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Plant Physiology Preview. Published on October 23, 2017, as DOI:10.1104/pp.17.01512
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Research Area: Biochemistry and Metabolism
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Short title: Lipid droplet localization
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*Corresponding author: Naoki Sato (naokisat@bio.c.u-tokyo.ac.jp)
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Department of Life Sciences, Graduate School of Arts and Sciences, University of Tokyo, Tokyo
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153-8902, Japan
Tel: +81-3-5454-6631
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Title :
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Revisiting the algal “chloroplast lipid droplet”: the absence of an entity that is unlikely to
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exist
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Authors:
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Takashi Moriyama1,2, Masakazu Toyoshima1,2,3, Masakazu Saito1,2,4, Hajime Wada1,2 and Naoki
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Sato1,2,*
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Affiliations:
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1
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Department of Life Sciences, Graduate School of Arts and Sciences, University of Tokyo, Tokyo
153-8902, Japan
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CREST, Japan Science and Technology Agency, Tokyo 102-0076, Japan
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One sentence summary
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Despite previous arguments on “chloroplast lipid droplet”, all lipid droplets are present in the
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cytosolic compartment and not in the chloroplast in Chlamydomonas reinhardtii.
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Author contributions
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N.S. conceived research; T.M., M.T, H.W. and N.S. designed research; T.M., M.T., M.S. and N.S.
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performed research (culture, observation, and image processing); T.M., M.T., H.W. and N.S. wrote
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the manuscript. All authors agreed to publish this manuscript.
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Funding information
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This work was supported in part by a KAKENHI (15K12433 and 17H03715 to NS) from the Japan
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Society for the Promotion of Science (JSPS), and a grant, Core Research for Evolutional Science and
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Technology, from Japan Science and Technology Agency to NS and HW.
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and Technology, Osaka University, 1-5 Yamadaoka, Suita, Osaka 565-0871, Japan
Present address: Department of Bioinformatic Engineering, Graduate School of Information Science
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Copyright 2017 by the American Society of Plant Biologists
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Present address: Department of Mathematical and Life Sciences, Graduate School of Science,
Hiroshima University, Higashi-Hiroshima 739-8526, Japan
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Abstract (233 words < 250)
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The precise localization of the lipid droplets and the metabolic pathways associated with the oil
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production are crucial to the engineering of microalgae for biofuel production. Several studies have
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reported detecting lipid droplets within the chloroplast of the microalga Chlamydomonas reinhardtii
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(Chlamydomonas), which accumulates considerable amount of triacylglycerol (TAG) and starch
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within the cell under nitrogen deprivation or high-light stress conditions. Starch undoubtedly
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accumulates within the chloroplast, but there have been debates on the localization of the lipid
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droplets, which are cytosolic organelles in other organisms. Although it is impossible to deny what
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never existed, we tried to repeat the experiments that pretended to find “chloroplast lipid droplets”.
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Here, we present microscopic results showing no evidence for the presence of lipid droplets within
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the chloroplast stroma, even though some lipid droplets existed in close association with the
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chloroplast or were even largely engulfed by the chloroplasts. Therefore, lipid droplets are cytosolic
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structures, distinct from the plastoglobules present in the chloroplast stroma. These results not only
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contrast with the old ideas, but also point out that the presumptive “chloroplast lipid droplets” are,
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in fact, lipid droplets embedded within chloroplast invaginations in association with the outer
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envelope of the chloroplast without intervention of the endoplasmic reticulum. This points to an
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intriguing possibility of a tight metabolic flow from the chloroplast to the lipid droplet through a
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close association rather than direct contact of both organelles.
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Key Words: Chlamydomonas reinhardtii, Chloroplast, Confocal microscopy, Electron microscopy,
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Lipid droplet.
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Introduction
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« Il est impossible de prouver la négative », answered Louis Pasteur (1922) to the partisans of the
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spontaneous generation. This famous phrase was meant to express the theoretical impossibility of
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denying what never exists. The same argument seems to be valid for the current debate on the
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presumptive presence of the lipid droplets within the chloroplast in the microalga Chlamydomonas
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reinhardtii (Chlamydomonas), a model alga that accumulates a considerable amount of
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triacylglycerol (TAG) and starch within the cell under nitrogen-deprived or high-light stress
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conditions (Siaut et al., 2011). Starch undoubtedly accumulates within the chloroplast, but there
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have been arguments on the localization of the lipid droplets, which are normally cytosolic
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organelles in plants and other organisms. Three studies on the presence of lipid droplets in the
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chloroplast have reported two lines of evidence: one based on starchless mutants under nitrogen
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deficient conditions (Goodson et al., 2011; Fan et al., 2011) and the other based on wild-type cells
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under high-light stress (Goold et al., 2016). Localization of the lipid droplets is crucial to
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engineering microalgae for biofuel production, which relies on the biochemical and cytological
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understanding of TAG synthesis and accumulation (Merchant et al., 2012; Johnson and Alric, 2013;
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Li-Beisson et al., 2015). Thus we sought to resolve the controversy by carefully repeating the
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previous electron microscopy experiments and using the confocal three-dimensional image
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reconstruction.
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The lipid droplet is a globular organelle enclosing TAG as a major constituent and is
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covered by a half unit membrane consisting of phospholipids and specific proteins. A homologous
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organelle is found in plants and algae, as well as in non-photosynthetic eukaryotes. In plants, the
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lipid droplet is known to emerge from the endoplasmic reticulum (ER) by budding. Many protein
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components of the lipid droplet membrane are known in plants and include oleosin, caleosin, and
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sterol dehydrogenase (Chapman et al., 2012), which are believed to stabilize the lipid droplet
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structure. In Chlamydomonas, different kinds of proteins are present in the limiting membrane of
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the lipid droplet (Moellering & Benning, 2010; Nguyen et al., 2011; Tsai et al., 2015). Lipid
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droplets can be small (<200 nm in diameter) or large (up to 1 µm in diameter). A number of
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microalgal biofuel production studies have focused on increasing oil production. TAG, a major
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component of the lipid droplet, is synthesized in the ER and the surface of the lipid droplet, with
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diacylglycerol and acyl-CoA or phosphatidylcholine as major precursors (Bates, 2016; Zienkiewicz
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et al., 2016). The pathway of TAG synthesis in plants and microalgae is supposed to be similar to
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that in animal cells and yeast with some additional pathways (Merchant et al., 2012; Johnson and
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Alric, 2013; Li-Beisson et al., 2015; Bates, 2016; Zienkiewicz et al., 2016).
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The chloroplast is the major site of fatty acid synthesis in plants and algae. Some of the
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fatty acids are transported to the cytosolic compartment and used for synthesis of phospholipids in
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the ER, whereas the remaining fatty acids are used to synthesize chloroplast lipids, such as
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galactolipids (Li-Beisson et al., 2015). Under nitrogen deprived conditions, fatty acids are
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mobilized from galactolipids to synthesize TAG (Zienkiewicz et al., 2016; Li et al., 2012). This
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raises a naïve suggestion that TAG could also be synthesized and accumulated within the
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chloroplast (Goodson et al., 2011; Fan et al., 2011). An argument for this comes from the presence
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of TAG in the plastoglobule, another type of lipid-containing structure that is ubiquitously present
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in the chloroplast (Bréhélin et al., 2007; Lohscheider and Bártulos, 2016). Cyanobacteria, which are
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considered descendants of an ancestral endosymbiont that engendered chloroplasts, also contain a
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lipidic structure apparently similar to plastoglobules (van de Meene et al., 2006). The plastoglobule
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is a small globular particle (100–200 nm in diameter) present in the vicinity of the photosynthetic
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membranes. It contains various isoprenoids, such as plastoquinones and tocopherols, but also TAG
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and membrane lipids (Tevini and Steinmüller, 1985). The enzyme, farnesyl ester synthetase
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(FES1/2), has the ability to synthesize TAG in chloroplasts (Lippold et al., 2012).
