also
supply the ATP required for CO2 fixation (Gaffron, 1942a,b;
Maione and Gibbs, 1986). These studies highlight that the
overall redox conditions of the environment may change
© 2015 The Authors
The Plant Journal © 2015 John Wiley & Sons Ltd, The Plant Journal, (2015), 82, 481–503
486 Wenqiang Yang et al.
CO2
4
ATP
NADH
NADPH
e–
Calvin-benson
cycle
3
FNR
NDH
1
Hydrogenase
2
mETC
6
FDX
7
H+
8
e–
CO2
Stroma
PQ
NADH
PSII
e–
TCA
cycle
Cyt b6f
?
O2
PFR1
Thylakoid
e–
5
H2
H2O
+
H+ H
H+
PSI
Pyruvate
Lumen
9
Acetate
ATP
Acetyl-CoA
Glyoxylate
cycle
Gluconeogenesis
Figure 1. Possible electron transport during the oxyhydrogen reaction. The cells are grown with a gas mixture that contains oxygen (< 2%), carbon dioxide and
hydrogen (approximately 3%). Lines and circles in various colors represent possible pathways occurring in the oxyhydrogen reactions. (1) Hydrogen may be oxidized by hydrogenases, and the electrons from H2 ultimately used to reduce O2 or CO2. Hydrogenases directly reduce ferredoxin (gray). (2) Reduced ferredoxin
serves as substrate for (FNR), which forms NADPH (pink). (3) NADPH may be used for carbon fixation (light blue). (4) NADPH may be converted to NADH and
used in the mETC to produce ATP (green). (5) The TCA cycle produces NADH, which is also used by the mETC (dark blue). (6) Electrons from NADPH can also
reduce the quinone pool, and this electron may be accepted by O2 or other unknown components in the lumen (purple). (7) Electrons can also be donated by
FDX to the cytochrome b6f complex (orange). Coupled to proton transfer, this process may facilitate generation of a pH gradient. (8) FDX may reduce PFR1 to
form pyruvate, which could be directed to gluconeogenesis (red). (9) Acetate can be converted to acetyl CoA for the glyoxylate cycle to produce organic compounds for gluconeogenesis. NDH, NAD(P)H dehydrogenase.
dramatically over the course of the day. These changes,
plus the availability of various substrates such as H2 and
O2, and light intensities, may trigger alternative electron
flow involving both mitochondrial and chloroplastic electron carriers, although many of these pathways are still
very poorly defined. The use of alternative pathways for
the generation of energy/reductant also highlights the metabolic flexibility of at least some soil-inhabiting algae,
organisms that must rapidly respond to dramatic changes
in redox conditions with changing temperature, light, O2
and nutrient levels.
DISRUPTION OF CHLAMYDOMONAS FERMENTATION
PATHWAYS
Anoxia and fermentation
The energetics of an ecosystem may be markedly affected
by O2 levels, which continually fluctuate over the course of
a day. Algal cells experiencing hypoxic/anoxic conditions
typically generate energy by substrate-level phosphorylation, which requires glycolytic catabolism of fixed carbon
(polysaccharides/sugars). If O2 cannot be used as a terminal electron acceptor to re-oxidize the NADH generated by
the glycolytic degradation of carbohydrates and TCA cycle
activity, the cells decrease the rate of glycolytic metabolism, causing decreased rates of ATP production. The
NADH that accumulates during anoxic glycolysis must then
be recycled by alternative mechanisms, which, in the case
of Chlamydomonas, typically involves metabolizing pyruvate to a variety of reduced, fermentative end-products
that are secreted from cells, e.g. formate, acetate, lactate,
succinate, glycerol, ethanol and H2 (Gfeller and Gibbs,
1984, 1985; Mus et al., 2007; Dubini et al., 2009; Philipps
et al., 2011; Catalanotti et al., 2012, 2013; Magneschi et al.,
2012; Yang et al., 2014a).
In both soil and aquatic environments, Chlamydomonas
may experience hypoxic/anoxic conditions, especially at
night when photosynthetic O2 evolution ceases and environmental O2 levels dramatically decrease because of
respiratory activity. Fermentation metabolism may also
occur in the light when cells experience anoxic conditions
(i.e. when respiratory consumption of O2 by the microbial
community exceeds photosynthetic O2 production). Chlamydomonas may rapidly acclimatize to anoxia (Gfeller and
Gibbs, 1984, 1985; Kreuzberg, 1984; Gibbs et al., 1986;
Ohta et al., 1987) by activating a variety of metabolic/fermentation pathways (Tsygankov et al., 2002; Hemschemeier and Happe, 2005; Atteia et al., 2006; Mus et al., 2007;
Dubini et al., 2009; Timmins et al., 2009) and regulating the
expression of genes encoding activities integral to those
pathways (Grossman et al., 2007; Merchant et al., 2007).
Furthermore, aspects of Chlamydomonas fermentation
© 2015 The Authors
The Plant Journal © 2015 John Wiley & Sons Ltd, The Plant Journal, (2015), 82, 481–503
Dark hypoxic growth of algae 487
tion of H2 and CO2 (Gfeller and Gibbs, 1984; Kreuzberg,
1984; Ohta et al., 1987; Mus et al., 2007; Dubini et al., 2009;
Catalanotti et al., 2012; Magneschi et al., 2012). In the environment, these released fermentation products probably
supply heterotrophic microbes with organic compounds
and reductants for growth and development.
Two major pyruvate-metabolizing enzymes of Chlamydomonas include the pyruvate formate lyase PFL1 and
the pyruvate:ferredoxin oxidoreductase PFR1 (the latter is
sometimes designated PFOR). PFL1 was localized to both
mitochondria and chloroplasts based on measurements of
activity, proteomic data and immunological analyses
(Kreuzberg et al., 1987; Atteia et al., 2006; Terashima et al.,
2010), whereas PFR1 was localized exclusively to chloroplasts (Terashima et al., 2010; van Lis et al., 2013). The PFL1
reaction catalyzes the conversion of pyruvate to acetyl CoA
and formate, and this appears to be the dominant enzyme
in pyruvate metabolism after Chlamydomonas acclimatizes
to dark anoxia (Mus et al., 2007; Philipps et al., 2011; Catalanotti et al., 2012), while in the PFR1 reaction, pyruvate is
converted to acetyl CoA, CO2 and reduced FDX. PFR1 was
metabolism appear to be highly flexible based on physiological/metabolic studies using wild-type and mutant
strains (Gfeller and Gibbs, 1984, 1985; Kreuzberg, 1984;
Gibbs et al., 1986; Ohta et al., 1987; Hemschemeier and
Happe, 2005; Atteia et al., 2006; Mus et al., 2007; Dubini
et al., 2009; Timmins et al., 2009; Grossman et al., 2011;
Philipps et al., 2011; Burgess et al., 2012; Catalanotti et al.,
2012, 2013; Magneschi et al., 2012; Meuser et al., 2012;
Yang et al., 2014b), global examination of gene expression
as cells acclimatize to anoxic conditions (Mus et al., 2007;
Hemschemeier et al., 2013a), and analysis of the Chlamydomonas genome through homology searches (Grossman et al., 2007, 2011; Merchant et al., 2007).
During dark fermentation, cellular carbohydrate reserves
are metabolized through glycolysis to generate ATP; the
NADH that is co-produced must be re-oxidized to sustain
energy production through the glycolytic breakdown of
sugars. Pyruvate, the end-product of glycolysis, is a substrate for many Chlamydomonas fermentation pathways
(Figure 2). The activities of these circuits are reflected by
the secretion of organic acids and alcohols, and the evolu-
Starch
Feedback regulation of glycolysis
Glycolysis
ATP
Pi
NAD + NADH + H+
ADP
GAP
DHAP
PEPC
Phosphoenol
pyruvate
ADP
Oxaloacetate
CO 2
ATP
NADH + H+
ADP
ADP
NAD +
CO 2
PYK
NAD(P)H + H+
ATP
PYC
ATP
Glycerol
MDH
NAD(P)+
α-ketoglutarate Glutamate
CO 2
Alanine
Pyruvate
ALAAT
NADH + H+
LDH PDC3
Acetaldehyde
NADH + H+
ADH1
Ethanol
NADH + H+
FDXred
PFR1red
FDXox
CO2
FUM
Fumarate
2H+
NADH + H+
HYDA1/2
FMR
H2
NAD+
Succinate
HYDEF/HYDG
Pi
2NADH + 2H+
PAT2/PAT1
3-hydroxybutyrate
CoASH
2NAD+
NAD+
PFR1ox
Acetyl-CoA
ADH1
Malate
NAD(P)+
PFR1
Formate
NAD+
ALDH
PFL1
CO2
Lactate
MME4
CoASH
NAD+
NAD(P)H + H+
Acetyl-P
ADP
CoASH
ACK1/ACK2
ATP
Acetate
Figure 2. Fermentative metabolism. Glycolysis (highlighted with a blue background and white lines) degrades photosynthetic hexoses (often from starch) to
pyruvate. In wild-type cells, under anoxic conditions, pyruvate can be used as a substrate by several enzymes, including PFL1 and PFR1 to form acetyl CoA,
which is the substrate for an acetate-producing pathway catalyzed by PAT1/2 and ACK1/2, highlighted with an orange background, or the ethanol-producing
pathway catalyzed by ADH1. Pyruvate can also be used as a substrate to produce ethanol via the PDC3/ADH1 pathway, in which acetaldehyde serves as an intermediate. PFR1 is an oxidoreductase that can reduce FDX during the conversion of pyruvate to CO2 and acetyl CoA. This reduced FDX can be used by HYDA1
and HYDA2 to generate H2. The compounds highlighted with a yellow background represent the major external metabolites excreted by anoxic wild-type cells,
while the compounds highlighted with a green background represent the metabolites that accumulate (both externally and internally) in various mutant strains
under anoxic conditions. The colors used to represent the enzymes indicate the subcellular localizations of the various proteins: purple, dual localization in chloroplasts and mitochondria; blue, chloroplast; red, mitochondria; black, cytosol or unknown. ALAAT, alanine aminotransferase; FMR, fumarate reductase; FUM,
fumarase; HYDEF, hydrogenase assembly factor EF; HYDG, hydrogenase assembly factor G; LDH, lactate dehydrogenase; MME4, malic enzyme 4; PEPC, phosphoenolpyruvate carboxylase; PYC, pyruvate carboxylase; PYK, pyruvate kinase. The black lines and arrows represent pathways occurring in wild-type cells,
while the red lines and arrows represent pathways occurring in various mutants. The pink line represents possible feedback regulation of the acetate-producing
pathway on glycolysis.
© 2015 The Authors
The Plant Journal © 2015 John Wiley & Sons Ltd, The Plant Journal, (2015), 82, 481–503
488 Wenqiang Yang et al.
shown to efficiently interact with both FDX1 and FDX2 (low
micromolar Km values) (Noth et al., 2013), but not the other
Chlamydomonas FDXs (ferredoxins) (van Lis et al., 2013).
Pyruvate oxidation by PFR1 is coupled to the generation of
two molecules of reduced FDX, which may be used by the
hydrogenases HYDA1 and HYDA2 (Mus et al., 2007; Dubini
et al., 2009; Meuser et al., 2012; Noth et al., 2013) to catalyze H2 production. This pathway has been reconstructed
in vitro using biochemically purified constituents (Chlamydomonas HYDA1, FDX1 and PFR1), with robust H2 production being observed in the presence of pyruvate.
Intriguingly, these in vitro reconstitution experiments demonstrated that PFR1 also oxidizes oxaloacetate (Noth et al.,
2013), which, if relevant in vivo, would have profound
implications regarding the ability of amino acid and lipid
catabolic pathways (and acetate assimilation to C4) to feed
into H2 production via PFR1 reduction of FDX. Reduced
FDX may be re-oxidized by several redox enzymes in addition to hydrogenases, including nitrite and sulfate/sulfite
reductases. PFL1 and PFR1 activities appear to occur simultaneously, with both enzymes acting on the same substrate (Mus et al., 2007; Atteia et al., 2013; van Lis et al.,
2013; Noth et al., 2013). This finding suggests the potential
for re-routing fermentative electron flow in Chlamydomonas toward PFR1-dependent production of H2 (potentially a sustainable, clean fuel). Such a possibility has been
tested by disrupting specific fermentation pathways (e.g.
eliminating PFL1) to potentially boost the rate of H2 generation (see below). The acetyl CoA produced as a consequence of PFL1 and PFR1 activities (Figure 2) is either
reduced to ethanol by the alcohol/acetaldehyde dehydrogenase ADH1 (Hemschemeier and Happe, 2005; Atteia
et al., 2006; Dubini et al., 2009) or metabolized to acetate
by the phosphate acetyltransferase (PAT) and acetate
kinase (ACK) reactions (Atteia et al., 2006; Yang et al.,
2014a,b); these latter reactions occur in both Chlamydomonas mitochondria (PAT1 and ACK2) and chloroplasts
(PAT2 and ACK1) (Mus et al., 2007; Grossman et al., 2011;
Catalanotti et al., 2013; Yang et al., 2014a,b). An alternative
pathway for ethanol production may involve direct decarboxylation of pyruvate to CO2 and acetaldehyde through
the action of pyruvate decarboxylase (PDC3). The acetaldehyde generated in this reaction may be reduced to ethanol
by alcohol dehydrogenase activity, with a recent study suggesting that Chlamydomonas ADH1 is able to generate ethanol from both acetyl CoA and acetaldehyde (Magneschi
et al., 2012). While the ADH1 reaction using acetyl CoA as
a substrate oxidizes two NADH molecules, only a single
NADH is oxidized in the reaction using acetaldehyde.
Mutants affected in fermentation metabolism
The hydEF mutant. The hydEF–1 mutant has been characterized in some detail over the last 10 years (Posewitz
et al., 2004a; Dubini et al., 2009). This mutant has no
hydrogenase activity, and consequently does not produce
H2 under anoxic conditions (Posewitz et al., 2004a).
Analyses of metabolites synthesized by this mutant
under anoxic conditions revealed lower levels of CO2,
extracellular formate, acetate and ethanol relative to wildtype cells, but increased carboxylation of pyruvate to
generate extracellular succinate, which sustains the recycling of NADH (Dubini et al., 2009). Transcript and metabolite analyses both strongly suggest that carboxylation of
pyruvate in the hydEF–1 mutant leads to generation of
either malate or OAA, which is subsequently converted
to succinate by reverse TCA cycle reactions; the succinate
is excreted from the cells (Figure 2) (Dubini et al., 2009).
In addition, by studying the hydEF–1 mutant, hydrogenase function was shown to be important for facilitating
photosynthetic processes under anoxic conditions (Ghysels et al., 2013).
pfl1 mutants. Several independent pfl1 mutants have
been isolated and analyzed (Philipps et al., 2011; Burgess
et al., 2012; Catalanotti et al., 2012). Under dark anoxic conditions, the mutants exhibited increases in pyruvate decarboxylation and accumulation of extracellular ethanol and
lactate, as well as increased intracellular levels of alanine,
succinate, malate and fumarate relative to wild-type cells
(Figure 2) (Philipps et al., 2011; Catalanotti et al., 2012).
Dark H2 production in the pfl-1 mutant isolated by Philips
et al. (2011) was either similar to or somewhat higher than
the level observed in wild-type cells, while the pfl1 mutants
characterized under the conditions used by Catalanotti et al.
(2012) exhibited lower H2 accumulation and in vitro activity
than wild-type cells; these differences may be a consequence of differences in the parental strains used to generate the mutants or in the assay/induction conditions for
dark anoxic H2 production. Interestingly, increased amounts
of 3–hydroxybutyrate were excreted into the medium in
pfl1–KD1 and pfl1–KD2 knockdown lines, suggesting the
build-up of acetyl CoA, which, as suggested by the authors,
may be the consequence of increased b-oxidation of fatty
acids or inhibition of the TCA cycle and/or the glyoxylate
shunt (Burgess et al., 2012).
The adh1 mutant. The Chlamydomonas alcohol/acetaldehyde dehydrogenase ADH1 is highly similar to the Escherichia coli AdhE enzyme. Immunoblot analyses showed
similar levels of pyruvate formate lyase, acetate kinase and
hydrogenase in wild-type cells and the adh1 mutant, and,
although the mutant appeared to express more PFR1, there
was no increase in H2 production. Furthermore, although
the adh1 mutant was unable to synthesize any ethanol or
CO2, it accumulated lower levels of formate and higher levels of acetate, lactate and especially glycerol relative to
wild-type cells, allowing effective re-oxidation of NADH
(Figure 2) (Magneschi et al., 2012).
© 2015 The Authors
The Plant Journal © 2015 John Wiley & Sons Ltd, The Plant Journal, (2015), 82, 481–503
Dark hypoxic growth of algae 489
sta mutants. Hydrogenase activity was reduced in two
Chlamydomonas mutants that are unable to accumulate
starch, sta6 (Zabawinski et al., 2001; Chochois et al., 2009)
and sta7 (Posewitz et al., 2004b), under dark anaerobic
conditions, and HYDA1 and HYDA2 transcript levels were
decreased in these strains (Posewitz et al., 2004b). This
indicates that signals other than simply the lack of O2
(potentially cellular redox status) are involved in activating
HYDA transcription. In contrast, under conditions of sulfur
starvation in the light, the sta6 mutant has hydrogenase
activity similar to that of wild-type cells (Chochois et al.,
2009). In addition, analysis of the sta6 mutant showed that
starch breakdown contributes to H2 production via donation of electrons to the PQ pool, and contribution of electrons from the oxidation of H2O by photosystem II also
occurs (Chochois et al., 2009).
The stm6 mutant. Disruption of the gene encoding a
homolog of the human mitochondrial transcription termination factor state transition mutant (STM6) in Chlamydomonas led to various phenotypes including inhibition of
CEF under anaerobic conditions (eliminating competition
between the hydrogenase and PSI-dependent CEF),
increased starch accumulation (providing additional reductant for PQ reduction), a decrease in the number of active
photosystem II reaction centers, an increased rate of respiration, and an elevated rate of sustained H2 photoproduction during sulfur deprivation relative to wild-type cells
(Kruse et al., 2005a,b; Rupprecht, 2009). Additional genetic
modifications generated in the stm6 genetic background
have also been examined. When this mutant is transformed with a gene encoding a glucose transporter, the
resulting stm6 Glc4 strain exhibits increased glucose
uptake and improved H2 photoproduction (Doebbe et al.,
2007). Furthermore, the stm6 Glc4 strain, which has a
reduced antenna size, exhibits an additional increase in the
level of H2 production (Doebbe et al., 2010). This mutant
has a complex pleiotropic phenotype that may be the
result of multiple primary and secondary defects.
The 2–on–2 hemoglobin mutant. Twelve hemoglobin homologs are encoded on the Chlamydomonas genome. A
2–on–2 hemoglobin, designated THB8, was shown to be
required for normal hypoxic growth and expression of
genes controlled by anoxia (Hemschemeier et al., 2013b).
This mutant is discussed further below with respect to O2
sensing.
pat/ack mutants. The PAT/ACK pathway promotes cellular
fitness during dark, anoxic acclimation, coupling the production of acetate to ATP synthesis (Atteia et al., 2006;
Yang et al., 2014b). Characterization of ack and pat
mutants (three single mutants and two double mutants) in
Chlamydomonas showed that the PAT/ACK pathway in
chloroplasts contributes more than that in mitochondria to
the health of cells experiencing hypoxic/anoxic conditions.
In these mutants, the block in acetate metabolism appears
to occur too far down the central metabolic pathway to
readily allow re-direction of metabolites to other pathways,
while the inability to sustain acetate and ATP production
slows down glycolytic metabolism (Figures 2 and 3) (Yang
et al., 2014b). Furthermore, acetate may be synthesized
under anoxic conditions even when both the chloroplastic
and mitochondrial PAT/ACK pathways are disrupted, suggesting that the cells have other metabolic routes for generating acetate, as discussed below.
ACETATE METABOLISM/FERMENTATION
General aspects
Acetate may be used as the sole energy source for growth
of Chlamydomonas when O2 is used as the terminal electron acceptor. Upon uptake (Figure 3), acetate is converted
to acetyl CoA via one of two pathways, both of which consume ATP. One pathway involves direct conversion of acetate to acetyl CoA by acetyl CoA synthetase (ACS), while
the second requires a two-step reaction catalyzed by ACK
and PAT, acting in the reverse direction to that of acetate
production during fermentation. Acetyl CoA then enters
the metabolic networks of the cell through the glyoxylate
cycle, combining with glyoxylate (to form malate) or with
OAA (to form citrate); the output for one ‘turn’ of the cycle
is a molecule of succinate.
Under anoxic/hypoxic conditions, photophosphorylation
appears to be necessary for sustained acetate assimilation
(Wiessner, 1965; Gibbs et al., 1986). The presence of acetate also helps to maintain cells in an anoxic state in the
light and under certain conditions of nutrient deprivation
because it promotes rapid catabolic consumption of O2
(Kosourov and Seibert, 2009; Morsy, 2011). This has been
demonstrated for sta mutants, in which acetate putatively
supports high respiratory rates, which show more rapid
anaerobiosis and H2 generation (Chochois et al., 2009); a
similar result was obtained for immobilized wild-type cells
(Kosourov and Seibert, 2009). Acetate is also a building
block used for storage of reduced carbon in the form of
triacylglycerides (Johnson and Alric, 2013). Finally, when
Chlamydomonas experiences anoxic/hypoxic conditions, it
produces acetate (which is excreted) through the PAT/ACK
pathway. This pathway recycles CoASH from acetyl CoA,
and, at the same time, generates ATP (one molecule per
acetate generated), which contributes to cell maintenance
(Mus et al., 2007; Tielens et al., 2010; Atteia et al., 2013;
Yang et al., 2014a,b).
Chlamydomonas has two parallel PAT/ACK pathways
involving four proteins: PAT1, PAT2, ACK1 and ACK2. Several reports have shown that PAT1/ACK2 are localized to
mitochondria while PAT2/ACK1 are located in chloroplasts
© 2015 The Authors
The Plant Journal © 2015 John Wiley & Sons Ltd, The Plant Journal, (2015), 82, 481–503
490 Wenqiang Yang et al.
Acetate
Cytosol
Acetate
YaaH
AMT
AMT
Acetate
Acetate
ATP
?
Acetylphosphate
AMP
AMP-ACS
CoASH
CoASH
Pi
O2
AcetylAMP
Acetyl-CoA
AMP
Pi
–O2
PPi
AMP-ACS
CoASH
Acetylphosphate
ATP
Acetate
AcetylAMP
AMP-ACS
PAT
CoASH
Acetate
PPi
AMP-ACS
ACK
Acetyl-CoA
Acetate
ATP
Acetate
ADP
Acetate
PAT
Acetate
ASCT
/SCL
ACT1
CGLD2
ACK
YaaH
ADP
ALDH
ATP
?
Acetate
NADH
Acetate
Acetaldehyde
NAD+
Mito or chloro
Acetate
Figure 3. Acetate metabolism under dark oxic and dark anoxic conditions. Various potential routes for acetate metabolism in Chlamydomonas are presented.
The outer black rectangle represents the plasma membrane, while the inner yellow rectangle represents the chloroplastic or mitochondrial membranes; acetate
metabolism occurs within these organelles. The pink line is used to separate oxic (top) and anoxic (bottom) conditions inside mitochondria and chloroplasts.
Double bars in various colors represent putative acetate transporters (the different colors were used to indicate that there may be different transporter types)
localized on the plasma membrane, chloroplastic and mitochondrial membranes. The enzymes shown in red are encoded by high-confidence gene models present in the Chlamydomonas genome, while the enzymes shown in gray represent gene models for which the function is not absolutely clear. Solid lines represent confirmed Chlamydomonas reactions, while dashed lines indicate proposed reactions based on gene model analyses and homology searches using
Phytozome 9.0 (http://www.phytozome.net/). ACT1, acyl CoA thioesterase; CGLD2, acyl CoA thioesterase; AMT, ammonium transporters; YaaH, members of the
GPR1/FUN34/YaaH family (putative acetate transporters).
(Atteia et al., 2006, 2009; Terashima et al., 2010; Yang
et al., 2014b). PAT/ACK activities of Chlamydomonas typically constitute the dominant pathways for acetate synthesis under dark anoxic conditions. The activities of these
pathways and the accumulation of acetate in cells experiencing anoxia may be affected by altering various
branches of fermentation metabolism through generation
of mutants. For example, accumulation of extracellular acetate during anoxia was diminished in pfl1 mutants (Burgess et al., 2012; Catalanotti et al., 2012) and by treating
anoxic cultures with a PFL inhibitor (Philipps et al., 2011).
However, acetate production increased in the adh1 single
and pfl1–1 adh1 double mutants (Catalanotti et al., 2012;
Magneschi et al., 2012). In the hydEF–1 mutant, acetate
production was reduced to half of that of wild-type cells as
much of the pyruvate was no longer converted to acetyl CoA but was carboxylated and then reduced to succinate by reverse TCA reactions (Dubini et al., 2009).
Fermentative pathways in ack and pat mutants
Insertional mutants of Chlamydomonas disrupted for
genes encoding the chloroplastic and mitochondrial acetate kinases ACK1 and ACK2 and the chloroplastic phosphate acetyltransferase PAT2 were recently isolated and
characterized (Yang et al., 2014b), revealing that fermentative acetate metabolism in Chlamydomonas was more
complicated than expected. The ack1 and pat2 strains
exhibited a more pronounced decrease in acetate secretion
under dark anoxic conditions compared with the ack2
strain (Yang et al., 2014b), suggesting a dominant role for
the chloroplast in acetate production. Among the chloroplastic enzyme mutants, the pat2–1 mutant consistently
produced less acetate than the ack1 mutant, as expected if
non-enzymatic hydrolysis of acetyl-P produced by PAT
contributes to the observed levels of secreted acetate in
the mutant strains. Two double mutants, ack1 ack2 and
pat2–1 ack2, were also generated; both chloroplastic and
mitochondrial PAT/ACK pathways are blocked in each of
these strains. Increases in lactate production were
observed in the pat2–1 and pat2–1 ack2 mutants, suggesting differences in regulation of fermentation metabolism in
the pat2 genetic backgrounds; re-routing of metabolites
was always observed in the pat2–1 genetic background.
This may be expected as the metabolic block in the chloroplast in pat2–1 strains occurs at the level of acetyl CoA,
contributing to increased pyruvate accumulation. This
pyruvate may be readily re-directed toward lactate production for redox balancing, as observed in the adh1 and pfl1–
1 mutants (Figure 3) (Catalanotti et al., 2012; Magneschi
et al., 2012). Double mutants that have neither the chloroplastic nor mitochondrial PAT/ACK acetate-producing pathways (ack1 ack2 and pat2–1 ack2 double mutants) still
© 2015 The Authors
The Plant Journal © 2015 John Wiley & Sons Ltd, The Plant Journal, (2015), 82, 481–503
Dark hypoxic growth of algae 491
accumulated acetate in the medium during exposure to
anoxic conditions, albeit at lower levels than in wild-type
cells (approximately 50% relative to wild-type cells). These
results suggest that routes other than the PAT/ACK pathway function in acetate generation in these mutants (Yang
et al., 2014b). Interestingly, acetate production is also
retained through undetermined activities in mutants of
Clostridium species lacking ACK activity (Sillers et al.,
2008; Kuit et al., 2012). There are a number of potential
routes that may account for acetate production in the Chlamydomonas mutants blocked in both the chloroplastic and
mitochondrial PAT/ACK pathways. First, there may be
spontaneous hydrolysis of acetyl-P to acetate and Pi (Koshland, 1952; Di Sabato and Jencks, 1961). This reaction
denies the cell the ATP that is generated by the action of
ACK, which may potentially result in a diminished rate of
glycolysis and concomitant reduction of secretion of fermentation metabolites (Figure 3). Second, acetyl CoA
hydrolase activity (Tielens et al., 2010) may release acetate
and CoASH from acetyl CoA without ATP production.
Genes encoding homologs of acetyl CoA hydrolase are
present on the Chlamydomonas genome. This activity may
be important when acetyl CoA accumulates in cells and is
not rapidly metabolized by alternative. Third, aldehyde
dehydrogenase (ALDH) activity may oxidize acetaldehyde
(from pyruvate decarboxylation) to acetate (Kirch et al.,
2004, 2005; Brocker et al., 2013). In Chlamydomonas, pyruvate may be decarboxylated by PDC3 to generate acetaldehyde, which may then be oxidized to acetate by ALDH
activity, similar to the reaction used by some yeast (Remize
et al., 2000). However, this reaction involves formation of
NADH, and accumulation of NADH would reduce the rate
of glycolysis unless the cells were able to rapidly re-oxidize
it by production and excretion of a reduced organic compound. The excretion of metabolites that serve as this
‘reductant sink’ was not observed, even though some
ALDH transcripts (e.g. ALDH3) show significant accumulation in the various pat ack mutant strains (Yang et al.,
2014b). However, no significant ALDH activity was detected
among mutant and wild-type strains at various times after
imposition of dark anoxic conditions despite transcript
increases (Yang et al., 2014b). Fourth, acetate may be generated by acetyl CoA synthetases functioning in the
reverse direction. Many organisms are able to catalyze this
reaction using ADP-forming acetyl CoA synthetase (ADPACS) (Tielens et al., 2010), and a few reports have even
suggested that acetate production may also be achieved
by AMP-forming acetyl CoA synthetases (AMP-ACS) (Takasaki et al., 2004; Yoshii et al., 2009). However, AMP-ACS
enzymes typically function exclusively in the direction of
acetyl CoA synthesis (Tielens et al., 2010). No gene model
for an ADP-ACS was identified on the Chlamydomonas
genome. If AMP-ACS were used for acetate production,
ATP production would be retained, but we did not observe
significant changes in the levels of any transcript encoding
this enzyme in any of the mutants following exposure to
dark anoxic conditions; however, this activity has been
reported to be under post-translational control (Takasaki
et al., 2004). The AMP-ACS pathway is functionally equivalent to the PAT/ACK pathway in that ATP production is
retained, and decreased glycolysis and acetate excretion
are not expected if this compensatory mechanism is triggered in pat ack mutants. Finally, acetate:succinate CoA
transferase (ASCT) and succinyl CoA ligase (SCL) (van
Grinsven et al., 2008; Millerioux et al., 2012) may be
involved in acetate accumulation. ASCT transfers the CoA
moiety of acetyl CoA to succinate, and SCL converts succinyl CoA back to succinate. Although SCL homologs are
encoded on the Chlamydomonas genome, no ASCT homologs have been identified, and it is therefore unlikely that
this pathway represents a viable alternative for acetate production in Chlamydomonas (Atteia et al., 2013).