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Localization of lipid droplets has been analyzed in detail by freeze-fracture electron
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microscopy in Chlamydomonas (Goodson et al., 2011). That study affirmed that the lipid droplet is
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present (A) in the cytosol between the nucleus and the chloroplast, (B) in the cytosol outside the
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chloroplast, or (C) within the chloroplast stroma (Fig. 1 panels A–C). The first two locations are
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commonly found in oil-accumulating Chlamydomonas, and this is consistent with the presence of
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lipid droplets within the cytosol in animal cells and yeast. The third location is reportedly found
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only in starchless mutants under “acetate-boost” conditions. This type of lipid droplet is called
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“cpst-LB” (for “chloroplast lipid body”). Another study that also reported lipid droplets within the
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chloroplast examined ultrathin sections by transmission electron microscopy (Fan et al., 2011). A
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more recent study argues that “plastidal lipid droplets” accumulate in the chloroplast under
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saturating light stress (Goold et al., 2016).
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The presence of lipid droplets in the chloroplast is currently being discussed by the lipid
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research community. There are several reasons to cast doubt about type C localization: First, all
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known enzymes (except FES1/2) involved in TAG synthesis are targeted to the ER. Second, there is
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no solid evidence that cyanobacteria have the ability to accumulate TAG as lipid droplets, such as is
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seen in eukaryotes. In addition, the lipid droplet and plastoglobule are different entities that must be
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distinguished.
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The present study was undertaken in an effort to find evidence for type C localization.
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According to the methodology of Louis Pasteur in denying spontaneous generation, it was
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important for us to repeat the previous experiments that showed type C localization with great care.
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If we obtained a positive result for the presence of lipid droplets within the chloroplast, this could
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be useful in algal engineering (Bhowmick et al., 2016; Gargouri et al., 2015; Misra et al., 2012).
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Even if we could not confirm the previous results, this would not mean that the type C localization
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never existed. Type C localization could have been present in other experiments under different
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conditions, at least theoretically. Nevertheless, if we continued to work in this manner, and finally
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did not find positive results for type C localization, we would be forced to make a decision to deny
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the reality of type C localization. Here, we present various relevant observations which lead us to be
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quite doubtful about type C localization; however, we did notice an intriguing possibility of direct
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metabolic flow between the chloroplast and lipid droplets.
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RESULTS
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High-light intensity-induced accumulation of lipid droplets in wild-type cells
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To confirm that the wild-type cells accumulate neutral lipids in response to high-light intensity as
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reported in strain CC-124 (Goold et al., 2016), we used strain CC-1010 (Sakurai et al., 2014) and
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examined lipid droplets stained with BODIPY after a shift from low-light (40 μmol m–2 s–1) to
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high-light (200 μmol m–2 s–1) (Fig. 2 panel A). Lipid droplet accumulation was estimated by
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quantifying Nile Red fluorescence (Fig. 2 panel B). The results showed that the fluorescence signals
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of the lipid droplets increased linearly with time under the high-light condition. As far as the
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microscopic images indicated, the lipid droplets detected corresponded to the “plastidal lipid
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droplets” reported by Goold et al. (2016), with respect to their size and the distribution within the
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cell. Quantification of TAG by chemical analysis (Table 1) gave consistent results with those of
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fluorometry, which revealed that CC-1010 cells also accumulated TAG upon transfer to the
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high-light condition, and that the continuous culture system used by Goold et al. (2016) was not
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obligatory for this phenomenon. The TAG contained only a low level of 16:4 and 18:3(9,12,15)
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fatty acids, suggesting that the contribution of chloroplast membrane lipids to the TAG pool was not
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important under this condition in contrast to the nitrogen-deficient conditions reported previously
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(Li et al., 2012; Sakurai et al., 2014).
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Subcellular localization of lipid droplets in cells 7–11 h after transfer to high light was
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examined by Z-stacking confocal fluorescence microscopy after staining with BODIPY or
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LipidTOX. The BODIPY and LipidTOX fluorescence signals coincided with each other (Fig. S1).
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The great majority of lipid droplets appeared in the cytosolic compartment in the two-dimensional
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images. In rare cases, the lipid droplets appeared entirely embedded in the chloroplast (Fig. 3 panels
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A–C). Note that the chloroplast images were obtained from blue autofluorescence (shown in red
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pseudocolor) and not from chlorophyll fluorescence. This was useful for avoiding contamination of
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lipid droplet fluorescence in the chloroplast image (for details, see Materials and Methods). The
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lipid droplets found in the cytosol and those apparent within the chloroplast were similar in size and
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appearance. We obtained the Z-stacking images containing such lipid droplets at intervals of 0.2–
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0.4 μm and carefully examined the structure of the chloroplast and lipid droplets in the three
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dimensional (3D) image with clipping of the X-, Y-, or Z-axes. We show three cases as examples in
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which the lipid droplets seemed to be present in the chloroplast. In Case 1 (Fig. 3) and Case 2 (Fig.
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S2), the lipid droplets indicated by arrows/arrowheads were present within an invagination of the
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chloroplast (Fig. 3 panels D–I and Fig. S2 panels D–F), but appeared entirely embedded within the
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chloroplast depending on the angle (Fig. 3 and Fig. S2, panels A–C). Case 3 (Fig. S3) shows that
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the lipid droplet indicated by the arrowhead penetrated the chloroplast, whereas another lipid
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droplet (arrow) was located within an invagination of the chloroplast. Sequential tiff files for
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rendering the view and selected movie files of the stereo images with rolling are provided as
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Supplemental Movies.
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Examination by transmission electron microscopy also revealed the infrequent occurrence
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of lipid droplets that appeared to be within the chloroplast. The presence of chloroplast envelope
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membranes, which separated the inside and outside of the chloroplast, is a good criterion to judge if
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a lipid droplet is located within or outside a chloroplast. Because the thickness of the thin sections
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was about ten times larger than the thickness of the chloroplast envelope membranes, the
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membranes were not always seen clearly (Fig. 1 panel D). Proper tilting of the section in the
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microscope made it possible to clearly trace the envelope membranes (Fig. 1 panel E). Figure 4
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panel A shows an example of a small lipid droplet apparently located within a chloroplast; however,
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an invagination was visualized by tilting (Fig. 4 panels B and C). This lipid droplet seemed to have
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a thin limiting membrane (Fig. 4 panel D). In addition, the lipid droplet was clearly different from
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the plastoglobules, which were about 100-nm particles present in the stroma in close association
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with starch granules and thylakoid membranes (Fig. 4 panel E). Although we examined many
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samples by confocal fluorescence and electron microscopy, we were unable to find any lipid droplet
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entirely enclosed within a chloroplast. In other words, all lipid droplets were present within the
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cytosolic compartment under high-light conditions.
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Starchless mutants under nitrogen deprivation
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We then examined subcellular localization of the lipid droplets in the starchless mutants cw15sta6
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and cw15sta7 (Siaut et al., 2011; Goodson et al., 2011; Fan et al., 2011; Work et al., 2010), which
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accumulate TAG after an acetate-boost under nitrogen deprivation. Figure 5 panel A shows an
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electron microscope image of a representative cw15sta6 cell containing lipid droplets, some of
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which were apparently present in the chloroplast (two lipid droplets were fused in this case). Tilted
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images of the lipid droplets (Fig. 5 panels B–D) allowed us to trace the envelope membranes
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enclosing the chloroplasts (Fig. 5 orange lines in panels E–G). A careful examination by connecting
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all traces revealed that the lipid droplets contacted with the cytosol at the sites indicated by the
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arrows. Figure S4 shows the lipid droplets in cw15sta7 cells. It was clear that the lipid droplets were
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present in close association with the chloroplast, but they existed outside the chloroplast and were
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never entirely embedded in the chloroplast stroma.
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We also examined starchless mutants by confocal microscopy and 3D image
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reconstruction (Fig. 6). In this case, the cells were stained with visualizing reagents for lipid and
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cytosol without fixing the cells. Cytosolic staining with fluorescein diacetate (FDA) was possible
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only in living cells, because the staining depends on the cytosolic esterase that hydrolyzes FDA.
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Some lipid droplets indicated by the arrows were apparently located within the chloroplast, but
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clipping at appropriate slices showed that they were present within the cytosolic compartment,
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although they were deeply embedded within the invaginations. As in the case of lipid droplets that
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accumulate under high light, we never found any lipid droplets present in the chloroplast in the
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starchless mutants.
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DISCUSSION
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Resolution of type C localization
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We were finally led to the idea that it is impossible to prove what never exists, as Pasteur thought
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about 150 years ago with respect to spontaneous generation (Pasteur 1922: original version was
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published in the 1860s). He thought that it was impossible to completely deny all possibilities of
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spontaneous generation. However, he inspected all of the experiments of the opponents and
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removed all possible pitfalls therein. We took the same strategy in our study on the presumptive
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lipid droplets in the chloroplast. We found various conditions in which lipid droplets were
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apparently present in the chloroplast (type C localization). We then carefully examined these lipid
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droplets by focusing on the presence of the chloroplast envelope membranes and the cytosol. All of
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our observations led us to conclude that the lipid droplets were not present within the chloroplast
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stroma, either under the high-light condition or under nitrogen deprivation.
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As Pasteur did for the spontaneous generation, we re-examined the results of previously
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published studies on type C localization of lipid droplets. Unfortunately, all previously presented
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data do not support type C localization. The lipid droplets in the fracture face reported by Goodson
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et al. (2011: Fig. 11A) were clearly enclosed by double membranes, which were unambiguously
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identified as envelope membranes; in other words, they were located outside the chloroplast. The
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lipid droplets reported by Fan et al. (2011) were present in a chloroplast invagination but not
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entirely enclosed by a chloroplast (Fig. S5 panel A). In the fluorescence micrographs of a more
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recent report (Goold et al., 2016), fluorescence of the lipid droplets (stained with Nile Red) also
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appeared in the chlorophyll fluorescence channel (Fig. S5 panel B: in fact, each channel was a
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monochromatic image in the confocal microscope). As a consequence, the site where lipid droplets
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reside was shown as a red fluorescent area, and the lipid droplet (found in the green channel) was
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erroneously located within the chloroplast (but the red fluorescence in this area must be
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fluorescence due to the lipid droplet itself contaminating the red channel). Nile Red has a broad
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absorption peak, with a very low absorbance, above 600 nm, and this resulted in an observable
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fluorescence excited at 635 nm (Fig. S6 panel A), which is normally used as an excitation beam for
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imaging chlorophylls. The same was essentially true for LipidTOX (Fig. S6 panel C, Fig. S7 panels
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D–F), whereas the problem was significantly alleviated with BODIPY (Fig. S6 panel B, Fig. S7
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panels G–I). All of these problems were avoided by using blue autofluorescence for imaging
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chloroplasts or thylakoid membranes (Fig. S7 panel A, and all other fluorescence figures in the
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present study), although the exact origin of this fluorescence is unknown. All of these
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measurements suggested that the lipid droplets seemingly located within the chloroplast as reported
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by Goold et al. (2016) were not really located within the chloroplast. Therefore, we conclude that
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there is currently no solid evidence for the presence of lipid droplets within the chloroplast in
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Chlamydomonas.
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We finally propose a realistic model for the “chloroplast lipid droplets” in
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Chlamydomonas (Fig. 7). The lipid droplets seemingly present within the chloroplast stroma are
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actually present in the invaginations of the chloroplast (Fig. 7 panel A). It is theoretically possible
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that they could be perfectly isolated from the cytosol (Fig. 7 panel B), but we never found such lipid
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droplets with confidence. If the lipid droplets were actually isolated from the bulk cytosol,
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biochemical activities, such as protein synthesis and the supply of important metabolites such as
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ATP, would be quite limited.
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Biochemical data arguments for the localization of lipid droplets
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In addition to electron microscopic images, Fan et al. (2011) presented biochemical data, which
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they tried to use as evidence for the chloroplast origin of lipid droplets. They analyzed the fatty acid
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composition of the TAG that accumulated in starchless mutants under nitrogen deprivation, and
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showed that C16 fatty acids were abundant. There is a prevailing belief that the C18/C16 lipid
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molecular species originate from the lipid biosynthetic pathway in the chloroplast, and that the
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C18/C18 molecular species are synthesized in the ER. These two pathways are often called the
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“prokaryotic” and “eukaryotic” pathways, respectively. In Chlamydomonas, this tradition began
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with a study by Eichenberger’s group in the 1980s (Giroud et al., 1988), which was motivated by
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the popular paradigm of the plant lipid field of the time (Roughan and Slack, 1982). More recently,
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Warakanont et al. (2015) showed that the chain length is not a key to classify the origin of lipids in
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the context of the lipid traffic in Chlamydomonas. C16 fatty acids can be provided from the ER for
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synthesis of chloroplast lipids. TAG contains both C16 and C18 fatty acids, which are provided
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from either ER lipids or chloroplast lipids. The 16:0 fatty acids in TAG are likely to be provided by
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diacylglyceryl-N,N,N-trimethylhomoserine (DGTS) whereas the 16:4 fatty acid is probably
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provided by monogalactosyl diacylglycerol (MGDG) because MGDG is the sole lipid class that
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contains 16:4 as an abundant fatty acid (Sakurai et al., 2014 and many other studies). This does not
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mean that TAG containing 16:4 fatty acid is present in the chloroplast, as stated by Fan et al. (2011).
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We imagine that the 16:4 fatty acid is hydrolyzed from MGDG (Li et al., 2012), transported to the
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cytosolic compartment, and incorporated into TAG by acyltransfer reactions.
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Another biochemical argument for localization of the lipid droplets is related to the
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possible use of a protein in the lipid droplet membrane as a marker for the cytosolic lipid droplet.,
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Various proteins in the lipid droplet membrane have been characterized in plants, and are used as
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markers for the lipid droplet membrane. In Chlamydomonas, major lipid droplet protein (MLDP)
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has been characterized as the major constituent of the lipid droplet membrane formed under
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nitrogen deprivation (Moellering and Benning, 2010; Tsai et al., 2015). We analyzed the level of the
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MLDP transcript in conditions known to favor the accumulation of lipid droplets (Fig. S9). Under
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nitrogen deprivation, expression of the MLDP gene increased about 15-fold, which was consistent
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with the results of Moellering and Benning (2010). However, expression of the MLDP gene under
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the high-light condition did not increase above the control level. The data in Goold et al. (2016)
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were ambiguous: the MLDP signal was scarcely found in the silver-stained gel, whereas MLDP was
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detected as the 14th most abundant lipid droplet protein by a proteome analysis. These results
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suggest that it is difficult to use MLDP as a general marker of lipid droplet membranes. In addition,
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all of the lipid droplet analytical data were obtained for lipid droplets isolated from whole cells,
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including both cytosolic lipid droplets and the putative “chloroplast lipid droplets” (if such entities
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exist). It is theoretically impossible to distinguish the cytosolic lipid droplet and the hypothetical
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“chloroplast lipid droplet” by using a marker developed with the lipid droplet preparation isolated
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in the published way. That is why we preferred microscopic methods to biochemical methods for
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identifying the cytosolic nature of the lipid droplets, which seemed to be “chloroplast lipid
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droplets”.
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Putative inter-organellar metabolic flow
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A remaining problem is why some lipid droplets are present in invagination of the chloroplast.
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Similar invaginations are not found in the chloroplasts of land plants. This phenomenon might be
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specific to Chlamydomonas or related algae having a single large chloroplast. Invaginations or
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holes might be naturally present in such large chloroplast, and the holes could facilitate transport of
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cytosolic materials to the outer cytosolic compartment between the chloroplast and the plasma
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membrane. Holes and invaginations might be hardly observable without carefully examining by a
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fluorescence microscope. A single chloroplast often appeared as several segments in the electron
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micrographs of wild-type cells under normal conditions without lipid droplet accumulation (Fig. S8,
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arrows), and these represented holes or the reticulated or branched architecture of the chloroplast.
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We identified several “holes” within the chloroplast in the confocal images (Fig. S7 C, arrowheads).
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Under the conditions in which TAG accumulates, these invaginations and holes are filled with
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growing lipid droplets. We still do not know if this filling of the invaginations by lipid droplets
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occurs passively as a result of mechanical packing within the cell. Alternatively, the tight
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association between the lipid droplet and the chloroplast at the site of an invagination or hole might
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have intriguing importance in the metabolic flow of fatty acids from the chloroplast to the lipid
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droplet. We will discuss this possibility further.