PAT and ACK activities are key enzymes of acetate-producing pathways in Chlamydomonas during hypoxia/
anoxia. Mutants defective for the genes encoding these
enzymes exhibit a reduction in the rate of glycolysis
(Yang et al., 2014b). Current data do not allow unambiguous conclusions regarding the origins of acetate production in the double mutants. However, the data do
suggest that acetate is formed without production of
ATP, as the rates of accumulation of all fermentation
products are attenuated. Two favored hypotheses consistent with the lack of ATP production during acetate synthesis include the possibility that acetyl-P is hydrolyzed
non-enzymatically in aqueous medium to acetate and
phosphate, as acetyl-P is not easily re-directed to other
metabolic pathways, or that acetyl CoA is hydrolyzed to
acetate and CoASH, which may be catalyzed by acyl/acetyl CoA hydrolases; other pathways for acetate production may also exist (Yang et al., 2014b).
Acetate transport and assimilation
AcpA, a member of the GPR1/FUN34/YaaH membrane protein family, is essential for acetate permease activity in the
hyphal fungus Aspergillus nidulans (Robellet et al., 2008).
In Saccharomyces cerevisiae, the ortholog of AcpA is Ady2
(Paiva et al., 2004). Based on homology to AcpA, five
genes encoding putative members of GPR1/FUN34/YaaH
family were identified on the Chlamydomonas genome
(Table 2 and Figure 3). The level of transcripts for two of
these putative acetate transporters increased during dark
to light transition (Duanmu et al., 2013). Ady2 was shown
to be important for the periodic ammonium export from
S. cerevisiae colonies observed during late development
(Palkova et al., 2002), indicating a potential relationship
between acetate and ammonium uptake/metabolism; these
relationships have not been explored in algae. Active acetate transport requires ATP, which has been used to
© 2015 The Authors
The Plant Journal © 2015 John Wiley & Sons Ltd, The Plant Journal, (2015), 82, 481–503
492 Wenqiang Yang et al.
Table 2 Enzymes potentially involved in acetate assimilation
Name
Phytozome9.0 ID
NCBI number
Annotation
Localization predication
TM domain number
ACK1
ACK2
PAT1
PAT2
ACS1
ACS2
ACS3
ACS4
ALDH1
ALDH2
ALDH3
ALDH4
ALDH5
ALDH6
ALDH7
ALDH8
SCLA1
SCLB1a
ACT1
CGLD2
YaaH-1
YaaH-2
yaaH-3
yaaH-4
yaaH-5
AMT1.1
AMT1.2
AMT1.3
AMT1.4
AMT1.5
AMT1.6
AMT1.7a
AMT1.7b
AMT1.8
Cre09.g396700
Cre17.g709850
Cre09.g396650
Cre17.g699000
g1290.t1
g1224.t1
Cre07.g353450
Cre01.g055500
g13400.t1
Cre16.g675650
Cre12.g500150
Cre12.g520350
Cre01.g033350
G8982.t1
g16809
Cre13.g605650
Cre03.g193850
g17060.t1
g16435.t1
g837.t1
Cre17.g700750
Cre17.g702900
Cre17.g702950
Cre17.g700450
Cre17.g700650
Cre03.g159254
Cre06.g293051
Cre14.g629920
Cre13.g569850
Cre09.g400750
Cre07.g355650
Cre02.g111050
Cre02.g111050
Cre12.g531000
XP_001694505
XP_001691682
XP_001691787
XP_001694504
XP_001700210
XP_001700230
XP_001702039
XP_001700230
XP_001694180
XP_001695943
XP_001690955
XP_001696928
XP_001690075
XP_001694332
XP_001698924
XP_001699134
XP_001693108
XP_001691581
XP_001692073
XP_001690113
XP_001691606
XP_001691586.1
XP_001691752
XP_001691608
XP_001691772
AF479643
AF530051
AF509497
AY542491
AY542492
AY548756
AY588244
AY548755
AY548754
Acetate kinase
Acetate kinase
Phosphate acetyltransferase
Phosphate acetyltransferase
Acetyl CoA synthetase
Acetyl CoA synthetase
Acetyl CoA synthetase
Acetyl CoA synthetase
Aldehyde dehydrogenase
Aldehyde dehydrogenase
Aldehyde dehydrogenase
Aldehyde dehydrogenase
Aldehyde dehydrogenase
Aldehyde dehydrogenase
Aldehyde dehydrogenase
Aldehyde dehydrogenase
Succinate CoA ligase
Succinate CoA ligase
Acyl CoA thioesterase
Acyl CoA thioesterase
GPR1/FUN34/yaaH family
GPR1/FUN34/yaaH family
GPR1/FUN34/yaaH family
GPR1/FUN34/yaaH family
GPR1/FUN34/yaaH family
Ammonium transporter
Ammonium transporter
Ammonium transporter
Ammonium transporter
Ammonium transporter
Ammonium transporter
Ammonium transporter
Ammonium transporter
Ammonium transporter
C
M
C
M
O
C
O
M or SP
O
M
M
M
SP
M
C
O or SP
M
M
O
M
Membrane
Membrane
Membrane
Membrane
Membrane
Membrane
Membrane
Membrane
Membrane
Membrane
Membrane
Membrane
Membrane
Membrane
0
0
0
0
0
0
0
0
0
0
0
0
0
0
0
0
0
0
0
0
6
6
6
6
6
11
11
10
11
10
11
11
11
10
The localization of ACS, ALDH, ammonium transporters (Fernandez and Galvan, 2007), the GPR1/FUN34/yaaH family of acetate transporters
and other possible acetate metabolism-related proteins were predicted using PredAlgo software (http://omictools.com/predalgo-s8353.html).
C, chloroplast; M, mitochondrion; SP, secretory pathway; O, other. The transmembrane (TM) domain number was predicted using the
TMHMM server v2.0 (http://www.cbs.dtu.dk/services/TMHMM/). ACS3 (Atteia et al., 2009; Terashima et al., 2010), ALD1/ALD2 (also named
ALDH3/ALDH8, Yang et al., 2014b) and SCLA1/SCLB1a are all localized to mitochondria (Atteia et al., 2009).
explain the low rates of Chlamydomonas acetate uptake
during anoxia in the dark (Gibbs et al., 1986). In the light,
CEF triggered by low O2 levels (Alric, 2010, 2014) maintains
cyclic photophosphorylation (ATP production) under hypoxic/anoxic conditions (Klob et al., 1973; Alric, 2014), and
the occurrence of cyclic photophosphorylation during
anoxia helps sustain acetate assimilation in Chlamydomonas mundana (Russell and Gibbs, 1968) and other
green algae (Wiessner, 1965), including Chlamydomonas
reinhardtii (Gibbs et al., 1986).
SUBCELLULAR LOCALIZATION AND
COMPARTMENTATION OF METABOLIC PATHWAYS
As discussed above, glycolysis is a conduit for eukaryotic
carbon and energy metabolism, leading to production of
pyruvate, ATP and NADH. When the cells become hypoxic
or anoxic, the pyruvate may be converted to metabolites
that serve as electron acceptors, allowing re-oxidization of
NADH formed as a consequence of glycolysis. These pathways for recycling electron carriers under hypoxic/anoxic
conditions (fermentation metabolism) may occur in different cellular compartments. For some eukaryotic organisms, including the protistan parasites such as Giardia and
€ ller et al., 2012), fermentation
Entamoeba species (Mu
occurs entirely in the cytosol. Fermentation may also occur
partly in hydrogenosomes, as is the case for Trichomonas
€ ller, 1993). In algae, end-products of glycolyvaginalis (Mu
sis may be metabolized in the cytosol, chloroplasts and
mitochondria. Furthermore, a number of metabolic reactions may occur in more than one cellular compartment,
with some enzymes routed to multiple locations or different isoforms of the enzyme targeted to different compart-
© 2015 The Authors
The Plant Journal © 2015 John Wiley & Sons Ltd, The Plant Journal, (2015), 82, 481–503
Dark hypoxic growth of algae 493
ments. For example, PFL1 is localized to both chloroplasts
and mitochondria in Chlamydomonas; dual localization of
proteins is not uncommon in eukaryotes (Atteia et al.,
€ ller et al., 2012). Furthermore,
2006; Martin, 2010; Mu
enzymes associated with glycolysis, the oxidative pentose
phosphate pathway and gluconeogenesis are located in
both the cytosol and chloroplasts. While the glycolytic and
oxidative pentose phosphate pathways in plants are localized in both the cytosol and chloroplasts (Plaxton, 1996;
Joyard et al., 2010), the glycolytic enzymes appear to be
differentially partitioned in Chlamydomonas; enzymes that
catalyze the formation of glyceraldehyde 3–phosphate
from glucose are located in chloroplasts, while enzymes
that transform 3–phosphoglycerate to pyruvate are located
in the cytosol (Ball, 1998; Johnson and Alric, 2013). Furthermore, while enzymes involved in acetate assimilation
and production are located in both mitochondria and chloroplasts, some Chlamydomonas TCA cycle enzymes are
localized to both mitochondria and to microbodies that
may represent glyoxysomes (Hayashi and Shinozaki, 2012;
Johnson and Alric, 2013); those present in the microbodies
probably participate in the glyoxylate cycle. Also, most cellular compartments require mechanisms for redox balancing and ATP synthesis, and some enzymes associated with
these activities may be shared among compartments, e.g.
the transhydrogenase is present in both mitochondria and
chloroplasts (Atteia et al., 2009; Terashima et al., 2011).
These considerations raise the fundamental question of
what events lead to the transfer of partial or entire metabolic pathways to new compartments. This issue is far
from resolved, and we are still uncertain of the localization
of many proteins in the cell. Furthermore, it is becoming
evident that, over evolutionary time, enzymes and pathways readily undergo re-compartmentation to the mitochondria, cytosol, hydrogenosomes or chloroplasts. In
some cases, gene duplications may lead to specialization
of the duplicated proteins, with each of the two paralogous
proteins targeted to a specific compartment and tailored
for function therein. Small changes in targeting sequences
resulting in mis-targeting may explain how individual
activities and even entire pathways become resident in
more than one cellular compartment (Martin, 2010). In
Chlamydomonas, the confirmed localization of PAT2 and
ACK1 in chloroplasts and PAT1 and ACK2 in mitochondria,
and the phenotypic consequences of lesions in genes
encoding each of these components, are helping to establish the functional importance of subcellular localization
and isoform specialization (Yang et al., 2014b).
CONTROL OF FERMENTATION METABOLISM
While progress has been made in defining algal fermentation pathways and elucidating their effect on cellular
metabolism, there are still numerous questions associated
with fermentation and anoxic/hypoxic metabolism. How
have fermentation pathways evolved in the algae? How
diverse are they among algae? How are they tailored to different environments? How are they regulated?
Regulation of genes encoding fermentative enzymes
Transcripts encoding many enzymes involved in fermentation accumulate in Chlamydomonas during anoxia (Mus
et al., 2007; Hemschemeier et al., 2013a) but others do not.
Expression of genes encoding the fermentative proteins
PDC3, lactate dehydrogenase and ADH2 was shown to be
primarily controlled by diurnal rhythms (Whitney et al.,
2011), while transcripts of genes encoding other proteins
such as PFR1 and HYDA1 exhibit marked accumulation at
the onset of anoxia (Mus et al., 2007). Furthermore, while
the level of PFR1 transcript increases at the onset of anoxia
(Mus et al., 2007), anoxia elicits an increase in PFL1 mRNA
levels, with no increase in PFL1 protein levels (Atteia et al.,
2006; Philipps et al., 2011; Catalanotti et al., 2012). These
results suggest that differences in the regulation of fermentation genes occur at both the transcriptional and translational levels. Interestingly, the molecular mass of PFL1
from anoxic cells was shown to be slightly less than the
molecular mass of the protein from cells maintained under
oxic conditions, suggesting that anoxic conditions trigger a
post-translational modification that may activate the
enzyme (Atteia et al., 2006; Catalanotti et al., 2012).
Several other factors also affect the transcriptional activity of fermentation genes. The rate of starch degradation
under anoxic conditions modulates intracellular NAD(P)H
levels and/or the oxidation state of the PQ pool, both of
which elicit changes in transcriptional activity of numerous
genes (Escoubas et al., 1995; Rutter et al., 2001; Pfannschmidt and Liere, 2005), while the production and detoxification of reactive oxygen species are probably also
important for controlling hypoxic responses (Antal et al.,
2003; Bailey-Serres and Chang, 2005; Guzy and Schumacker, 2006). For example, hydrogen peroxide (H2O2) synthesized by a NADPH oxidase is required for induction of ADH
in Arabidopsis (Baxter-Burrell et al., 2002).
Although few molecular studies have been performed to
elucidate the regulatory features controlling the genes and
proteins responsive to hypoxia/anoxia, exciting details concerning this regulation are beginning to emerge. A 21–
128 bp region upstream of the HYDA1 gene transcription
start site was initially shown to be involved in controlling
the expression of HYDA1 (Stirnberg and Happe, 2004).
Reporter gene analysis and electrophoretic mobility shift
assays demonstrated that CRR1, the copper-responsive
regulatory factor (Sommer et al., 2010), plays a role in
HYDA1 transcriptional control through its squamosa promoter-binding protein domain (Pape et al., 2012). Two consensus CRR1-binding GTAC motifs are present in the
HYDA1 promoter, and are necessary for full promoter
activity under hypoxic conditions; CRR1 binds to one of
© 2015 The Authors
The Plant Journal © 2015 John Wiley & Sons Ltd, The Plant Journal, (2015), 82, 481–503
494 Wenqiang Yang et al.
these GTAC cores in vitro (Pape et al., 2012). The same
GTAC motifs are present in the promoter of FDX5, which is
also regulated by CRR1 in response to copper levels and
O2 conditions (Lambertz et al., 2010). CRR1 plays an important role in regulating several genes encoding key proteins
(e.g. HYDA1 and PRF1) that are involved in dark, hypoxic
metabolism in Chlamydomonas, and, in particular, influences a subset of proteins that are also regulated under
conditions of copper deficiency (Hemschemeier et al.,
2013a). The importance of CRR1 in hypoxic metabolism is
underscored by the observation that crr1 mutants exhibit a
severe growth attenuation phenotype during hypoxia in
the light (Hemschemeier et al., 2013a). However, additional
hypoxia/anoxia regulatory strategies must exist that are
independent of CRR1, as the majority of the mRNAs that
differentially accumulate after dark, hypoxic acclimation
appear to be relatively insensitive to CRR1. For example,
approximately 1400 transcripts differentially accumulated
after acclimation of Chlamydomonas cells to dark, hypoxic
conditions, but only approximately 40 of these were aberrantly regulated in crr1 mutants under the same conditions
(Hemschemeier et al., 2013a). Moreover, HYDA1 transcript
accumulation is still observed in the crr1 mutant in
response to anoxia, albeit at attenuated levels (Pape et al.,
2012; Hemschemeier et al., 2013a). Overall, these data indicate that CRR1 has an important role in regulating the transcript levels of a subset of hypoxia-/anoxia-responsive
genes, but additional regulatory factors that have yet to be
identified must also play a significant role in transcriptional
responses to O2 availability.
O2 sensing/regulation in various organisms
In animals, prolyl 4–hydroxylases directly sense O2 and are
involved in controlling responses to anoxia (Guzy and Schumacker, 2006). A constitutively expressed hypoxia-inducing
factor is hydroxylated on conserved proline residues in the
presence of O2. This modification targets the hypoxia-inducing factor for ubiquitin-dependent degradation. In the
absence of O2, hydroxylation of the hypoxia-inducing factor
ceases, and the protein accumulates and triggers expression
of several target genes. Prolyl 4–hydroxylases may have
other protein targets that accumulate under anoxic conditions (not necessarily transcription/regulatory factors) and
are rapidly degraded as cells transition from anoxic to oxic
conditions (Semenza, 2011). In Arabidopsis and rice (Oryza
sativa), the levels of prolyl 4–hydroxylase transcripts are
strongly induced by O2 deprivation (Lasanthi-Kudahettige
et al., 2007; Vlad et al., 2007), raising the possibility that their
role as sensing elements may be conserved in plants.