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We were impressed by the close positioning of the lipid droplets with the outer envelope
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membrane, which formed the inner surface of the chloroplast invagination (Fig. 5, Fig. S4, Fig. S10
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panels B and C). This close relationship between the lipid droplet and the chloroplast envelope
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membrane could facilitate efficient transport of fatty acids synthesized within the chloroplast to the
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growing lipid droplet. In general, lipid droplets emerge from the ER membrane by budding. The
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precursors of TAG, such as diacylglycerol, phosphatidylcholine or acyl-CoA, are provided by
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enzymes localized in the ER. However, no ER membranes were detected between the lipid droplet
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and the chloroplast envelope at the site of their close association. The space between the chloroplast
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envelope membrane and the surface of the lipid droplet was very narrow. It was narrower than the
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thickness of a typical ER cisterna. All precursors for synthesizing TAG must be supplied directly
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from the chloroplast to the lipid droplet in the lipid droplets located within the invagination of
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chloroplast,. This was an intriguing new aspect of lipid droplet formation. The problem of the
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“chloroplast lipid droplet” will have to be studied under a different perspective, namely, lipid traffic
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(just as in Block and Jouhet, 2015, for a somewhat different type of traffic) from the chloroplast to
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the lipid droplet. We still do not know how the initial lipid droplet is formed, which finally localize
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to the chloroplast invagination.
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Tsai et al. (2015) noted the presence of plastid proteins and the plastid lipid digalactosyl
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diacylglycerol (DGDG) in the lipid droplets of Chlamydomonas, and discussed a putative origin of
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lipid droplets from the chloroplast envelope membranes. Many outer envelope proteins, such as
348
TOC75, TOC34, TGD2, and MGD1, co-immunoprecipitated with MLDP. This is explicitly depicted
349
in Figure 2 of Liu and Benning (2015), who suggested a possible flow of materials from the
350
chloroplast envelope to the lipid droplet. An integration of the lipid droplet within the chloroplast
351
envelope, rather than contact of the membrane and the lipid droplet, was presented in Figure 3 of
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352
Li-Beisson et al. (2015). Although Liu and Benning (2015) were very cautious about the probable
353
non-specific contamination, the presence of light-harvesting chlorophyll proteins and ATPase
354
subunits, which are often detected in the proteomics of subcellular fractions, suggested that we need
355
different types of evidence, such as fluorescence energy transfer results, to conclude the presence of
356
envelope membrane proteins in the lipid droplet. Despite the figure in Liu and Benning (2015) that
357
showed direct contact or complete fusion of the lipid droplet membrane and the outer envelope, no
358
such image has ever been either presented in the literature or detected by our observations. Most
359
electron micrographs presented in the literature were taken at low magnifications to show the entire
360
structure of the cell (about ×10,000), but the relationship between the lipid droplet and the envelope
361
must be observed at a higher magnification, such as ×50,000 or higher. As shown schematically in
362
Figure 1 for the two envelope membranes, two parallel membranes can be seen as fused in the
363
examination of thin sections. The same is true for the lipid droplet membrane and the envelope
364
membranes. We occasionally observed a lipid droplet apparently fused with the envelope
365
membranes in thin sections, but the two objects were separated if we observed the sections by
366
tilting them in a manner shown in Figure 5. In this sense, no image has ever been presented
367
showing direct contact or fusion of the lipid droplet membrane and the chloroplast envelope.
368
The proteomics or lipid analysis data of the isolated lipid droplet could be interpreted to
369
show that during the fractionation the lipid droplet present within the chloroplast invagination was
370
released as a complex structure consisting of the lipid droplet wrapped by the chloroplast envelope
371
membranes, with the inner envelope outside, and the outer envelope inside. This is a more realistic
372
model of isolated lipid droplets bearing DGDG and outer envelope proteins, which are present in a
373
minor proportion (the majority of the lipid droplets are certainly those that originate from the
374
cytosolic compartment). Therefore, a realistic image of the “chloroplast lipid droplet” is a lipid
375
droplet, which is present within the chloroplast invagination, in close association with the
376
chloroplast envelope membrane by keeping a very narrow layer of cytosol. We cannot exclude the
377
possibility that some structures bridge the lipid droplet and the envelope. In this regard, it is
378
interesting to note that we also found close associations between the Golgi apparatus, the lipid
379
droplets, and the chloroplast (Fig. 4 panel A and Fig. S10). It appears that the Golgi apparatus
380
provides enzymes TAG synthesis while the chloroplast provides fatty acids.
381
382
Distinguishing the plastoglobule from the lipid droplet
383
384
The plastoglobule is known to contain TAG, which is supposed to be synthesized by FES1/2
385
(Lippold et al., 2012), also called DGAT3/4. As clearly stated by Schimper (1885, p. 182), the
386
plastoglobule and lipid droplet must be clearly distinguished. He showed that the former is soluble
387
in ethanol, whereas the latter is not. The plastoglobule is different from a lipid droplet in that it
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388
contains various compounds other than TAG, such as plastoquinones and tocopherols, among others.
389
The plastoglobule contains various types of specific proteins known as fibrillins (FBN) (Lundquist
390
et al., 2012; Lohscheider and Bártulos, 2016). In the list of proteins detected by the proteomic
391
analysis of plastoglobules in Arabidopsis thaliana, we found no protein that was known to localize
392
to the cytosolic lipid droplets. In the discussion provided by groups of both Fan et al (2011) and
393
Goold et al. (2016), the ability of the chloroplast to synthesize TAG by FES1/2 was taken as
394
evidence for the presence of a lipid droplet within the chloroplast. If a significant number of lipid
395
droplets are present within the chloroplast, and if TAG is provided by the plastoglobules as
396
suggested, then FBN might be detected in the proteome of lipid droplets. However, there is no
397
protein in plastoglobules, such as FBN, in the list of proteins detected by the proteomic analysis of
398
Chlamydomonas lipid droplets (Moellering and Benning, 2010; Nguyen et al., 2011; Tsai et al.,
399
2015). The only putative FBN-related protein is g7551 (530 aa), described by Tsai et al. (2015). It
400
has a partial homology to plant FBN, but this entry was changed to a short (130 aa) protein
401
(A8JH10), and is not included in the list of FBNs in Lohscheider and Bártulos (2016). Therefore, no
402
FBNs are detected in isolated lipid droplets, and it is unlikely that even a small population of lipid
403
droplets is localized in the chloroplast and supplied with TAG synthesized by plastoglobules.
404
During our electron microscopic sample preparation the cells were dehydrated with
405
ethanol; thus, TAG remained as a liquid material mixed with the resin within the section for
406
observation. Some of the electron micrographs showed wavy appearing lipid droplets due to
407
inefficient cutting of the TAG/resin mixture. The plastoglobules were seen as small particles (Fig. 4
408
panel E). Goold et al (2016) presented “plastidal lipid droplets”, but according to our examination,
409
these are lipid droplets not plastoglobules: the size of the droplets was smaller than the typical large
410
lipid droplets accumulated in nitrogen-starved cells, but they were fairly larger (>200 nm in
411
diameter) than typical plastoglobules, which were about 100 nm in diameter (Fig. 4 panel E). The
412
lipid droplets presumptively located within the chloroplast were similar in size to those undoubtedly
413
present in the cytosol. There is no reason to assume that the “plastidal lipid droplets” were
414
something other than typical lipid droplets. All lipid droplets, whether “chloroplast lipid droplets”
415
or not, must be present within the cytosolic compartment.
416
We noted that lipid droplets that appear to localize within the chloroplast retain a thin
417
limiting membrane. This is very difficult to see, but a thin membrane was visible in a limited region
418
depending on the tilt angles (Fig. 4 panel D, Fig. S10 panel B). No limiting membrane is known in
419
the plastoglobule. This observation is also evidence that the lipid droplets are considered to be
420
cytosolic and not located in the chloroplast.