There are also clear examples of the involvement of protein degradation in responses to anoxia in plants. The
N–end rule reflects an evolutionarily conserved mechanism
for eliciting protein degradation, whereby the N-terminal
amino acid is an important factor in determining the
half-life of the protein. In Arabidopsis, hypoxia-responsive
transcription factors are targeted for N–end degradation;
substrates for the pathway include ethylene response factor group VII transcription factors, which are susceptible
through a motif at their N-terminus, starting with Met-Cys.
In some plants, the regulators are not susceptible to this
degradation, and such plants are generally more tolerant
to hypoxic conditions. In rice, the dominant regulator of
hypoxia, SUB1A–1, is not a substrate for the N–end rule
degradation pathway (Gibbs et al., 2011; Licausi, 2011; Sasidharan and Mustroph, 2011).
In E. coli, there are two pathways that function in O2
sensing. One pathway is through Fnr (Bunn and Poyton,
1996), a global regulator of a large number of E. coli genes
that acts as either a transcriptional activator or repressor
(Spiro and Guest, 1991). The second pathway involves the
ArcA/ArcB two-component regulators (Bunn and Poyton,
1996). The response regulator ArcA may be phosphorylated by the sensor protein ArcB (Iuchi and Lin, 1992); the
latter is a histidine kinase that undergoes autophosphorylation under anoxic conditions (Kato et al., 1997). This phosphorylation cascade promotes acclimation to low O2
conditions by activating or repressing specific genes. In
Rhizobium meliloti, FixL and FixJ are two-component regulators that mediate the bacterium’s response to O2 conditions; these have been extensively studied with respect to
induction of nitrogen fixation genes under anaerobic conditions. The FixL protein is an O2 sensor (membrane protein) that behaves like ArcB in E. coli, phosphorylating the
response regulator FixJ (Monson et al., 1992). Phosphorylated FixJ activates nifA and fixK (Gilles-Gonzalez et al.,
1994), which encode two regulatory elements. NifA is
involved in expression of genes encoding subunits of
nitrogenase (and factors required to synthesize active
nitrogenase), while FixK controls expression of genes
required for microaerobic growth (Dixon and Kahn, 2004).
Yeast has more complicated O2 sensing regulatory mechanisms, involving the regulatory elements Hap1–5p, Mot3p,
Rox1p, Upc2p and Ecm22p (Kwast et al., 1998; Poyton,
1999; Hughes et al., 2005; Davies and Rine, 2006; Todd
et al., 2006; Hughes and Espenshade, 2008; Grahl and Cramer, 2010); a detailed description of the intricacies of this
system is beyond the scope of this review.
O2 sensing/regulation in Chlamydomonas
In Chlamydomonas, no O2 sensing regulatory factors analogous to mammalian hypoxia-inducing factors have been
identified. However, several of the 22 prolyl 4–hydroxylases encoded on the Chlamydomonas genome are significantly up-regulated in response to anoxia (Mus et al., 2007;
Hemschemeier et al., 2013a), and a subset of these are regulated to a degree by CRR1 (Hemschemeier et al., 2013a).
Although still highly speculative, determination of the
activity of one or more of these prolyl 4–hydroxylases in
© 2015 The Authors
The Plant Journal © 2015 John Wiley & Sons Ltd, The Plant Journal, (2015), 82, 481–503
Dark hypoxic growth of algae 495
Chlamydomonas may provide insight into how this alga
senses O2, maintains genes in an inactive state when O2 is
present, and targets key proteins involved in fermentation
metabolism for destruction as algal cells transition from
anoxic to oxic conditions. However, many (or all) of these
hydroxylases may not function in regulating hypoxic/
anoxic responses, but instead may modify the cell-wall
structure through proline hydroxylation; Chlamydomonas
has a proteinaceous, hydroxyproline-rich cell wall.
Murthy et al. (2012) recently used Chlamydomonas genome inspection to identify nine proteins with homology to
the O2 sensing, Per-Arnt-Sim (PAS)-heme domains present
in the FixL proteins of rhizobia (Murthy et al., 2012). Transcript levels for most of these proteins increase during
anoxia (Mus et al., 2007; Hemschemeier et al., 2013a), and
the PAS domains of two FixL-like proteins (FXL1 and FXL5)
were heterologously expressed in E. coli and shown to
bind heme and O2 at physiologically relevant concentrations. Although the FXL proteins, which are large proteins
(>1000 amino acids) with multiple predicted transmembrane domains, are candidate O2 sensors, experiments
directly linking them to physiological roles in O2 sensing
and signal transduction have yet to be reported.
As mentioned above, it has also been demonstrated that
a 2–on–2 hemoglobin designated THB8 has a critical role
in the Chlamydomonas anaerobic response (Hemschemeier et al., 2013b). Silencing of the THB8 gene causes both a
growth defect under anoxic conditions in the light and
mis-regulation of several genes that respond to hypoxic
conditions, including HYDA2 and CYG2 (encoding an adenylate/guanylate cyclase). The growth defect is exacerbated
by an NO scavenger, suggesting that the hypoxic/anoxic
responses in Chlamydomonas are at least partially controlled by both the 2–on–2 hemoglobin and an NO-dependent signaling pathway (Hemschemeier et al., 2013b). A
role for nitric oxide in O2 sensing has also been reported
for pea (Pisum sativum) (Borisjuk et al., 2007), and may
also be involved in the degradation of photosynthetic proteins in N-deprived Chlamydomonas cells (Wei et al.,
2014). However, it is still not clear whether the THB8 protein is part of the sensing mechanism, and more work is
required to determine whether the other 2–on–2 hemoglobins in Chlamydomonas function in anoxic/hypoxic acclimation and/or sensing of O2 levels.
Finally, there are also some studies showing that the
acclimation of plants to anoxic conditions may involve ethylene response factor transcriptional elements (Bailey-Serres and Voesenek, 2010; Gibbs et al., 2011; Licausi, 2011 ).
Chlamydomonas has putative ethylene response factor
transcription factors (Merchant et al., 2007), but none have
the cysteine at the N–terminus that has been associated
with O2 sensing in plants.
Together, these results suggest that there are a number
of different factors and mechanisms involved in regulating
fermentative processes in photosynthetic organisms. The
use of multiple mechanisms may enable metabolic versatility and fine tuning of the responses to dynamic environmental conditions. Full clarification of the regulatory
pathways, especially for Chlamydomonas, requires significantly more work in order to elucidate modes of sensing
an oxic/anoxic environment, and the diversity of transcriptional and post-transcriptional processes responsible for
eliciting acclimation responses.
Other metabolic strategies to cope with hypoxia/anoxia
Many photosynthetic organisms have evolved a set of
pathways, some of which generate a modest amount of
energy, that function during exposure to anoxic conditions
€ ller et al., 2012; Catalanotti et al., 2013). In addition,
(Mu
microbes and plants have also evolved a set of specific
strategies that they use to cope with hypoxia/anoxia. During starch breakdown, amylase levels increase in some
species to satisfy the increased carbon demand under hypoxic/anoxic conditions (Weigelt et al., 2009). In some species, to increase energy use efficiency, sucrose
degradation shifts from invertase to sucrose synthase to
form UDP-glucose, which uses pyrophosphate as the substrate to synthesize UTP/ATP; this shift helps to increase
net ATP production (Zeng et al., 1999). Pyruvate and glutamate may be converted to alanine and 2–oxoglutarate by
the alanine/2-oxoglutarate shunt, which prevents the loss
of carbon through fermentation pathways and yields ATP
through substrate level phosphorylation (Araujo et al.,
2012). Additionally, glutamic acid decarboxylase uses protons as its substrate and may help stabilize cytosolic pH
via the c–aminobutyric acid (GABA) shunt (Miyashita and
Good, 2008; Bailey-Serres et al., 2012), while a reduction in
the respiratory rate results from down-regulation of net
NADH production via the TCA cycle, reduced mETC activity
and/or triggering of mechanisms associated with O2 conservation (Chang et al., 2012).
Many plants produce ethanol as well as lactate during
hypoxia. The regulation of these pathways appears to be
under pH control; ethanol production appears to be critical
for the maintenance of cytosolic pH, as supported by data
demonstrating that a decrease in cytosolic pH of approximately 0.6 units favors ethanol production (Roberts et al.,
1989; Bailey-Serres and Voesenek, 2008; Catalanotti et al.,
2013). A less common fermentation process largely occurring in marine environments involves cytosolic opine formation. In this redox reaction, a pyruvate–amino acid
condensation regenerates NAD+. Possible advantages of
this reaction are redox balancing, cytosolic pH control, and
maintenance of osmotic equilibrium (Ballantyne, 2004).
Increased de-nitrification may also occur when cells experience anoxia. In fungi and other eukaryotic organisms,
there are two de-nitrification pathways; one is typically
localized to mitochondria and usually occurs under low O2
© 2015 The Authors
The Plant Journal © 2015 John Wiley & Sons Ltd, The Plant Journal, (2015), 82, 481–503
496 Wenqiang Yang et al.
conditions, while the other, often referred to as ammonia
fermentation, is localized in the cytosol (Takasaki et al.,
2004) and is activated under strict anoxic conditions. The
latter pathway involves reduction of nitrate to ammonium
using reductant generated by the catabolic oxidation of
ethanol (the donor of electrons) and concomitant acetate
synthesis, coupled to substrate-level phosphorylation
(Zhou et al., 2002). Nitrate respiration has been reported in
diatoms as a mechanism to survive dark, anoxic conditions
(Kamp et al., 2011). Finally, the generation of H2 in algal
chloroplasts may serve as a redox valve, although H2 production may also occur in mitochondria-like organelles in
the stramenopiles and in hydrogenosomes in the amoebo€ ller et al., 2012 and Catalanotti et al., 2013).
zoa (Mu
Mitochondrial respiration and chlororespiration
Inhibition of mitochondrial respiration appears to have at
least two major metabolic consequences. First, the flow of
electrons to O2 is blocked, leading to NADH accumulation
in the mitochondrion and cytosol, which probably results
in inhibition of the TCA cycle and glycolysis (e.g. at the
level of the pyruvate dehydrogenase complex). Second,
depletion of ATP during dark maintenance may promote
the glycolytic breakdown of starch/sugars, as ATP is an
allosteric inhibitor of hexokinase and phosphofructokinase
(Klock and Kreuzberg, 1991); when mitochondria are performing aerobic respiration, the substrate of phosphofructokinase, fructose-6–phosphate, is more abundant than its
product, fructose-1,6–bisphosphate, while this equilibrium
is reversed under anaerobic conditions when NAD(P)H is
not readily recycled (Klock and Kreuzberg, 1991). The consequences of the simultaneous slowing of respiratory NAD
(P)H oxidation and stimulation of upstream glycolytic steps
has an additive effect resulting in an elevated cellular
redox state. Furthermore, inhibition of both mitochondrial
respiration and chlororespiration leads to reduction of
chloroplastic electron carriers, including the PQ pool;
either of these respiratory processes appears to be sufficient to re-oxidize most NAD(P)H produced by glycolysis
(Alric, 2010, 2014). In addition, another enzyme that is
likely to have a major effect on NAD(P)H accumulation in
the cytosol and chloroplasts during anoxic growth is the
glycolytic enzyme glyceraldehyde phosphate dehydrogenase. Under dark aerobic conditions, a downstream product of this reaction, 3–phosphoglycerate, is more abundant
than glyceraldehyde-3-phosphate (the substrate of glyceraldehyde phosphate dehydrogenase), suggesting rapid
oxidation of NADPH in the presence of O2. This equilibrium
is reversed under anaerobic conditions (Klock and Kreuzberg, 1991), when cellular NAD(P)H levels increase.
Another reaction that is likely to affect cellular redox to
some extent is catalyzed by glucose-6–phosphate dehydrogenase and inhibited by NADPH (Lendzian and Bassham,
1975). MDH, which is associated with the glyoxylate cycle
(in the cytosol), the TCA cycle (in the mitochondrion) and
chloroplast metabolism, may also contribute to cellular
redox conditions, although, in the absence of net acetate
assimilation (cells maintained in minimal medium), the
production of NAD(P)H by MDH in the dark is negligible.
Redox regulation
The redox conditions of photosynthetic organisms have
profound effects on their physiological and metabolic processes. Changes in activities of catalytic processes as well
as the organization of macromolecular complexes in membranes may accompany redox changes associated with the
dark and hypoxic/anoxic and high-light conditions. Anoxia
creates a more reduced stromal redox poise, which has
been shown to enhance CEF measured in the presence of
3–(3,4-dichlorophenyl)-1,1–dimethylurea (DCMU) in wildtype Chlamydomonas cells. This CEF enhancement was
not observed in a pgrl1 mutant (Tolleter et al., 2011). The
association of PGRL1 with the PSI–light-harvesting complex I supercomplex, which is involved in CEF, was favored
in Chlamydomonas cells maintained under anoxic conditions in the light (Iwai et al., 2010; Takahashi et al., 2013).
In the moss Physcomitrella patens, quantitative proteomics
demonstrated severe down-regulation of the photosystems
but up-regulation of the chloroplastic NADH dehydrogenase complex, plastocyanin, and Ca2+ sensors in the pgrl1
mutant, indicating that, in the absence of PGRL1, a set of
metabolic reactions may be elicited to compensate for
decreased CEF under anoxic light conditions (Kukuczka
et al., 2014). Furthermore, Ca2+ sensor (CAS) and Anaerobic Response 1 (ANR1) proteins showed increased abundance under anoxic conditions, associate with each other
and with PGRL1, and all become part of a large active PSI–
cytochrome b6f complex performing CEF (Terashima et al.,
2012). Furthermore, pgrl1 knockdown lines exhibited
hypersensitivity to iron deficiency, linking Fe limitation to
the formation/remodeling of the supercomplex associated
with CEF (Petroutsos et al., 2009). It was also shown that
conformational changes in the PGRL1 protein are linked to
the cellular redox state (Johnson et al., 2014). Phenotypic
comparative analyses have demonstrated that PGRL1 is
crucial for acclimation of Chlamydomonas cells to high
light and anoxia; analyses of the double mutant pgrl1 npq4
(where the gene disrupted in npq4 encodes LHCSR3) confirmed a complementary role of PGRL1 and LHCSR3 in
managing excess absorbed excitation energy (Kukuczka
et al., 2014). In addition, both proteins are required for
photoprotection and for survival of the cells under low O2
(Kukuczka et al., 2014). The integrated interactions between
redox, high light and anoxia are still being decoded; however, it is becoming clear that the overlapping features of
these conditions elicit overlapping regulatory processes
and use of at least some shared regulatory elements to tailor the activities of the metabolic machinery to cellular con-
© 2015 The Authors
The Plant Journal © 2015 John Wiley & Sons Ltd, The Plant Journal, (2015), 82, 481–503
Dark hypoxic growth of algae 497
ditions. There are several other energetic and redox considerations that distinguish light from dark growth in photosynthetic organisms. NADPH plays a critical role in
driving anabolic processes, including the synthesis of lipids, amino acids and nucleotides, and is directly produced
by the activity of FDX:NADP+ oxidoreductase. During dark
metabolism, many reactions including those of the TCA
cycle and glycolysis generate NADH; the oxidative pentose
phosphate pathway produces NADPH. Interconversion
between NADH and NADPH may be achieved by the pyridine nucleotide transhydrogenase (Agledal et al., 2010;
Holm et al., 2010). This enzyme regulates the NAD(H)/
NADP(H) ratio through a reversible hydride transfer that
occurs in either an energy-dependent or energy-independent manner (Olausson et al., 1992; Pedersen et al., 2008);
the NAD(H)/NADP(H) ratio helps to control the extent to
which the cells perform catabolic and anabolic processes.
Some bacteria, including E. coli, rely heavily on pyridine
nucleotide transhydrogenase activity to modulate metabolism (Sauer et al., 2004; Fuhrer and Sauer, 2009).
Other electron carriers, such as the FDXs and thioredoxins, are small redox carriers that supply electrons to a range
of cellular processes, as previously discussed. Furthermore,
while some of the FDX proteins may be efficiently reduced
by NADH or NADPH, FDXs with a very negative redox
potential may only be able to acquire electrons through PSI,
which suggests tailoring of redox components in the light
and the dark. As mentioned above, we isolated a mutant of
Chlamydomonas that does not grow in the dark (but does
grow in the light) and is null for FDX5 (W. Yang, unpublished results). This result supports the concept that the
FDX family in Chlamydomonas represents a group of proteins with a specialized function as electron carriers, but
their functions may only be possible in the light (when PSI
through FDX1 supplies much of the reductant) or dark
(where NADH supplies most of the reductant). More information is required with respect to the redox potential of the
various FDXs and the affinities with which they interact with
their specific target proteins. Other redox carriers such as
thioredoxins may also be critical for ‘dark’ metabolism.
PERSPECTIVES
Chlamydomonas is a metabolically versatile organism that
performs photosynthetic CO2 fixation, aerobic respiration
and anaerobic fermentation. This alga is a model for
examining many aspects of photosynthetic metabolism,
and has been the subject of numerous metabolic studies.
Many pathways and enzymes associated with fermentation metabolism in this organism are only now being
defined, and almost nothing is known about mechanisms
by which these pathways are regulated, or the ways in
which fermentation products are partitioned among the
various cell compartments. The generation of lesions that
block some of these pathways is providing new insights
into compensatory responses that allow sustained ATP
production while eliminating reducing equivalents
through generation of reduced carbon compounds that
are excreted from cells. Initial characterizations of Chlamydomonas have demonstrated that this alga has flexible, mixed-acid fermentation pathways, with features
common to bacterial-, plant- and yeast-type fermentation.
Most enzymes for fermentative metabolism in the algae,
inferred from genomic and metabolic studies, have not
been biochemically characterized. Expression patterns of
genes encoding these enzymes, the biochemical properties of these enzymes (including potential interactions
with each other), and the diversity of fermentation pathways plus the extent to which they are used under various conditions, require further examination in a broader
spectrum of algal systems. Additionally, the diversity of
external and internal end-products accumulated by various algae during fermentation is still mostly unknown.
Such information is critical for developing a clear understanding of metabolic diversity both within and among
the various algal groups, and the ways in which fermentation pathways have been shaped by environmental conditions. Furthermore, there are many technologies,
including flux balance analysis, mass flux analysis, timeresolved fluorescence measurements and the use of O2
microsensors that may help to evaluate the redox conditions of cells and correlate those conditions with the activities of both oxic and anoxic metabolisms. An
understanding of the various pathways critical for dark
metabolism and the ways in which these pathways are
controlled constitutes a domain of metabolism that must
be fully described if we are to understand the energy budget of photosynthetic microbes in the environment and
potential ways to manipulate carbon cycling. Finally, fermentation metabolism in algae appears to represent a significant ecological component of carbon flux in soils (and
sediments) that strongly affects its content of organic
acids, alcohols and H2, which in turn affects the biotic
composition of the ecosystem.
ACKNOWLEDGEMENTS
The work performed in our laboratories and described here was
supported by grants from the US the Department of Energy (numbers DE-FG02-12ER16338 and DE-FG02-12ER16339). Aspects of the
work were also funded by US National Science Foundation grants
to A.R.G. (MCB0824469 and MCB-0951094).
REFERENCES
Agledal, L., Niere, M. and Ziegler, M. (2010) The phosphate makes a difference: cellular functions of NADP+. Redox Rep. 15, 2–10.
Aksoy, M., Pootakham, W., Pollock, S.V., Moseley, J.L., Gonzalez-Ballester,
D. and Grossman, A.R. (2013) Tiered regulation of sulfur deprivation
responses in Chlamydomonas reinhardtii and identification of an associated regulatory factor. Plant Physiol. 162, 195–211.
Allen, F.L. and Horwitz, L. (1957) Oxygen evolution and photoreduction in
adapted Scenedesmus. Arch. Biochem. Biophys. 66, 45–63.
© 2015 The Authors
The Plant Journal © 2015 John Wiley & Sons Ltd, The Plant Journal, (2015), 82, 481–503
498 Wenqiang Yang et al.
Alric, J. (2010) Cyclic electron flow around photosystem I in unicellular
green algae. Photosyn. Res. 106, 47–56.
Alric, J. (2014) Redox and ATP control of photosynthetic cyclic electron
flow in Chlamydomonas reinhardtii: (II) involvement of the PGR5-PGRL1
pathway under anaerobic conditions. Biochim. Biophys. Acta 1837, 825–
834.
Ananyev, G., Carrieri, D. and Dismukes, G.C. (2008) Optimization of metabolic capacity and flux through environmental cues to maximize hydrogen production by the cyanobacterium “Arthrospira (Spirulina)
maxima”. Appl. Environ. Microbiol. 74, 6102–6113.
Anderson, J.W. and House, C.M. (1979) Polarographic study of oxaloacetate
reduction by isolated pea chloroplasts. Plant Physiol. 64, 1058–1063.
Antal, T.K., Krendeleva, T.E., Laurinavichene, T.V., Makarova, V.V., Ghirardi,
M.L., Rubin, A.B., Tsygankov, A.A. and Seibert, M. (2003) The dependence of algal H2 production on Photosystem II and O2 consumption
activities in sulfur-deprived Chlamydomonas reinhardtii cells. Biochim.
Biophys. Acta 1607, 153–160.
Araujo, W.L., Tohge, T., Nunes-Nesi, A., Daloso, D.M., Nimick, M., Krahnert,
I., Bunik, V.I., Moorhead, G.B. and Fernie, A.R. (2012) Phosphonate analogs of 2-oxoglutarate perturb metabolism and gene expression in illuminated Arabidopsis leaves. Front. Plant Sci. 3, 114.
Atteia, A., van Lis, R., Gelius-Dietrich, G., Adrait, A., Garin, J., Joyard, J.,
Rolland, N. and Martin, W. (2006) Pyruvate formate-lyase and a novel
route of eukaryotic ATP synthesis in Chlamydomonas mitochondria. J.
Biol. Chem. 281, 9909–9918.
Atteia, A., Adrait, A., Brugiere, S. et al. (2009) A proteomic survey of Chlamydomonas reinhardtii mitochondria sheds new light on the metabolic
plasticity of the organelle and on the nature of the alpha-proteobacterial
mitochondrial ancestor. Mol. Biol. Evol. 26, 1533–1548.
Atteia, A., van Lis, R., Tielens, A.G. and Martin, W.F. (2013) Anaerobic
energy metabolism in unicellular photosynthetic eukaryotes. Biochim.
Biophys. Acta 1827, 210–223.
Badin, E.J. and Calvin, M.J. (1950) The path of carbon in photosynthesis. IX.
Photosynthesis, photoreduction and the hydrogenoxygen-carbon dioxide
dark reduction. Am. Chem. Soc. 72, 5266–5270.
Bailey-Serres, J. and Chang, R. (2005) Sensing and signaling in response
to oxygen deprivation in plants and other organisms. Ann. Bot. 96, 507–
518.
Bailey-Serres, J. and Voesenek, L.A. (2008) Flooding stress: acclimations
and genetic diversity. Annu. Rev. Plant Biol. 59, 313–339.
Bailey-Serres, J. and Voesenek, L.A. (2010) Life in the balance: a signaling network controlling survival of flooding. Curr. Opin. Plant Biol. 13, 489–494.
Bailey-Serres, J., Fukao, T., Gibbs, D.J., Holdsworth, M.J., Lee, S.C., Licausi,
F., Perata, P., Voesenek, L.A. and van Dongen, J.T. (2012) Making sense
of low oxygen sensing. Trends Plant Sci. 17, 129–138.
Ball, S. (1998) Regulation of starch biosynthesis. In The Molecular Biology
of Chloroplasts and Mitochondria in Chlamydomonas. Advances in photosynthesis. Govindjee (ed) vol 7. (Rochaix, J.D., Goldschmidt-Clermont,
M. and Merchant, S., eds). Dortrecht, Netherlands: Kluwer Academic
Publishers, pp, 550–567.
Ballantyne, J.S. (2004) Mitochondria: aerobic and anaerobic design–lessons
from molluscs and fishes. Comp. Biochem. Physiol. 139, 461–467.
Barbieri, M.R., Larosa, V., Nouet, C., Subrahmanian, N., Remacle, C. and Hamel, P.P. (2011) A forward genetic screen identifies mutants deficient for
mitochondrial complex I assembly in Chlamydomonas reinhardtii.
Genetics, 188, 349–358.
Baxter-Burrell, A., Yang, Z., Springer, P.S. and Bailey-Serres, J. (2002) RopGAP4-dependent Rop GTPase rheostat control of Arabidopsis oxygen
deprivation tolerance. Science, 296, 2026–2028.
Bedhomme, M., Zaffagnini, M., Marchand, C.H., Gao, X.H., Moslonka-Lefebvre, M., Michelet, L., Decottignies, P. and Lemaire, S.D. (2009) Regulation
by glutathionylation of isocitrate lyase from Chlamydomonas reinhardtii.
J. Biol. Chem. 284, 36282–36291.
Bennoun, P. (2002) The present model for chlororespiration. Photosynth.
Res. 73, 273–277.
Borisjuk, L., Macherel, D., Benamar, A., Wobus, U. and Rolletschek, H.
(2007) Low oxygen sensing and balancing in plant seeds: a role for nitric
oxide. New Phytol. 176, 813–823.
Boschetti, A. and Schmid., K. (1998) Energy supply for ATP-synthase deficient chloroplasts of Chlamydomonas reinhardtii. Plant Cell Physiol. 39,
160–168.
Brocker, C., Vasiliou, M., Carpenter, S. et al. (2013) Aldehyde dehydrogenase (ALDH) superfamily in plants: gene nomenclature and comparative
genomics. Planta, 237, 189–210.
Bulte, L., Gans, P., Rebeille , F. and Wollman, F.A. (1990) ATP control on
state transitions in vivo in Chlamydomonas reinhardtii. Biochim. Biophys. Acta, 1020, 72–80.
Bunn, H.F. and Poyton, R.O. (1996) Oxygen sensing and molecular adaptation to hypoxia. Physiol. Rev. 76, 839–885.
Burgess, S.J., Tredwell, G., Molnar, A., Bundy, J.G. and Nixon, P.J. (2012)
Artificial microRNA-mediated knockdown of pyruvate formate lyase
(PFL1) provides evidence for an active 3-hydroxybutyrate production
pathway in the green alga Chlamydomonas reinhardtii. J. Biotech. 162,
57–66.
Cao, M., Meng, D., Wang, L., Bei, S., Snell, W.J. and Pan, J. (2013) Activation loop phosphorylation of a protein kinase is a molecular marker of
organelle size that dynamically reports flagellar length. Proc. Natl Acad.
Sci. USA, 110, 12337–12342.
Cardol, P., Matagne, R.F. and Remacle, C. (2002) Impact of mutations affecting ND mitochondria-encoded subunits on the activity and assembly of
complex I in Chlamydomonas. Implication for the structural organization
of the enzyme. J. Mol. Biol. 319, 1211–1221.
Cardol, P., Lapaille, M., Minet, P., Franck, F., Matagne, R.F. and Remacle, C.
(2006) ND3 and ND4L subunits of mitochondrial complex I, both nucleus
encoded in Chlamydomonas reinhardtii, are required for activity and
assembly of the enzyme. Eukaryot. Cell 5, 1460–1467.
Cardol, P., Boutaffala, L., Memmi, S., Devreese, B., Matagne, R.F. and Remacle, C. (2008) In Chlamydomonas, the loss of ND5 subunit prevents the
assembly of whole mitochondrial complex I and leads to the formation
of a low abundant 700 kDa subcomplex. Biochim. Biophys. Acta, 1777,
388–396.
Cardol, P., Alric, J., Girard-Bascou, J., Franck, F., Wollman, F.A. and Finazzi,
G. (2009) Impaired respiration discloses the physiological significance of
state transitions in Chlamydomonas. Proc. Natl Acad. Sci. USA, 106,
15979–15984.
Carrieri, D., Momot, D., Brasg, I.A., Ananyev, G., Lenz, O., Bryant, D.A. and
Dismukes, G.C. (2010) Boosting autofermentation rates and product
yields with sodium stress cycling: application to production of renewable
fuels by cyanobacteria. Appl. Environ. Microbiol. 76, 6455–6462.
Catalanotti, C., Dubini, A., Subramanian, V., Yang, W., Magneschi, L., Mus,
F., Seibert, M., Posewitz, M.C. and Grossman, A.R. (2012) Altered fermentative metabolism in Chlamydomonas reinhardtii mutants lacking
pyruvate formate lyase and both pyruvate formate lyase and alcohol
dehydrogenase. Plant Cell, 24, 692–707.
Catalanotti, C., Yang, W., Posewitz, M.C. and Grossman, A.R. (2013) Fermentation metabolism and its evolution in algae. Front. Plant Sci. 4, 150.
Chang, R., Jang, C.J., Branco-Price, C., Nghiem, P. and Bailey-Serres, J.
(2012) Transient MPK6 activation in response to oxygen deprivation and
reoxygenation is mediated by mitochondria and aids seedling survival in
Arabidopsis. Plant Mol. Biol. 78, 109–122.
Chen, C. and Gibbs, M. (1992) Coupling of carbon dioxide fixation to the
oxyhydrogen reaction in the isolated chloroplast of Chlamydomonas
reinhardtii. Plant Physiol. 100, 1361–1365.
Chochois, V., Dauvillee, D., Beyly, A., Tolleter, D., Cuine, S., Timpano, H.,
Ball, S., Cournac, L. and Peltier, G. (2009) Hydrogen production in Chlamydomonas: photosystem II-dependent and -independent pathways differ in their requirement for starch metabolism. Plant Physiol. 151, 631–
640.
Colin, M., Dorthu, M.P., Duby, F., Remacle, C., Dinant, M., Wolwertz, M.R.,
Duyckaerts, C., Sluse, F. and Matagne, R.F. (1995) Mutations affecting
the mitochondrial genes encoding the cytochrome oxidase subunit I and
apocytochrome b of Chlamydomonas reinhardtii. Mol. Gen. Genet. 249,
179–184.
Cournac, L., Redding, K., Ravenel, J., Rumeau, D., Josse, E.M., Kuntz, M.
and Peltier, G. (2000) Electron flow between photosystem II and oxygen in chloroplasts of photosystem I-deficient algae is mediated by a
quinol oxidase involved in chlororespiration. J. Biol. Chem. 275, 17256–
17262.
Dang, K.V., Plet, J., Tolleter, D. et al. (2014) Combined increases in mitochondrial cooperation and oxygen photoreduction compensate for deficiency in cyclic electron flow in Chlamydomonas reinhardtii. Plant Cell,
26, 3036–3050.