421
422
Concluding remarks
423
Based on the electron microscopic examination and the 3D reconstruction by confocal fluorescence
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424
microscopy, we conclude that there is no evidence for the presence of lipid droplets in the
425
chloroplast of Chlamydomonas. We also provide various biochemical arguments using published
426
data on the lipid and protein compositions of the lipid droplets and plastoglobules, which are taken
427
as convincing evidence that the lipid droplets are essentially different from plastoglobules. In
428
addition, we found a new aspect of research on the lipid droplets, namely, the traffic from the
429
chloroplast to the lipid droplet without an intervening ER membrane.
430
431
MATERIALS AND METHODS
432
433
Algal strains and growth conditions
434
435
The green alga C. reinhardtii strains CC-1010, CC-4348 (cw15sta6-1; BAFJ5), and CC-4334
436
(cw15sta7-1) were obtained from the Chlamydomonas Resource Center (St. Paul, MN, USA).
437
CC-1010 cells were grown in modified Bristol’s medium (MBM) (Watanabe, 1960) with aeration
438
by 2% CO2 in air at 25°C. For lipid accumulation under high light irradiation, the culture was first
439
illuminated at a photon flux of 40 µmol m–2 s–1 until OD750 became 0.2–0.4, and then illuminated at
440
200 µmol m–2 s–1 according to Goold et al. (2016). The starchless mutants, sta6 and sta7 cells, were
441
grown in TAP medium (Gorman and Levine, 1965) at 25°C under light (50 µmol m–2 s–1) with
442
shaking. The cells were transferred to nitrogen-depleted TAP medium containing 20 mM potassium
443
acetate (TAP-N+Ac) for 24 h to accumulate lipids.
444
445
Microscopic examination
446
447
CC-1010 cells stained with BODIPY were observed under a fluorescence microscope as described
448
previously (Sakurai et al., 2014). For confocal microscopy, CC-1010 cells were fixed in 0.25%
449
glutaraldehyde and stained with 10 μg mL–1 BODIPY or 0.5% LipidTOX Red solution (Invitrogen,
450
Carlsbad, CA, USA). Unfixed sta6 and sta7 cells were stained with 0.5% LipidTOX Red solution
451
and 1 μM fluorescein diacetate (FDA) cytosolic staining. The samples were placed on a glass-based
452
dish (cover glass thickness 0.08–0.12 mm) and were imaged with a Nikon C2+ confocal imaging
453
system (Nikon Instech, Tokyo, Japan) mounted on a Nikon Eclipse Ti-E inverted microscope.
454
Fluorescence signals of chloroplasts and LipidTOX were acquired sequentially with 408-nm
455
excitation and 417–477-nm emission (see below), and 561-nm excitation and 575–615-nm emission,
456
respectively. BODIPY and FDA fluorescence signals were acquired by 488-nm excitation and 500–
457
550-nm emission. Images were obtained as Z-stacks, and processed with Fiji software
458
(http://fiji.sc/).
459
(http://www.sci.utah.edu/software/fluorender.html).
Reconstructed
3D
images
were
visualized
by
FluoRender
14
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software
460
Chloroplasts are usually imaged with chlorophyll fluorescence, typically using excitation
461
at 640 nm and emission at 660–1000 nm (Fig. S7 B). As pointed out in Figure S5, fluorescence of
462
Nile Red or LipidTOX can enter the chlorophyll channel (Fig. S6 and Fig. S7). This is inevitable if
463
chlorophyll fluorescence is used for imaging chloroplasts (or thylakoid membranes). We discovered
464
that the blue region can be used to obtain images of chloroplasts (excitation at 408 nm, and
465
emission at 417–477 nm). The image obtained with blue fluorescence was identical to the image
466
obtained with conventional red fluorescence (Fig. S7 panels A–C). We still do not know the exact
467
origin of this blue fluorescence of chloroplasts, but this technique is useful for avoiding
468
fluorescence of lipid stains (Nile Red, BODIPY, or LipidTOX) from making “fake” images on the
469
chloroplast image.
470
471
Fluorometric quantification of neutral lipids
472
473
The cells were fixed with 0.25% glutaraldehyde and stored at 4°C until measurement. 2 × Nile Red
474
solution (1 μg mL–1 Nile Red and 50% DMSO) was added to each sample, and the mixture was
475
incubated for 5 min at room temperature. Fluorescence emission at 604 nm of Nile Red bound to
476
neutral lipid was measured under excitation at 530 nm in an EnSpire plate reader (PerkinElmer,
477
Waltham, MA, USA). Background fluorescence was also measured and subtracted.
478
479
Fluorescence spectral analysis
480
481
Fluorescence of Nile Red, LipidTOX, and BODIPY dissolved in TAG was measured using a
482
Shimadzu RF-5300PC fluorescence spectrophotometer (Kyoto, Japan).
483
484
Lipid analysis
485
486
Extraction of lipids from cells, thin-layer chromatographic separation of lipid classes, and gas
487
chromatographic determination of fatty acid methyl esters were performed essentially as described
488
previously (Sakurai et al., 2014).
489
490
Transmission electron microscopy
491
492
Cells were fixed in 0.125% glutaraldehyde for 10 min and then with 1% glutaraldehyde for 30 min.
493
They were centrifuged at 400 × g for 10 min, and the precipitate was embedded in 1% low-melting
494
agarose. Dehydration through an ethanol series, embedding in Epon resin, sectioning, and staining
495
were performed as described previously (Sato et al., 2014; Toyoshima and Sato, 2015).
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496
497
Acknowledgments
498
The authors thank Drs. Fan, J. and Xu, C., Brookhaven National Laboratory, for discussion and
499
sharing unpublished data. They also thank Dr. Li-Beisson, Y., CEA Cadarache, for discussion on
500
their published data. Despite these discussions, disaccords still remained, unfortunately. The authors
501
are grateful to Mr. Takashi Hirashima in our laboratory for discussion on plastoglobules.
502
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503
Table 1. Fatty acid composition and accumulation of triacylglycerol in C. reinhardtii CC1010
504
cells subjected to high-light irradiation for 7 hours (n = 3). The values show the composition of
505
fatty acids in percentage. The level of TAG is shown at the bottom. It was noted that the fatty acids
506
specific to chloroplast lipids, namely, 16:4 and 18:3(9,12,15), did not appreciably accumulate in
507
TAG. In Sakurai et al. (2014), 18:4 was supposed to be a mixture of 18:4(5,9,12,15) and
508
18:4(6,9,12,15), but, as reported in Sato et al. (2016), 18:4(6,9,12,15) was not detected in C.
509
reinhardtii CC-1010.
510
Fatty acid
14:0
14:1
16:0
16:1(7)
16:2(7,10)
16:3(4,7,10)
16:3(7,10,13)
16:4(4,710,13)
17:0
18:1(9)
18:1(11)
18:2(9,12)
18:3(5,9,12)
18:3(9,12,15)
18:4(5,9,12,15)
TAG % of total
lipids
Total lipids
0h
7h
0.4±0.1
0.4±0.0
0.3±0.1
0.2±0.0
24.2±1.8
24.7±0.3
2.0±0.4
2.5±0.6
4.8±1.0
3.3±0.7
0.9±0.2
0.9±0.0
7.8±0.3
7.7±0.8
10.1±2.3
10.0±1.3
0.2±0.1
0.1±0.0
4.1±0.2
5.1±1.2
2.9±0.7
3.0±0.3
16.1±2.8
14.7±2.3
8.7±0.5
9.0±0.5
15.9±1.9
16.9±1.1
1.7±0.2
1.6±0.2
—
—
Triacylglycerol
0h
7h
4.4±0.7
2.2±0.8
1.4±0.3
0.9±0.7
35.4±2.5
30.6±3.2
1.7±0.1
2.0±0.5
1.1±0.3
2.1±0.8
0.3±0.3
0.4±0.1
6.7±0.8
6.9±1.5
2.3±0.2
3.3±0.1
0.4±0.4
0.4±0.2
12.6±1.9
16.1±1.7
4.2±1.0
3.9±0.4
13.1±1.3
14.5±1.8
8.8±1.1
7.3±0.5
6.1±0.6
8.0±0.3
1.5±0.4
1.7±0.2
3.5±0.2
7.8±1.3
511
512
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513
Figure legends
514
515
Figure 1. Schematic model of the relationship between the lipid droplet and the chloroplast. A
516
– C, Schematic views of hypothetical different localizations of lipid droplet (LD). Nuc, nucleus; Cp,
517
chloroplast; CW, cell wall; D and E, Schematic explanation of different views of the chloroplast
518
envelope membranes (CpEnv) in an ultrathin section. Detection of the chloroplast envelope
519
membranes is crucial in identifying the localization of lipid droplet. We consider the case in which a
520
lipid droplet is present within a cytosolic compartment enclosed within the chloroplast, as shown by
521
the broken line in B or C. In this case, the lipid droplet must be surrounded by the chloroplast
522
envelope membranes. If a section is cut at the center of the lipid droplet which appears to be located
523
within the chloroplast, we can recognize that the lipid droplet is surrounded by the chloroplast
524
envelope membranes, and therefore, is located within the cytosolic compartment (section 1). But if
525
a section (section 2) is cut out of the center, then the chloroplast envelope membranes and the lipid
526
droplet half-membrane become obscure, because the thickness of the section (usually 70 – 100 nm)
527
is much larger than the thickness of the membranes (7 – 10 nm for envelope membranes, and 3 – 4
528
nm for lipid droplet half-membrane). If the section is appropriately tilted (E), then we can identify
529
the membranes partly.