© 2015 The Authors
The Plant Journal © 2015 John Wiley & Sons Ltd, The Plant Journal, (2015), 82, 481–503
Dark hypoxic growth of algae 499
Davies, B.S. and Rine, J. (2006) A role for sterol levels in oxygen sensing in
Saccharomyces cerevisiae. Genetics, 174, 191–201.
Di Sabato, G. and Jencks, W.P. (1961) Mechanism and catalysis of reactions
of acyl phosphates. II. Hydrolysis1. J. Am. Chem. Soc. 83, 4400–4405.
Dixon, R. and Kahn, D. (2004) Genetic regulation of biological nitrogen fixation. Nat. Rev. Microbiol. 2, 621–631.
Doebbe, A., Rupprecht, J., Beckmann, J., Mussgnug, J.H., Hallmann, A.,
Hankamer, B. and Kruse, O. (2007) Functional integration of the HUP1
hexose symporter gene into the genome of C. reinhardtii: Impacts on
biological H2 production. J. Biotechnol. 131, 27–33.
Doebbe, A., Keck, M., La Russa, M., Mussgnug, J.H., Hankamer, B., Tekce,
E., Niehaus, K. and Kruse, O. (2010) The interplay of proton, electron,
and metabolite supply for photosynthetic H2 production in Chlamydomonas reinhardtii. J. Biol. Chem. 285, 30247–30260.
Dorthu, M.P., Remy, S., Michel-Wolwertz, M.R., Colleaux, L., Breyer, D.,
Beckers, M.C., Englebert, S., Duyckaerts, C., Sluse, F.E. and Matagne,
R.F. (1992) Biochemical, genetic and molecular characterization of new
respiratory-deficient mutants in Chlamydomonas reinhardtii. Plant Mol.
Biol. 18, 759–772.
Duanmu, D., Casero, D., Dent, R.M. et al. (2013) Retrograde bilin signaling
enables Chlamydomonas greening and phototrophic survival. Proc. Natl
Acad. Sci. USA, 110, 3621–3626.
Dubini, A., Mus, F., Seibert, M., Grossman, A.R. and Posewitz, M.C. (2009)
Flexibility in anaerobic metabolism as revealed in a mutant of Chlamydomonas reinhardtii lacking hydrogenase activity. J. Biol. Chem. 284, 7201–
7213.
Duby, F. and Matagne, R.F. (1999) Alteration of dark respiration and reduction of phototrophic growth in a mitochondrial DNA deletion mutant of
Chlamydomonas lacking cob, nd4, and the 3’ end of nd5. Plant Cell, 11,
115–125.
Dutcher, S.K. (1995) Flagellar assembly in two hundred and fifty easy-to-follow steps. Trends Genet. 11, 398–404.
Erbes, D.L. and Gibbs, M. (1981) Kinetics of the oxyhydrogen reaction in the
presence and absence of carbon dioxide in Scenedesmus obliquus. Plant
Physiol. 67, 129–132.
Eriksen, N.T. (2008) Production of phycocyanin – a pigment with applications in biology, biotechnology, foods and medicines. Appl. Microbiol.
Biotechnol. 80, 1–4.
Escoubas, J.M., Lomas, M., LaRoche, J. and Falkowski, P.G. (1995) Light intensity regulation of cab gene transcription is signaled by the redox state of
the plastoquinone pool. Proc. Natl Acad. Sci. USA, 92, 10237–10241.
Fang, W., Si, Y., Douglass, S., Casero, D., Merchant, S.S., Pellegrini, M., Ladunga, I., Liu, P. and Spalding, M.H. (2012) Transcriptome-wide changes
in Chlamydomonas reinhardtii gene expression regulated by carbon
dioxide and the CO2-concentrating mechanism regulator CIA5/CCM1.
Plant Cell, 24, 1876–1893.
(2007) Inorganic nitrogen assimilation in ChlaFernandez, E. and Galva n, A.
mydomonas. J. Exp. Bot. 58, 2279–2287.
Frenkel, A., Gaffron, H. and Battley, E.H. (1949) Photosynthesis and photoreduction by a species of blue-green algae. Biol. Bull. 97, 269.
Fuhrer, T. and Sauer, U. (2009) Different biochemical mechanisms ensure
network-wide balancing of reducing equivalents in microbial metabolism. J. Bacteriol. 191, 2112–2121.
Gaffron, H. (1939) Reduction of carbon dioxide with molecular hydrogen in
green algae. Nature, 123, 204–205.
Gaffron, H. (1940) The oxyhydrogen reaction in green algae and the reduction of carbon dioxide in the dark. Science, 91, 529–530.
Gaffron, H. (1942a) Reduction of carbon dioxide coupled with the oxyhydrogen reaction in Algae. J. Gen. Physiol. 26, 241–267.
Gaffron, H. (1942b) The effect of specific poisons upon the photo-reduction
with hydrogen in green algae. J. Gen. Physiol. 26, 195–217.
Gans, P. and Rebeille, F. (1990) Control in the dark of the plastoquinone
redox state by mitochondrial activity in Chlamydomonas reinhardtii. Biochim. Biophys. Acta, 1015, 150–155.
Gfeller, R.P. and Gibbs, M. (1984) Fermentative metabolism of Chlamydomonas reinhardtii: I. Analysis of fermentative products from starch in
dark and light. Plant Physiol. 75, 212–218.
Gfeller, R.P. and Gibbs, M. (1985) Fermentative metabolism of Chlamydomonas reinhardtii: II. Role of plastoquinone. Plant Physiol. 77, 509–511.
Ghirardi, M.L., Togasaki, R.K. and Seibert, M. (1997) Oxygen sensitivity of
algal H2-production. Appl. Biochem. Biotechnol. 63–65, 141–151.
Ghirardi, M.L., Dubini, A., Yu, J. and Maness, P.C. (2009) Photobiological
hydrogen-producing systems. Chem. Soc. Rev. 38, 52–61.
Ghysels, B., Godaux, D., Matagne, R.F., Cardol, P. and Franck, F. (2013)
Function of the chloroplast hydrogenase in the microalga Chlamydomonas: the role of hydrogenase and state transitions during photosynthetic activation in anaerobiosis. PLoS ONE, 8, e64161.
Gibbs, M., Gfeller, R.P. and Chen, C. (1986) Fermentative metabolism of
Chlamydomonas reinhardii: III. Photoassimilation of acetate. Plant Physiol. 82, 160–166.
Gibbs, D.J., Lee, S.C., Isa, N.M. et al. (2011) Homeostatic response to
hypoxia is regulated by the N-end rule pathway in plants. Nature, 479,
415–418.
Gilles-Gonzalez, M.A., Gonzalez, G., Perutz, M.F., Kiger, L., Marden, M.C.
and Poyart, C. (1994) Heme-based sensors, exemplified by the kinase
FixL, are a new class of heme protein with distinctive ligand binding and
autoxidation. Biochemistry, 33, 8067–8073.
€ rkman, O. (1995) Temperature-sensitive coupling and
Gilmore, A.M. and Bjo
uncoupling of ATPase-mediated, nonradiative energy dissipation: similarities between chloroplasts and leaves. Planta, 197, 646–654.
Gingras, G., Goldsby, R.A. and Calvin, M. (1963) Carbon dioxide metabolism
in hydrogen-adapted Scenedesmus. Arch. Biochem. Biophys. 100, 178–
184.
Gonzalez-Ballester, D., Casero, D., Cokus, S., Pellegrini, M., Merchant, S.S.
and Grossman, A.R. (2010) RNA-seq analysis of sulfur-deprived Chlamydomonas cells reveals aspects of acclimation critical for cell survival.
Plant Cell, 22, 2058–2084.
Gonzalez-Ballester, D., Pootakham, W., Mus, F. et al. (2011) Reverse genetics in Chlamydomonas: a platform for isolating insertional mutants. Plant
Methods, 7, 24.
Grahl, N. and Cramer, R.A. Jr (2010) Regulation of hypoxia adaptation: an
overlooked virulence attribute of pathogenic fungi? Med. Mycol. J. 48,
1–15.
Gray, M.W. and Boer, P.H. (1988) Organization and expression of algal
(Chlamydomonas reinhardtii) mitochondrial DNA. Philos. Trans. R. Soc.
Lond. 319, 135–147.
van Grinsven, K.W., Rosnowsky, S., van Weelden, S.W., Putz, S., van der
Giezen, M., Martin, W., van Hellemond, J.J., Tielens, A.G. and Henze, K.
(2008) Acetate:succinate CoA-transferase in the hydrogenosomes of
Trichomonas vaginalis: identification and characterization. J. Biol. Chem.
283, 1411–1418.
Grossman, A.R., Croft, M., Gladyshev, V.N., Merchant, S.S., Posewitz, M.C.,
Prochnik, S. and Spalding, M.H. (2007) Novel metabolism in Chlamydomonas through the lens of genomics. Curr. Opin. Plant Biol. 10, 190–198.
Grossman, A.R., Catalanotti, C., Yang, W., Dubini, A., Magneschi, L., Subramanian, V., Posewitz, M.C. and Seibert, M. (2011) Multiple facets of
anoxic metabolism and hydrogen production in the unicellular green
alga Chlamydomonas reinhardtii. New Phytol. 190, 279–288.
Guzy, R.D. and Schumacker, P.T. (2006) Oxygen sensing by mitochondria at
complex III: the paradox of increased reactive oxygen species during
hypoxia. Exp. Physiol. 91, 807–819.
Harris, E.H. (2001) Chlamydomonas as a Model Organism. Annu. Rev. Plant
Physiol. Plant Mol. Biol. 52, 363–406.
Hayashi, Y. and Shinozaki, A. (2012) Visualization of microbodies in Chlamydomonas reinhardtii. J. Plant. Res. 125, 579–586.
Heineke, D., Riens, B., Grosse, H., Hoferichter, P., Peter, U., Flugge, U.I. and
Heldt, H.W. (1991) Redox transfer across the inner chloroplast envelope
membrane. Plant Physiol. 95, 1131–1137.
Heinnickel, M.L., Alric, J., Wittkopp, T., Yang, W., Catalanotti, C., Dent, R.,
Niyogi, K.K., Wollman, F.A. and Grossman, A.R. (2013) Novel thylakoid
membrane GreenCut protein CPLD38 impacts accumulation of the cytochrome b6f complex and associated regulatory processes. J. Biol. Chem.
288, 7024–7036.
Hemschemeier, A. and Happe, T. (2005) The exceptional photofermentative
hydrogen metabolism of the green alga Chlamydomonas reinhardtii.
Biochem. Soc. Trans. 33, 39–41.
Hemschemeier, A., Casero, D., Liu, B., Benning, C., Pellegrini, M., Happe, T.
and Merchant, S.S. (2013a) Copper response regulator1-dependent and independent responses of the Chlamydomonas reinhardtii transcriptome
to dark anoxia. Plant Cell, 25, 3186–3211.
Hemschemeier, A., Duner, M., Casero, D., Merchant, S.S., Winkler, M. and
Happe, T. (2013b) Hypoxic survival requires a 2–on–2 hemoglobin in a
© 2015 The Authors
The Plant Journal © 2015 John Wiley & Sons Ltd, The Plant Journal, (2015), 82, 481–503
500 Wenqiang Yang et al.
process involving nitric oxide. Proc. Natl Acad. Sci. USA, 110, 10854–
10859.
Hoefnagel, M.H. and Wiskich, J.T. (1998) Activation of the plant alternative
oxidase by high reduction levels of the Q-pool and pyruvate. Arch. Biochem. Biophys. 355, 262–270.
Hoehler, T.M., Albert, D.B., Alperin, M.J., Bebout, B.M., Martens, C.S. and
Des Marais, D.J. (2002) Comparative ecology of H2 cycling in sedimentary and phototrophic ecosystems. Antonie Van Leeuwenhoek, 81, 575–
585.
Holm, A.K., Blank, L.M., Oldiges, M., Schmid, A., Solem, C., Jensen, P.R.
and Vemuri, G.N. (2010) Metabolic and transcriptional response to cofactor perturbations in Escherichia coli. J. Biol. Chem. 285, 17498–17506.
Hoober, J.K., Boyd, C.O. and Paavola, L.G. (1991) Origin of thylakoid membranes in Chlamydomonas reinhardtii y-1 at 38 degrees C. Plant Physiol.
96, 1321–1328.
Horwitz, L. (1957) Observations on the oxyhydrogen reaction in Scenedesmus and its relation to respiration and photosynthesis. Arch. Biochem.
Biophys. 66, 23–44.
Houchins, J.P. and Burris, R.H. (1981a) Light and dark reactions of the
uptake hydrogenase in Anabaena 7120. Plant Physiol. 68, 712–716.
Houchins, J.P. and Burris, R.H. (1981b) Occurrence and localization of two
distinct hydrogenases in the heterocystous cyanobacterium Anabaena
sp. strain 7120. J. Bacteriol. 146, 209–214.
Houille-Vernes, L., Rappaport, F., Wollman, F.A., Alric, J. and Johnson, X.
(2011) Plastid terminal oxidase 2 (PTOX2) is the major oxidase involved
in chlororespiration in Chlamydomonas. Proc. Natl Acad. Sci. USA, 108,
20820–20825.
Hughes, B.T. and Espenshade, P.J. (2008) Oxygen-regulated degradation of
fission yeast SREBP by Ofd1, a prolyl hydroxylase family member. EMBO
J. 27, 1491–1501.
Hughes, A.L., Todd, B.L. and Espenshade, P.J. (2005) SREBP pathway
responds to sterols and functions as an oxygen sensor in fission yeast.
Cell, 120, 831–842.
Iuchi, S. and Lin, E.C. (1992) Mutational analysis of signal transduction by
ArcB, a membrane sensor protein responsible for anaerobic repression
of operons involved in the central aerobic pathways in Escherichia coli.
J. Bacteriol. 174, 3972–3980.
Iwai, M., Takizawa, K., Tokutsu, R., Okamuro, A., Takahashi, Y. and Minagawa, J. (2010) Isolation of the elusive supercomplex that drives cyclic electron flow in photosynthesis. Nature, 464, 1210–1213.
Jacobs, J., Pudollek, S., Hemschemeier, A. and Happe, T. (2009) A novel,
anaerobically induced ferredoxin in Chlamydomonas reinhardtii. FEBS
Lett. 583, 325–329.
Jans, F., Mignolet, E., Houyoux, P.A., Cardol, P., Ghysels, B., Cuine, S.,
Cournac, L., Peltier, G., Remacle, C. and Franck, F. (2008) A type II NAD
(P)H dehydrogenase mediates light-independent plastoquinone reduction in the chloroplast of Chlamydomonas. Proc. Natl Acad. Sci. USA,
105, 20546–20551.
Jiang, Y., Chen, F. and Liang, S.Z. (1999) Production potential of docosahexaenoic acid by the heterotrophic marine dinoflagellate Crypthecodinium cohnii. Process Biochem. 34, 633.
Johnson, X. and Alric, J. (2013) Central carbon metabolism and electron
transport in Chlamydomonas reinhardtii: metabolic constraints for carbon partitioning between oil and starch. Eukaryot. Cell, 12, 776–793.
Johnson, X., Steinbeck, J., Dent, R.M. et al. (2014) Proton gradient regulation 5-mediated cyclic electron flow under ATP- or redox-limited conditions: a study of DeltaATpase pgr5 and DeltarbcL pgr5 mutants in the
green alga Chlamydomonas reinhardtii. Plant Physiol. 165, 438–452.
Joliot, P. and Joliot, A. (1980) Dependence of delayed luminescence upon
adenosine triphosphatase activity in Chlorella. Plant Physiol. 65, 691–696.