530
531
Figure 2. Neutral lipid accumulation in C. reinhardtii CC1010 cells subjected to high-light
532
irradiation. A, Observation of BODIPY-stained cells. The cells grown under low light were shifted
533
to high light and grown for 11 h. DIC, differential interference contrast microscope image; Merged,
534
merged image of BODIPY and DIC; B, Measurement of fluorescent signal of Nile Red. The cells
535
grown as in A were stained with Nile Red and the fluorescent signal was measured by a plate reader.
536
537
Figure 3. 3D reconstruction by confocal fluorescence microscopy of wild-type C. reinhardtii
538
CC-1010 cell, after the transfer from low light to high light. The cells grown under high light for
539
7 h were stained with BODIPY and observed by confocal microscopy with Z-stacking. The
540
fluorescence of BODIPY and blue autofluorescence are pseudo-colored. A-C, 2D images of a slice,
541
in which a lipid droplet (white arrow) seemed to be entirely enclosed by the chloroplast. Images A
542
and B show the fluorescence of BODIPY and blue autofluorescence, respectively. Merged; merged
543
image of BODIPY and autofluorescence images. D-I, Rendering images of the same cell as the one
544
shown in A-C. D-F show the images viewed from an identical angle. G-I show the images viewed
545
from a different angle. E and F are clipped images of D at X- (E) or X- and Z-axes (F). H and I are
546
clipped images of G at Y- (H) or Y- and Z-axes (I). The white arrows indicate an identical lipid
547
droplet in A, C, E, F, H, and I.
548
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549
Figure 4. Electron micrograph of a lipid droplet entrapped by a chloroplast in wild-type C.
550
reinhardtii CC-1010 cells, seven hours after the transfer from low light to high light. A, Image
551
of a chloroplast seemingly including a lipid droplet in its stroma; B, Image of the same chloroplast
552
with 17˚ tilting; C, Same as B with tracing chloroplast envelope. D, Enlarged lipid droplet, showing
553
a putative limiting membrane (white arrowhead); E, Plastoglobules within the chloroplast
554
(arrowheads). Cyto, cytosol; Golgi, Golgi apparatus; Mt, mitochondrion; LD, lipid droplet; Starch,
555
starch granule; Stroma, chloroplast stroma.
556
557
Figure 5. Electron micrographs of lipid droplets in the starchless mutant (cw15sta6) cells of C.
558
reinhardtii, deprived of nitrogen for 24 hours with supplementation of acetate. A, Whole cell;
559
B–D, tilted images as indicated along the directions shown above. Traces of identifiable parts of
560
chloroplast envelope membranes (single lines for a set of inner and outer membranes) are presented
561
in the lower panels (E – G). Visibility of the membranes depended on the tilt angle. Cp, chloroplast;
562
N, nucleus; LD, lipid droplet. Red arrows indicate the cytoplasmic ribosomes.
563
564
Figure 6. 3D reconstruction by confocal fluorescence microscopy of the starchless mutant cells
565
of C. reinhardtii, deprived of nitrogen for 24 hours with acetate supplementation. Upper panels,
566
a representative cw15sta6 cell; Lower panels, a representative cw15sta7 cell. All images are
567
pseudocolored. A, Localization of lipid droplets detected by LipidTOX fluorescence (green); B,
568
Localization of chloroplast detected by blue autofluorescence (but shown in red); C. Localization of
569
cytosol detected by FDA fluorescence (blue); D. Merged images of A–C; E–G, Views of 3D
570
reconstructed images by different clipping. The staining with FDA in living cells will give uniform
571
fluorescence after a long time due to hydrolysis of FDA by esterases leaked from broken cells. We
572
had to work rapidly with a lowest concentration of FDA. That is why the intensity of fluorescence
573
was weak in this observation.
574
575
Figure 7. Models of localization of lipid droplet in C. reinhardtii. A, A plausible model of a lipid
576
droplet seemingly located within a chloroplast, but localized within the cytosolic compartment,
577
which is present in a chloroplast invagination. The close association of chloroplast envelope and the
578
lipid droplet half membrane supports the transfer of fatty acids from the chloroplast to the lipid
579
droplet. B, A hypothetical model of a lipid droplet seemingly embedded within a chloroplast. The
580
cytosol is shown by cyan. Note that the lipid droplet is surrounded by cytosol. Cp, chloroplast;
581
CpEnv, chloroplast envelope; Nuc, nucleus; LD, lipid droplet. This figure shows only a lipid droplet
582
seemingly present within the chloroplast (type C localization). Besides this type of lipid droplets,
583
there are a number of lipid droplets located in the cytosol between the nucleus and the chloroplast
584
(type A) or outside the chloroplast (type B).
19
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585
586
587
Supplemental data
588
589
Supplemental Figure S1. Comparison of LipidTOX- and BODIPY-staining.
590
591
Supplemental Figure S2. 3D reconstruction by confocal fluorescence microscopy of another
592
wild-type cell of C. reinhardtii CC-1010.
593
594
Supplemental Figure S3. 3D reconstruction by confocal fluorescence microscopy of a third
595
wild-type cell of C. reinhardtii CC-1010.
596
597
Supplemental Figure S4. Electron micrographs of lipid droplets in the starchless mutant (cw15sta7)
598
cells of C. reinhardtii, deprived of nitrogen for 24 hours with acetate supplementation.
599
600
Supplemental Figure S5. Re-examination of the published figures of chloroplast localization of lipid
601
droplets in C. reinhardtii.
602
603
Supplemental Figure S6. Fluorescence spectra of representative pigments for lipid droplet staining
604
in pure triacylglycerol.
605
606
Supplemental Figure S7. Confocal imaging of chloroplasts and lipid droplets.
607
608
Supplemental Figure S8. Putative holes in the chloroplast in the strain CC-1010.
609
610
Supplemental Figure S9. Levels of MLDP gene transcripts under various growth conditions in C.
611
reinhardtii CC-1010.
612
613
Supplemental Figure S10. Close association of Golgi apparatus – lipid droplet – chloroplast.
614
615
Supplemental Data S1 – S3. Sequential TIF files of the 3D reconstruction of LD-accumulating
616
CC-1010 cells for Figure 3, Figure S2 and Figure S3.
617
618
Supplemental Data S4 and S5. Sequential TIF files of the 3D reconstruction of LD-accumulating
619
sta6 and sta7 cells, respectively, for the upper and lower parts of Figure 6.