Joyard, J., Ferro, M., Masselon, C., Seigneurin-Berny, D., Salvi, D., Garin, J.
and Rolland, N. (2010) Chloroplast proteomics highlights the subcellular
compartmentation of lipid metabolism. Prog. Lipid Res. 48, 128–158.
Kamp, A., deBeer, D., Nitsch, J.C., Lavik, G. and Stief, P. (2011) Diatoms
respire nitrate to survive dark and anoxic conditions. Proc. Natl Acad.
Sci. USA, 108, 5649–5654.
Kato, M., Mizuno, T., Shimizu, T. and Hakoshima, T. (1997) Insights into
multistep phosphorelay from the crystal structure of the C-terminal HPt
domain of ArcB. Cell, 88, 717–723.
Kirch, H.H., Bartels, D., Wei, Y., Schnable, P.S. and Wood, A.J. (2004) The
ALDH gene superfamily of Arabidopsis. Trends Plant Sci. 9, 371–377.
Kirch, H.H., Schlingensiepen, S., Kotchoni, S., Sunkar, R. and Bartels, D.
(2005) Detailed expression analysis of selected genes of the aldehyde
dehydrogenase (ALDH) gene superfamily in Arabidopsis thaliana. Plant
Mol. Biol. 57, 315–332.
Klob, W., Kandler, O. and Tanner, W. (1973) The role of cyclic photophosphorylation in vivo. Plant Physiol. 51, 825–827.
Klock, G. and Kreuzberg, K. (1991) Compartmented metabolite pools in protoplasts from the green alga Chlamydomonas reinhardtii: changes after
transition from aerobiosis to anaerobiosis in the dark. Biochim. Biophys.
Acta, 1073, 410–415.
Kobayashi, M., Kakizono, T., Yamaguchi, K., Nishio, N. and Nagai, S. (1992)
Growth and astaxanthin formation of Haematococcus pluvialis in heterotrophic and mixotrophic conditions. J. Ferment. Bioeng. 74, 17–20.
Kornberg, H.L. and Krebs, H.A. (1957) Synthesis of cell constituents from
C2-units by a modified tricarboxylic acid cycle. Nature 179, 988–991.
Koshland, D.E. (1952) Effect of catalysts on the hydrolysis of acetyl phosphate. Nucleophilic displacement mechanisms in enzymatic reactions. J.
Am. Chem. Soc. 74, 2286–2292.
Kosourov, S.N. and Seibert, M. (2009) Hydrogen photoproduction by nutrient-deprived Chlamydomonas reinhardtii cells immobilized within thin
alginate films under aerobic and anaerobic conditions. Biotechnol. Bioeng. 102, 50–58.
Kreuzberg, K. (1984) Starch fermentation via formate producing pathway in
Chlamydomonas reinhardtii, Chlorogonium elongatum and Chlorella
fusca. Physiol. Plant. 61, 87–94.
€ ck, G. and Grobheiser, D. (1987) Subcellular distribution of
Kreuzberg, K., Klo
pyruvate-degrading enzymes in Chlamydomonas reinhardtii studied by an
improved protoplast fractionation procedure. Physiol. Plant. 69, 481–488.
Kro€ mer, S. (1995) Respiration during photosynthesis. Annu. Rev. Plant
Physiol. Plant Mol. Biol. 46, 45–70.
Kro€ mer, S. and Heldt, H.W. (1991) On the role of mitochondrial oxidative
phosphorylation in photosynthesis metabolism as studied by the effect
of oligomycin on photosynthesis in protoplasts and leaves of barley
(Hordeum vulgare). Plant Physiol. 95, 1270–1276.
Kruse, O., Rupprecht, J., Mussgnug, J., Dismukes, G. and Hankamer, B.
(2005a) Photosynthesis: a blueprint for solar energy capture and biohydrogen production technologies. Photochem. Photobiol. Sci. 4, 957–970.
Kruse, O., Rupprecht, J., Bader, K.P., Thomas-Hall, S., Schenk, P.M., Finazzi,
G. and Hankamer, B. (2005b) Improved photobiological H2 production in
engineered green algal cells. J. Biol. Chem. 280, 34170–34177.
Kuit, W., Minton, N.P., Lopez-Contreras, A.M. and Eggink, G. (2012) Disruption of the acetate kinase (ack) gene of Clostridium acetobutylicum
results in delayed acetate production. Appl. Microbiol. Biotechnol. 94,
729–741.
Kukuczka, B., Magneschi, L., Petroutsos, D., Steinbeck, J., Bald, T., Powikrowska, M., Fufezan, C., Finazzi, G. and Hippler, M. (2014) Proton gradient regulation5-like1-mediated cyclic electron flow is crucial for
acclimation to anoxia and complementary to nonphotochemical quenching in stress adaptation. Plant Physiol. 165, 1604–1617.
Kwast, K.E., Burke, P.V. and Poyton, R.O. (1998) Oxygen sensing and the
transcriptional regulation of oxygen-responsive genes in yeast. J. Exp.
Biol. 201, 1177–1195.
Lambertz, C., Hemschemeier, A. and Happe, T. (2010) Anaerobic expression
of the ferredoxin-encoding FDX5 gene of Chlamydomonas reinhardtii is
regulated by the Crr1 transcription factor. Eukaryot. Cell, 9, 1747–1754.
Lapaille, M., Thiry, M., Perez, E., Gonzalez-Halphen, D., Remacle, C. and Cardol, P. (2010) Loss of mitochondrial ATP synthase subunit beta (Atp2)
alters mitochondrial and chloroplastic function and morphology in Chlamydomonas. Biochim. Biophys. Acta, 1797, 1533–1539.
Lasanthi-Kudahettige, R., Magneschi, L., Loreti, E., Gonzali, S., Licausi, F.,
Novi, G., Beretta, O., Vitulli, F., Alpi, A. and Perata, P. (2007) Transcript
profiling of the anoxic rice coleoptile. Plant Physiol. 144, 218–231.
Lecler, R., Vigeolas, H., Emonds-Alt, B., Cardol, P. and Remacle, C. (2012)
Characterization of an internal type-II NADH dehydrogenase from Chlamydomonas reinhardtii mitochondria. Curr. Genet. 58, 205–216.
Lendzian, K. and Bassham, J.A. (1975) Regulation of glucose-6-phosphate
dehydrogenase in spinach chloroplasts by ribulose 1,5-diphosphate and
NADPH/NADP+ ratios. Biochim. Biophys. Acta, 396, 260–275.
Levine, P.P. (1969) The analysis of photosynthesis using mutant strains of
algae and higher plants. Annu. Rev. Plant Physiol. Plant Mol. Biol. 20,
523–540.
© 2015 The Authors
The Plant Journal © 2015 John Wiley & Sons Ltd, The Plant Journal, (2015), 82, 481–503
Dark hypoxic growth of algae 501
Li, X., Moellering, E.R., Liu, B., Johnny, C., Fedewa, M., Sears, B.B., Kuo,
M.H. and Benning, C. (2012) A galactoglycerolipid lipase is required for
triacylglycerol accumulation and survival following nitrogen deprivation
in Chlamydomonas reinhardtii. Plant Cell, 24, 4670–4686.
Licausi, F. (2011) Regulation of the molecular response to oxygen limitations in plants. New Phytol. 190, 550–555.
van Lis, R., Baffert, C., Coute, Y., Nitschke, W. and Atteia, A. (2013) Chlamydomonas reinhardtii chloroplasts contain a homodimeric pyruvate:ferredoxin
oxidoreductase that functions with FDX1. Plant Physiol. 161, 57–71.
Magneschi, L., Catalanotti, C., Subramanian, V., Dubini, A., Yang, W., Mus,
F., Posewitz, M.C., Seibert, M., Perata, P. and Grossman, A.R. (2012) A
mutant in the ADH1 gene of Chlamydomonas reinhardtii elicits metabolic
restructuring during anaerobiosis. Plant Physiol. 158, 1293–1305.
Maione, T.E. and Gibbs, M. (1986) Association of the chloroplastic respiratory and photosynthetic electron transport chains of Chlamydomonas reinhardii with photoreduction and the oxyhydrogen reaction. Plant
Physiol. 80, 364–368.
Martin, W. (2010) Evolutionary origins of metabolic compartmentalization in
eukaryotes. Philos. Trans. R. Soc. Lond. 365, 847–855.
Matagne, R.F., Michel-Wolwertz, M.R., Munaut, C., Duyckaerts, C. and
Sluse, F. (1989) Induction and characterization of mitochondrial DNA
mutants in Chlamydomonas reinhardtii. J. Cell Biol. 108, 1221–1226.
Melis, A. and Happe, T. (2001) Hydrogen production. Green algae as a
source of energy. Plant Physiol. 127, 740–748.
Melis, A. and Happe, T. (2004) Trails of green alga hydrogen research – from
Hans Gaffron to new frontiers. Photosyn. Res. 80, 401–409.
Merchant, S.S., Allen, M.D., Kropat, J., Moseley, J.L., Long, J.C., Tottey, S.
and Terauchi, A.M. (2006) Between a rock and a hard place: trace element
nutrition in Chlamydomonas. Biochim. Biophys. Acta, 1763, 578–594.
Merchant, S.S., Prochnik, S.E., Vallon, O. et al. (2007) The Chlamydomonas
genome reveals the evolution of key animal and plant functions. Science,
318, 245–250.
Meuser, J.E., D’Adamo, S., Jinkerson, R.E., Mus, F., Yang, W., Ghirardi,
M.L., Seibert, M., Grossman, A.R. and Posewitz, M.C. (2012) Genetic disruption of both Chlamydomonas reinhardtii [FeFe]-hydrogenases: Insight
into the role of HYDA2 in H2 production. Biochem. Biophys. Res. Commun. 417, 704–709.
Meyer, M. and Griffiths, H. (2013) Origins and diversity of eukaryotic CO2-concentrating mechanisms: lessons for the future. J. Exp. Bot. 64, 769–786.
Michaelis, G., Vahrenholz, C. and Pratje, E. (1990) Mitochondrial DNA of
Chlamydomonas reinhardtii: the gene for apocytochrome b and the complete functional map of the 15.8 kb DNA. Mol. Gen. Genet. 223, 211–216.
Millerioux, Y., Morand, P., Biran, M. et al. (2012) ATP synthesis-coupled and
-uncoupled acetate production from acetyl-CoA by mitochondrial acetate:succinate CoA-transferase and acetyl-CoA thioesterase in Trypanosoma. J. Biol. Chem. 287, 17186–17197.
Miyashita, Y. and Good, A.G. (2008) Contribution of the GABA shunt to
hypoxia-induced alanine accumulation in roots of Arabidopsis thaliana.
Plant Cell Physiol. 49, 92–102.
Monson, E.K., Weinstein, M., Ditta, G.S. and Helinski, D.R. (1992) The FixL
protein of Rhizobium meliloti can be separated into a heme-binding oxygen-sensing domain and a functional C-terminal kinase domain. Proc.
Natl Acad. Sci. USA, 89, 4280–4284.
Morsy, F.M. (2011) Acetate versus sulfur deprivation role in creating anaerobiosis in light for hydrogen production by Chlamydomonas reinhardtii
and Spirulina platensis: two different organisms and two different mechanisms. Photochem. Photobiol. 87, 137–142.
Moseley, J.L., Gonzalez-Ballester, D., Pootakham, W., Bailey, S. and Grossman, A.R. (2009) Genetic interactions between regulators of Chlamydomonas phosphorus and sulfur deprivation responses. Genetics, 181,
889–905.
€ ller, M. (1993) The hydrogenosome. J. Gen. Microbiol. 139, 2879–2889.
Mu
€ ller, M., Mentel, M., van Hellemond, J.J., Henze, K., Woehle, C., Gould,
Mu
S.B., Yu, R.Y., van der Giezen, M., Tielens, A.G. and Martin, W.F. (2012)
Biochemistry and evolution of anaerobic energy metabolism in eukaryotes. Microbiol. Mol. Biol. Rev. 76, 444–495.
Murthy, U.M., Wecker, M.S., Posewitz, M.C., Gilles-Gonzalez, M.A. and
Ghirardi, M.L. (2012) Novel FixL homologues in Chlamydomonas reinhardtii bind heme and O2. FEBS Lett. 586, 4282–4288.
Mus, F., Dubini, A., Seibert, M., Posewitz, M.C. and Grossman, A.R. (2007)
Anaerobic acclimation in Chlamydomonas reinhardtii: anoxic gene
expression, hydrogenase induction, and metabolic pathways. J. Biol.
Chem. 282, 25475–25486.
Nagel, G., Ollig, D., Fuhrmann, M., Kateriya, S., Musti, A.M., Bamberg, E.
and Hegemann, P. (2002) Channelrhodopsin-1: a light-gated proton channel in green algae. Science, 296, 2395–2398.
Neupert, W. (1997) Protein import into mitochondria. Annu. Rev. Biochem.
66, 863–917.
Noth, J., Krawietz, D., Hemschemeier, A. and Happe, T. (2013) Pyruvate:ferredoxin oxidoreductase is coupled to light-independent hydrogen production in Chlamydomonas reinhardtii. J. Biol. Chem. 288, 4368–4377.
Ogbonna, J.C., Tomiyama, S. and Tanaka, H. (1998) Heterotrophic cultivation of Euglena gracilis Z for efficient production of a-tocopherol. J. Appl.
Phycol. 10, 67–74.
Ohad, I., Siekevitz, P. and Palade, G.E. (1967) Biogenesis of chloroplast
membranes II. Plastid differentiation during greening of a dark grown
algal mutant (Chlamydomonas reinharddi). J. Cell Biol. 35, 553–584.
Ohta, S., Miyamoto, K. and Miura, Y. (1987) Hydrogen evolution as a consumption mode of reducing equivalents in green algal fermentation.
Plant Physiol. 83, 1022–1026.
Olausson, T., Nordling, M., Karlsson, G., Meuller, J., Lundberg, L. and Rydstrom, J. (1992) Structure and function of proton-translocating nicotinamide nucleotide transhydrogenase. Acta Physiol. Scand. 607, 13–22.
Page, M.D., Kropat, J., Hamel, P.P. and Merchant, S.S. (2009) Two Chlamydomonas CTR copper transporters with a novel cys-met motif are
localized to the plasma membrane and function in copper assimilation.
Plant Cell, 21, 928–943.
Paiva, S., Devaux, F., Barbosa, S., Jacq, C. and Casal, M. (2004) Ady2p is
essential for the acetate permease activity in the yeast Saccharomyces
cerevisiae. Yeast, 21, 201–210.
Palkova, Z., Devaux, F., Icicova, M., Minarikova, L., Le Crom, S. and Jacq, C.
(2002) Ammonia pulses and metabolic oscillations guide yeast colony
development. Mol. Biol. Cell, 13, 3901–3914.
Pape, M., Lambertz, C., Happe, T. and Hemschemeier, A. (2012) Differential
expression of the Chlamydomonas [FeFe]-hydrogenase-encoding HYDA1
gene is regulated by the copper response regulator1. Plant Physiol. 159,
1700–1712.
Pedersen, A., Karlsson, G.B. and Rydstrom, J. (2008) Proton-translocating
transhydrogenase: an update of unsolved and controversial issues. J.
Bioenerg. Biomembr. 40, 463–473.
Peltier, G. and Cournac, L. (2002) Chlororespiration. Annu. Rev. Plant Biol.
53, 523–550.
Perez-Garcia, O., Escalante, F.M., de-Bashan, L.E. and Bashan, Y. (2011) Heterotrophic cultures of microalgae: metabolism and potential products.
Water Res. 45, 11–36.