620
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621
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Besagni C, Kessler F, Stymne S, Dörmann P (2012) Fatty acid phytyl ester synthesis in chloroplasts
677
of Arabidopsis. Plant Cell 24: 2001–2014
678
679
Liu B and Benning C (2013) Lipid metabolism in microalgae distinguishes itself. Curr Opin
680
Biotechnol 24: 300–309
681
682
Lohscheider JN, Bártulos CR (2016) Plastoglobules in algae: A comprehensive comparative study
683
of the presence of major structural and functional components in complex plastids. Marine
684
Genomics 28: 127–136
685
686
Lundquist PK, Poliakov A, Bhuiyan NH, Zybailov B, Sun Q, van Wijk KJ (2012) The functional
687
network of the Arabidopsis plastoglobule proteome based on quantitative proteomics and
688
genome-wide co-expression analysis. Plant Physiol 158: 1172–1192
689
690
Merchant SA, Kropat J, Liu B, Shaw J, Warakanont J (2012) TAG, You’re it! Chlamydomonas as a
691
reference organism for understanding algal triacylglycerol accumulation. Curr Opin Biotechnol 23:
692
352–363
22
Downloaded from on October 25, 2017 - Published by www.plantphysiol.org
Copyright © 2017 American Society of Plant Biologists. All rights reserved.
693
694
Misra N, Panda PK, Parida BK, Mishra BK (2012) Phylogenomic study of lipid genes involved in
695
microalgal biofuel production—candidate gene mining and metabolic pathway analyses. Evol
696
Bioinformatics 8: 545–564
697
698
Moellering ER, Benning C (2010) RNA interference silencing of a major lipid droplet protein
699
affects lipid droplet size in Chlamydomonas reinhardtii. Eukaryot Cell 9: 97–106.
700
701
Nguyen HM, Baudet M, Cuiné S, Adriano JM, Barthe D, Billon E, Bruley C, Beisson F, Peltier G,
702
Ferro M, Li-Beisson Y (2011) Proteomic profiling of oil bodies isolated from the unicellular green
703
microalga Chlamydomonas reinhardtii: With focus on proteins involved in lipid metabolism.
704
Proteomics 11: 4266–4273
705
706
Pasteur L (1922) Œuvres de Pasteur, vol. II, p. 295 and p. 459, Masson, Paris, available from
707
Bibliothèque National de France.
708
709
Roughan PG, Slack CR (1982) Cellular organization of glycerolipid metabolism. Annu Rev Plant
710
Physiol 33: 97–132.
711
712
Sakurai K, Moriyama T, Sato N (2014) Detailed identification of fatty acid isomers sheds light on
713
the probable precursors of triacylglycerol accumulation in photoautotrophically grown
714
Chlamydomonas reinhardtii. Eukaryot Cell 13: 256–266
715
716
Sato N, Katsumata Y, Sato K, Tajima N (2014) Cellular dynamics drives the emergence of
717
supracellular structure in the cyanobacterium, Phormidium sp. KS. Life 4: 819–836
718
719
Sato N, Mori N, Hirashima T, Moriyama T (2016) Diverse pathways of biosynthesis of
720
phosphatidylcholine in algae as estimated by labeling studies and genomic sequence analysis. Plant
721
J 87: 281–292
722
723
Schimper AFW (1885) Untersuchungen über die Chlorophyllkörper und die ihnen homologen
724
Gebilde. Jarb F wiss Botanik 16: 1–247
725
726
Siaut M, Cuiné, S, Cagnon C, Fessler B, Nguyen M, Carrier P, Beyly A, Beisson F, Triantaphylidès
727
C, L-Beisson Y, Peltier G (2011) Oil accumulation in the model green alga Chlamydomonas
728
reinhardtii: characterization, variability between common laboratory strains and relationship with
23
Downloaded from on October 25, 2017 - Published by www.plantphysiol.org
Copyright © 2017 American Society of Plant Biologists. All rights reserved.
729
starch reserves. BMC Biotechnol 11: 7
730
731
Tevini M, Steinmüller D (1985) Composition and function of plastoglobuli. II. Lipid composition of
732
leaves and plastoglobuli during beech leaf senescence. Planta 163: 91–96
733
734
Toyoshima M, Sato N (2015) High-level accumulation of triacylglycerol and starch in
735
photoautotrophically grown Chlamydomonas debaryana NIES-2212. Plant Cell Physiol 56: 2447–
736
2456
737
738
Tsai C-H, Zienkiewicz K, Amstutz CL, Brink BG, Warakanont J, Roston R, Benning C (2015)
739
Dynamics of protein and polar lipid recruitment during lipid droplet assembly in Chlamydomonas
740
reinhardtii. Plant J 83: 650–660
741
742
van de Meene AML, Hohmann-Marriott MF, Vermaas WFJ, Roberson RW (2006) The
743
three-dimensional structure of the cyanobacterium Synechocystis sp. PCC 6803. Arch Microbiol
744
184: 259–270
745
746
Warakanont J, Tsai C-H, Michel EJS, Murphy GR, Hsueh PY, Roston RL, Sears BB, Benning C
747
(2015) Chloroplast lipid transfer processes in Chlamydomonas reinhardtii involving a
748
TRIGALACTOSYLDIACYLGLYCEROL 2 (TGD2) orthologue. Plant J 84: 1005–1020
749
750
Watanabe A (1960) List of algal strains in collection at the Institute of Applied Microbiology,
751
University of Tokyo. J Gen Appl Microbiol 6: 283–292
752
753
Work VH, Radakovits R, Jinkerson RE, Meuser JE, Elliott LG, Vinyard DJ, Laurens LMI,
754
Dismukes GC, Posewitz MC (2010) Increased lipid accumulation in the Chlamydomonas
755
reinhardtii sta7-10 starchless isoamylase mutant and increased carbohydrate synthesis in
756
complemented strains. Eukaryot Cell 9: 1251–1261
757
758
Zienkiewicz K, Du ZY, Ma W, Vollheyde K, Benning C (2016) Stress-induced neutral lipid
759
biosynthesis in microalgae — Molecular, cellular and physiological insights. Biochim Biophys Acta
760
1861: 1269–1281
761
24
Downloaded from on October 25, 2017 - Published by www.plantphysiol.org
Copyright © 2017 American Society of Plant Biologists. All rights reserved.
(A) LD between Nuc and Cp
(B) LD between Cp and CW
(C) LD inside Cp?
Nuc
Cp
LD
(D) Ultrathin section
Side view
(E) 20˚ tilt
Side view
CpEnv
Stroma
Lipid droplet
Thin section 1
(80 nm)
Thin section 2
tion
ec
in s
Th
Cytosol
Top view 1 (EM image)
Top view 1 (EM image)
Top view 2 (Membranes become obscure)
Downloaded from on October 25, 2017 - Published by www.plantphysiol.org
Copyright © 2017 American Society of Plant Biologists. All rights reserved.
2
Downloaded from on October 25, 2017 - Published by www.plantphysiol.org
Copyright © 2017 American Society of Plant Biologists. All rights reserved.
BODIPY
A
B
Blue autofluorescence
C
Merged
5 µm
D
Rendering view by depth
Y
E
Clipping of X-axis
Y
G
F
Clipping of X- and Z-axes
Y
X
X
X
Z
Z
Z
Rendering view by depth
Y
H
Clipping of Y-axis
Y
Z
Clipping of Y- and Z-axes
Y
Z
X
I
Z
X
X
Downloaded from on October 25, 2017 - Published by www.plantphysiol.org
Copyright © 2017 American Society of Plant Biologists. All rights reserved.
Downloaded from on October 25, 2017 - Published by www.plantphysiol.org
Copyright © 2017 American Society of Plant Biologists. All rights reserved.
Downloaded from on October 25, 2017 - Published by www.plantphysiol.org
Copyright © 2017 American Society of Plant Biologists. All rights reserved.
sta6
FDA
LipidTOX Autofluorescence
A
B
Merged
C
D
5 µm
E Rendering view by depth
F Clipping of X-axis
G Clipping of X- and Z-axes
Y
Y
Y
XX
ZZ
sta7
X
X
ZZ
A
B
X
X
ZZ
C
D
5 µm
E
Rendering view by depth
Y
F
Clipping of X-axis
Y
X
Z
G Clipping of X- and Z-axes
Y
X
Z
X
Z
Downloaded from on October 25, 2017 - Published by www.plantphysiol.org
Copyright © 2017 American Society of Plant Biologists. All rights reserved.
5 µm
A
B
Nuc
Nuc
Cp
LD
Cp
LD
CpEnv
Downloaded from on October 25, 2017 - Published by www.plantphysiol.org
Copyright © 2017 American Society of Plant Biologists. All rights reserved.
CpEnv
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Liu B and Benning C (2013) Lipid metabolism in microalgae distinguishes itself. Curr Opin Biotechnol 24: 300–309
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CrossRef: Author and Title
Google Scholar: Author Only Title Only Author and Title
Lohscheider JN, Bártulos CR (2016) Plastoglobules in algae: A comprehensive comparative study of the presence of major structural
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CrossRef: Author and Title
Google Scholar: Author Only Title Only Author and Title
Lundquist PK, Poliakov A, Bhuiyan NH, Zybailov B, Sun Q, van Wijk KJ (2012) The functional network of the Arabidopsis plastoglobule
proteome based on quantitative proteomics and genome-wide co-expression analysis. Plant Physiol 158: 1172–1192
Pubmed: Author and Title
CrossRef: Author and Title
Google Scholar: Author Only Title Only Author and Title
Merchant SA, Kropat J, Liu B, Shaw J, Warakanont J (2012) TAG, You’re it! Chlamydomonas as a reference organism for understanding
algal triacylglycerol accumulation. Curr Opin Biotechnol 23: 352–363
Pubmed: Author and Title
CrossRef: Author and Title
Google Scholar: Author Only Title Only Author and Title
Misra N, Panda PK, Parida BK, Mishra BK (2012) Phylogenomic study of lipid genes involved in microalgal biofuel production—
candidate gene mining and metabolic pathway analyses. Evol Bioinformatics 8: 545–564
Pubmed: Author and Title
CrossRef: Author and Title
Google Scholar: Author Only Title Only Author and Title
Moellering ER, Benning C (2010) RNA interference silencing of a major lipid droplet protein affects lipid droplet size in
Chlamydomonas reinhardtii. Eukaryot Cell 9: 97–106.
Pubmed: Author and Title
CrossRef: Author and Title
Google Scholar: Author Only Title Only Author and Title
Nguyen HM, Baudet M, Cuiné S, Adriano JM, Barthe D, Billon E, Bruley C, Beisson F, Peltier G, Ferro M, Li-Beisson Y (2011) Proteomic
profiling of oil bodies isolated from the unicellular green microalga Chlamydomonas reinhardtii: With focus on proteins involved in
lipid metabolism. Proteomics 11: 4266–4273
Pubmed: Author and Title
CrossRef: Author and Title
Google Scholar: Author Only Title Only Author and Title
Pasteur L (1922) Œuvres de Pasteur, vol. II, p. 295 and p. 459, Masson, Paris, available from Bibliothèque National de France.
Pubmed: Author and Title
CrossRef: Author and Title
Google Scholar: Author Only Title Only Author and Title
Roughan PG, Slack CR (1982) Cellular organization of glycerolipid metabolism. Annu Rev Plant Physiol 33: 97–132.
Pubmed: Author and Title
CrossRef: Author and Title
Google Scholar: Author Only Title Only Author and Title
Sakurai K, Moriyama T, Sato N (2014) Detailed identification of fatty acid isomers sheds light on the probable precursors of
triacylglycerol accumulation in photoautotrophically grown Chlamydomonas reinhardtii. Eukaryot Cell 13: 256–266
Pubmed: Author and Title
Downloaded from on October 25, 2017 - Published by www.plantphysiol.org
Copyright © 2017 American Society of Plant Biologists. All rights reserved.
CrossRef: Author and Title
Google Scholar: Author Only Title Only Author and Title
Sato N, Katsumata Y, Sato K, Tajima N (2014) Cellular dynamics drives the emergence of supracellular structure in the cyanobacterium,
Phormidium sp. KS. Life 4: 819–836
Pubmed: Author and Title
CrossRef: Author and Title
Google Scholar: Author Only Title Only Author and Title
Sato N, Mori N, Hirashima T, Moriyama T (2016) Diverse pathways of biosynthesis of phosphatidylcholine in algae as estimated by
labeling studies and genomic sequence analysis. Plant J 87: 281–292
Pubmed: Author and Title
CrossRef: Author and Title
Google Scholar: Author Only Title Only Author and Title
Schimper AFW (1885) Untersuchungen über die Chlorophyllkörper und die ihnen homologen Gebilde. Jarb F wiss Botanik 16: 1–247
Pubmed: Author and Title
CrossRef: Author and Title
Google Scholar: Author Only Title Only Author and Title
Siaut M, Cuiné, S, Cagnon C, Fessler B, Nguyen M, Carrier P, Beyly A, Beisson F, Triantaphylidès C, L-Beisson Y, Peltier G (2011) Oil
accumulation in the model green alga Chlamydomonas reinhardtii: characterization, variability between common laboratory strains and
relationship with starch reserves. BMC Biotechnol 11: 7
Pubmed: Author and Title
CrossRef: Author and Title
Google Scholar: Author Only Title Only Author and Title
Tevini M, Steinmüller D (1985) Composition and function of plastoglobuli. II. Lipid composition of leaves and plastoglobuli during
beech leaf senescence. Planta 163: 91–96
Pubmed: Author and Title
CrossRef: Author and Title
Google Scholar: Author Only Title Only Author and Title
Toyoshima M, Sato N (2015) High-level accumulation of triacylglycerol and starch in photoautotrophically grown Chlamydomonas
debaryana NIES-2212. Plant Cell Physiol 56: 2447–2456
Pubmed: Author and Title
CrossRef: Author and Title
Google Scholar: Author Only Title Only Author and Title
Tsai C-H, Zienkiewicz K, Amstutz CL, Brink BG, Warakanont J, Roston R, Benning C (2015) Dynamics of protein and polar lipid
recruitment during lipid droplet assembly in Chlamydomonas reinhardtii. Plant J 83: 650–660
Pubmed: Author and Title
CrossRef: Author and Title
Google Scholar: Author Only Title Only Author and Title
van de Meene AML, Hohmann-Marriott MF, Vermaas WFJ, Roberson RW (2006) The three-dimensional structure of the cyanobacterium
Synechocystis sp. PCC 6803. Arch Microbiol 184: 259–270
Pubmed: Author and Title
CrossRef: Author and Title
Google Scholar: Author Only Title Only Author and Title
Warakanont J, Tsai C-H, Michel EJS, Murphy GR, Hsueh PY, Roston RL, Sears BB, Benning C (2015) Chloroplast lipid transfer
processes in Chlamydomonas reinhardtii involving a TRIGALACTOSYLDIACYLGLYCEROL 2 (TGD2) orthologue. Plant J 84: 1005–1020
Pubmed: Author and Title
CrossRef: Author and Title
Google Scholar: Author Only Title Only Author and Title
Watanabe A (1960) List of algal strains in collection at the Institute of Applied Microbiology, University of Tokyo. J Gen Appl Microbiol 6:
283–292
Pubmed: Author and Title
CrossRef: Author and Title
Google Scholar: Author Only Title Only Author and Title
Work VH, Radakovits R, Jinkerson RE, Meuser JE, Elliott LG, Vinyard DJ, Laurens LMI, Dismukes GC, Posewitz MC (2010) Increased
lipid accumulation in the Chlamydomonas reinhardtii sta7-10 starchless isoamylase mutant and increased carbohydrate synthesis in
complemented strains. Eukaryot Cell 9: 1251–1261
Pubmed: Author and Title
CrossRef: Author and Title
Google Scholar: Author Only Title Only Author and Title
Zienkiewicz K, Du ZY, Ma W, Vollheyde K, Benning C (2016) Stress-induced neutral lipid biosynthesis in microalgae — Molecular,
cellular and physiological insights. Biochim Biophys Acta 1861: 1269–1281
Pubmed: Author and Title
CrossRef: Author and Title
Google Scholar: Author Only Title Only Author and Title
Downloaded from on October 25, 2017 - Published by www.plantphysiol.org
Copyright © 2017 American Society of Plant Biologists. All rights reserved.
Downloaded from on October 25, 2017 - Published by www.plantphysiol.org
Copyright © 2017 American Society of Plant Biologists. All rights reserved.
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