Petroutsos, D., Terauchi, A.M., Busch, A., Hirschmann, I., Merchant, S.S., Finazzi, G. and Hippler, M. (2009) PGRL1 participates in iron-induced
remodeling of the photosynthetic apparatus and in energy metabolism
in Chlamydomonas reinhardtii. J. Biol. Chem. 284, 32770–32781.
Pfannschmidt, T. and Liere, K. (2005) Redox regulation and modification of
proteins controlling chloroplast gene expression. Antioxid. Redox Signal.
7, 607–618.
Philipps, G., Krawietz, D., Hemschemeier, A. and Happe, T. (2011) A pyruvate formate lyase-deficient Chlamydomonas reinhardtii strain provides
evidence for a link between fermentation and hydrogen production in
green algae. Plant J. 66, 330–340.
Plancke, C., Vigeolas, H., Hohner, R. et al. (2014) Lack of isocitrate lyase in Chlamydomonas leads to changes in carbon metabolism and in the response to
oxidative stress under mixotrophic growth. Plant J. 77, 404–417.
Plaxton, W.C. (1996) The organization and regulation of plant glycolysis.
Annu. Rev. Plant Physiol. Plant Mol. Biol. 45, 185–214.
Pootakham, W., Gonzalez-Ballester, D. and Grossman, A.R. (2010) Identification and regulation of plasma membrane sulfate transporters in Chlamydomonas. Plant Physiol. 153, 1653–1668.
Posewitz, M.C., King, P.W., Smolinski, S.L., Zhang, L., Seibert, M. and Ghirardi, M.L. (2004a) Discovery of two novel radical S-adenosylmethionine
proteins required for the assembly of an active [Fe] hydrogenase. J. Biol.
Chem. 279, 25711–25720.
Posewitz, M.C., Smolinski, S.L., Kanakagiri, S., Melis, A., Seibert, M. and
Ghirardi, M.L. (2004b) Hydrogen photoproduction is attenuated by disruption of an isoamylase gene in Chlamydomonas reinhardtii. Plant Cell,
16, 2151–2163.
© 2015 The Authors
The Plant Journal © 2015 John Wiley & Sons Ltd, The Plant Journal, (2015), 82, 481–503
502 Wenqiang Yang et al.
Poyton, R.O. (1999) Models for oxygen sensing in yeast: implications for
oxygen-regulated gene expression in higher eukaryotes. Respir. Physiol.
115, 119–133.
Purton, S. (2007) Tools and techniques for chloroplast transformation of
Chlamydomonas. Adv. Exp. Med. Biol. 616, 34–45.
Remacle, C., Baurain, D., Cardol, P. and Matagne, R.F. (2001a) Mutants of
Chlamydomonas reinhardtii deficient in mitochondrial complex I: characterization of two mutations affecting the nd1 coding sequence. Genetics,
158, 1051–1060.
Remacle, C., Duby, F., Cardol, P. and Matagne, R.F. (2001b) Mutations inactivating mitochondrial genes in Chlamydomonas reinhardtii. Biochem.
Soc. Trans. 29, 442–446.
Remacle, C., Cardol, P., Coosemans, N., Gaisne, M. and Bonnefoy, N. (2006)
High-efficiency biolistic transformation of Chlamydomonas mitochondria
can be used to insert mutations in complex I genes. Proc. Natl Acad. Sci.
USA, 103, 4771–4776.
Remacle, C., Coosemans, N., Jans, F., Hanikenne, M., Motte, P. and Cardol,
P. (2010) Knock-down of the COX3 and COX17 gene expression of cytochrome c oxidase in the unicellular green alga Chlamydomonas reinhardtii. Plant Mol. Biol. 74, 223–233.
Remize, F., Andrieu, E. and Dequin, S. (2000) Engineering of the pyruvate
dehydrogenase bypass in Saccharomyces cerevisiae: role of the cytosolic
Mg2+ and mitochondrial K+ acetaldehyde dehydrogenases Ald6p and
Ald4p in acetate formation during alcoholic fermentation. Appl. Environ.
Microbiol. 66, 3151–3159.
Robellet, X., Flipphi, M., Pegot, S., Maccabe, A.P. and Velot, C. (2008) AcpA,
a member of the GPR1/FUN34/YaaH membrane protein family, is essential for acetate permease activity in the hyphal fungus Aspergillus nidulans. Biochem. J. 412, 485–493.
Roberts, J.K., Chang, K., Webster, C., Callis, J. and Walbot, V. (1989) Dependence of ethanolic fermentation, cytoplasmic pH regulation, and viability
on the activity of alcohol dehydrogenase in hypoxic maize root tips.
Plant Physiol. 89, 1275–1278.
Rochaix, J.D. (2001) Assembly, function, and dynamics of the photosynthetic
machinery in Chlamydomonas reinhardtii. Plant Physiol. 127, 1394–1398.
Rochaix, J.D. (2007) Role of thylakoid protein kinases in photosynthetic
acclimation. FEBS Lett. 581, 2768–2775.
Rupprecht, J. (2009) From systems biology to fuel–Chlamydomonas reinhardtii as a model for a systems biology approach to improve biohydrogen production. J. Biotech. 142, 10–20.
Russell, G.K. and Gibbs, M. (1968) Evidence for the participation of the
reductive pentose phosphate cycle in photoreduction and the oxyhydrogen reaction. Plant Physiol. 43, 649–652.
Rutter, J., Reick, M., Wu, L.C. and McKnight, S.L. (2001) Regulation of clock
and NPAS2 DNA binding by the redox state of NAD cofactors. Science,
293, 510–514.
Sager, R. (1955) Inheritance in the Green Alga Chlamydomonas reinhardii.
Genetics, 40, 476–489.
Salinas, T., Larosa, V., Cardol, P., Marechal-Drouard, L. and Remacle, C.
(2014) Respiratory-deficient mutants of the unicellular green alga Chlamydomonas: a review. Biochimie, 100, 207–218.
Sasidharan, R. and Mustroph, A. (2011) Plant oxygen sensing is mediated
by the N-end rule pathway: a milestone in plant anaerobiosis. Plant Cell,
23, 4173–4183.
Sauer, U., Canonaco, F., Heri, S., Perrenoud, A. and Fischer, E. (2004) The
soluble and membrane-bound transhydrogenases UdhA and PntAB have
divergent functions in NADPH metabolism of Escherichia coli. J. Biol.
Chem. 279, 6613–6619.
Scheibe, R. (1987) NADP+-malate dehydrogenase in C3-plants: regulation
and role of a light-activated enzyme. Physiol. Plant. 71, 393–400.
Semenza, G.L. (2011) Oxygen sensing, homeostasis and disease. N. Engl. J.
Med. 365, 537–547.
Sillers, R., Chow, A., Tracy, B. and Papoutsakis, E.T. (2008) Metabolic engineering of the non-sporulating, non-solventogenic Clostridium acetobutylicum strain M5 to produce butanol without acetone demonstrate the
robustness of the acid-formation pathways and the importance of the
electron balance. Metabol. Engin. 10, 321–332.
Sommer, F., Kropat, J., Malasarn, D., Grossoehme, N.E., Chen, X., Giedroc,
D.P. and Merchant, S.S. (2010) The CRR1 nutritional copper sensor in
Chlamydomonas contains two distinct metal-responsive domains. Plant
cell, 22, 4098–4113.
Spear, J.R., Walker, J.J., McCollom, T.M. and Pace, N.R. (2005) Hydrogen
and bioenergetics in the Yellowstone geothermal ecosystem. Proc. Natl
Acad. Sci. USA, 102, 2555–2560.
Spiro, S. and Guest, J.R. (1991) Adaptive responses to oxygen limitation in
Escherichia coli. Trends Biochem. Sci. 16, 310–314.
Stirnberg, M. and Happe, T. (2004) Identification of a cis-acting element
controlling anaerobic expression of the HydA gene from Chlamydomonas reinhardtii. In Biohydrogen III (Miyake, J., Igarashi, Y. and Roegner, M., eds). Oxford: Elsevier Science, pp. 117–127.
Takahashi, H., Clowez, S., Wollman, F.A., Vallon, O. and Rappaport, F.
(2013) Cyclic electron flow is redox-controlled but independent of state
transition. Nat. Commun. 4, 1954.
Takasaki, K., Shoun, H., Yamaguchi, M., Takeo, K., Nakamura, A., Hoshino,
T. and Takaya, N. (2004) Fungal ammonia fermentation, a novel metabolic mechanism that couples the dissimilatory and assimilatory pathways of both nitrate and ethanol. Role of acetyl CoA synthetase in
anaerobic ATP synthesis. J. Biol. Chem. 279, 12414–12420.
Terashima, M., Specht, M., Naumann, B. and Hippler, M. (2010) Characterizing the anaerobic response of Chlamydomonas reinhardtii by quantitative proteomics. Mol. Cell Proteomics, 9, 1514–1532.
Terashima, M., Specht, M. and Hippler, M. (2011) The chloroplast proteome:
a survey from the Chlamydomonas reinhardtii perspective with a focus
on distinctive features. Curr. Genet. 57, 151–168.
€ dig, M. et al. (2012) Calcium-dependent
Terashima, M., Petroutsos, D., Hu
regulation of cyclic photosynthetic electron transfer by a CAS, ANR1,
and PGRL1 complex. Proc. Natl Acad. Sci. USA, 109, 17717–17722.
Terauchi, A.M., Lu, S.F., Zaffagnini, M. et al. (2009) Pattern of expression
and substrate specificity of chloroplast ferredoxins from Chlamydomonas reinhardtii. J. Biol. Chem. 284, 25867–25878.
Tielens, A.G., van Grinsven, K.W., Henze, K., van Hellemond, J.J. and Martin, W. (2010) Acetate formation in the energy metabolism of parasitic
helminths and protists. Int. J. Parasitol. 40, 387–397.
Timmins, M., Thomas-Hall, S.R., Darling, A., Zhang, E., Hankamer, B., Marx,
U.C. and Schenk, P.M. (2009) Phylogenetic and molecular analysis of
hydrogen-producing green algae. J. Exp. Bot. 60, 1691–1702.
Todd, B.L., Stewart, E.V., Burg, J.S., Hughes, A.L. and Espenshade, P.J.
(2006) Sterol regulatory element binding protein is a principal regulator of
anaerobic gene expression in fission yeast. Mol. Cell. Biol., 26, 2817–2831.
Tolleter, D., Ghysels, B., Alric, J. et al. (2011) Control of hydrogen photoproduction by the proton gradient generated by cyclic electron flow in Chlamydomonas reinhardtii. Plant Cell, 23, 2619–2630.
Tsygankov, A.A., Fedorov, A.S., Kosourov, S.N. and Rao, K.K. (2002) Hydrogen production by cyanobacteria in an automated outdoor photobioreactor under aerobic conditions. Biotech. Bioeng. 80, 777–783.
Vlad, F., Spano, T., Vlad, D., Daher, F.B., Ouelhadj, A. and Kalaitzis, P.
(2007) Arabidopsis prolyl 4-hydroxylases are differentially expressed in
response to hypoxia, anoxia and mechanical wounding. Physiol. Plant.
130, 471–483.
Wagner, V., Ullmann, K., Mollwo, A., Kaminski, M., Mittag, M. and Kreimer,
G. (2008) The phosphoproteome of a Chlamydomonas reinhardtii eyespot fraction includes key proteins of the light signaling pathway. Plant
Physiol. 146, 772–788.
Wang, W.Y., Wang, W.L., Boynton, J.E. and Gillham, N.W. (1974) Genetic
control Of chlorophyll biosynthesis in chlamydomonas analysis of
mutants at two loci mediating the conversion of protoporphyrin-IX to
magnesium protoporphyrin. J Cell Biol., 63, 806–823.
Weber, A., Menzlaff, E., Arbinger, B., Gutensohn, M., Eckerskorn, C. and
Flugge, U.I. (1995) The 2-oxoglutarate/malate translocator of chloroplast
envelope membranes: molecular cloning of a transporter containing a
12-helix motif and expression of the functional protein in yeast cells. Biochemistry, 34, 2621–2627.
Wei, L., Derrien, B., Gautier, A. et al. (2014) Nitric oxide-triggered remodeling of chloroplast bioenergetics and thylakoid proteins upon nitrogen
starvation in Chlamydomonas reinhardtii. Plant Cell, 26, 353–372.
Weigelt, K., Kuster, H., Rutten, T. et al. (2009) ADP-glucose pyrophosphorylase-deficient pea embryos reveal specific transcriptional and metabolic
changes of carbon-nitrogen metabolism and stress responses. Plant
Physiol. 149, 395–411.
Wen, Z.-Y. and Chen, F. (2003) Heterotrophic production of eicosapentaenoic acid by microalgae. Biotechnol. Adv. 21, 273–294.
© 2015 The Authors
The Plant Journal © 2015 John Wiley & Sons Ltd, The Plant Journal, (2015), 82, 481–503
Dark hypoxic growth of algae 503
White, R.A. and Hoober, J.K. (1994) Biogenesis of thylakoid membranes in
Chlamydomonas reinhardtii y1 (a kinetic study of initial greening). Plant
Physiol. 106, 583–590.
Whitney, L.A., Loreti, E., Alpi, A. and Perata, P. (2011) Alcohol dehydrogenase and hydrogenase transcript fluctuations during a day-night cycle in
Chlamydomonas reinhardtii: the role of anoxia. New Phytol. 190, 488–
498.
Wieckowski, M.R. and Wojtczak, L. (1997) Involvement of the dicarboxylate
carrier in the protonophoric action of long-chain fatty acids in mitochondria. Biochem. Biophys. Res. Commun. 232, 414–417.
Wiessner, W. (1965) Quantum requirement for acetate assimilation and its
significance for quantum measurements in photophosphorylation. Nature, 205, 56–57.
Wiseman, A., Gillham, N.W. and Boynton, J.E. (1977) Nuclear mutations
affecting mitochondrial structure and function in Chlamydomonas. J.
Cell Biol. 73, 56–77.
Yang, W., Catalanotti, C., Posewitz, M., Alric, J. and Grossman, A.R. (2014a)
Insight into Algal Fermentation Metabolism. Springer: Low Oxygen
Stress in Plants, 21, pp. 35–163.
Yang, W., Catalanotti, C., D’Adamo, S. et al. (2014b) Alternative acetate production pathways in Chlamydomonas reinhardtii during dark anoxia and
the dominant role of chloroplasts in fermentative acetate production.
Plant Cell, 26, 4499–4518.
Yoshii, Y., Furukawa, T., Yoshii, H. et al. (2009) Cytosolic acetyl-CoA synthetase affected tumor cell survival under hypoxia: the possible function in
tumor acetyl-CoA/acetate metabolism. Cancer Sci. 100, 821–827.
Zabawinski, C., Van Den Koornhuyse, N., D’Hulst, C., Schlichting, R., Giersch, C., Delrue, B., Lacroix, J.M., Preiss, J. and Ball, S. (2001) Starchless
mutants of Chlamydomonas reinhardtii lack the small subunit of a heterotetrameric ADP-glucose pyrophosphorylase. J. Bacteriol. 183, 1069–1077.
Zeng, Y., Wu, Y., Avigne, W.T. and Koch, K.E. (1999) Rapid repression of
maize invertases by low oxygen. Invertase/sucrose synthase balance,
sugar signaling potential, and seedling survival. Plant Physiol. 121, 599–
608.
Zhou, Z., Takaya, N., Nakamura, A., Yamaguchi, M., Takeo, K. and Shoun,
H. (2002) Ammonia fermentation, a novel anoxic metabolism of nitrate
by fungi. J. Biol. Chem. 277, 1892–1896.
© 2015 The Authors
The Plant Journal © 2015 John Wiley & Sons Ltd, The Plant Journal, (2015), 82, 481–503