Chloroplasts are photosynthetic organelles that evolved through the endosymbiosis between cyanobacteria-like symbionts and hosts. Many studies have attempted to isolate intact chloroplasts to analyze their morphological characteristics and photosynthetic activity. Although several studies introduced isolated chloroplasts into the cells of different species, their photosynthetic activities have not been confirmed. In this study, we isolated photosynthetically active chloroplasts from the primitive red alga Cyanidioschyzon merolae and incorporated them in cultured mammalian cells via co-cultivation. The incorporated chloroplasts retained their thylakoid structure in intracellular vesicles and were maintained in the cytoplasm, surrounded by the mitochondria near the nucleus. Moreover, the incorporated chloroplasts maintained electron transport activity of photosystem II in cultured mammalian cells for at least 2 days after the incorporation. Our top-down synthetic biology-based approach may serve as a foundation for creating artificially photosynthetic animal cells.

Photosynthesis originated 3.5 billion years ago, which is a billion years after the Earth formed.¹⁾ Photosynthesis is one of the metabolic processes that have influenced global environmental conditions and evolution on Earth. Cyanobacteria-like bacteria that emerged 1.8 billion years ago were taken up and maintained in host cells 1.2–1.6 billion years ago, which led to the evolution of chloroplasts in algae.^2),3) In the field of synthetic biology, researchers have attempted to artificially create photosynthesis, which has evolved over the long history of the Earth, through bottom-up and top-down approaches. A study is currently underway to synthesize artificial chloroplasts chemosynthetically at the molecular level as a bottom-up approach that involves constructing organelles from natural and synthetic molecules.⁴⁾ In contrast, many studies have been conducted to modify native chloroplasts to enable their use in heterologous environments as a top-down approach that aims to improve the function of native organelles.^5)-7)

It was discovered 80 years ago that chloroplasts isolated from plants can maintain their photosynthetic activity.⁸⁾ Unfortunately, how isolated chloroplasts can maintain the activity for a certain period of time has remained a mystery considering the loss of most genes necessary for photosynthesis, protein synthesis, and growth from the chloroplast genome via the evolution of algae and plants.⁹⁾ Although the mechanisms underlying chloroplast or algal uptake and survival within host cells remain unclear, there are reports of the transfer of photosynthetic activities to unrelated cellular systems. Examples include kleptoplasty in sacoglossan sea slugs and algal uptake by the yellow-spotted salamander.^10)-13) Therefore, many researchers have tried to mimic these phenomena in vitro by introducing isolated chloroplasts into the cells of other species.^14)-16) However, these incorporated chloroplasts can maintain their correct morphology in the host cells for only a few hours, and their photosynthetic activities remain unconfirmed. To date, there has been no report of the transfer of isolated chloroplasts into a heterologous host cell that resulted in detectable photosynthesis in the host cell.

Research on photosynthetic therapy has been conducted to treat diseases via the induction of photosynthesis in animal tissues and organs using injected algae or thylakoid membranes.^13),17)-19) In a recent study, nano-sized thylakoids injected into degenerative disease (e.g., osteoarthritis)-related chondrocytes, in which NADPH and ATP are depleted, produced energy, thereby contributing to the activation of animal cell anabolism.⁷⁾ Therefore, we speculated that introducing chloroplasts directly into animal cells without any modifications may be a cost-effective method for increasing energy production.

To clarify the mechanism underlying the evolution of algae and self-sustained chloroplasts to enhance future applications related to photosynthetic therapies, we introduced native chloroplasts into cultured animal cells, which were subsequently analyzed in terms of their photosynthetic activity. In this study, chloroplasts of the red alga Cyanidioschyzon merolae (schyzon) were introduced into cultured mammalian cells.²⁰⁾ Schyzon, which has retained the primitive characteristics of ancestral algae, exist in highly acidic and volcanic hot springs that seem to reflect prehistoric Earth environmental conditions.²¹⁾ In fact, schyzon is normally cultured at 42℃ in a medium with a pH of 2.3. Algal chloroplast genomes typically contain only approximately 100 genes, but the schyzon genome includes 243 genes.²²⁾ This suggests that the schyzon chloroplast may remain viable in animal cells for long periods. Notably, many algal cells become inactive at temperatures below 37℃, which is the optimal temperature for cultured mammalian cells, but schyzon cells remain active at this temperature, making it useful for synthetic biology research. Unlike chloroplasts in flowering plants, algal chloroplasts rarely differentiate into other plastids, such as leucoplasts or etioplasts, in response to environmental changes.²³⁾ Considering future applications, we speculated that chloroplasts incorporated into animal cells may remain stable and continuously produce energy regardless of environmental conditions. Moreover, we successfully developed a method for isolating chloroplasts from schyzon and have demonstrated through an X-ray analysis that the inner structure of isolated chloroplasts remains intact.²⁴⁾ As the host cultured mammalian cells, we used the Chinese hamster ovary (CHO)-K1 cell line, which has been used for the bioproduction of valuable substances and antibodies.^25),26) CHO-K1 cells are highly receptive to exogenous substances, making them suitable for maintaining algal chloroplasts. Finally, we measured the photosynthetic activity of algal chloroplasts in CHO-K1 cells using imaging pulse amplitude modulation (PAM) fluorometry.^27),28)

To isolate chloroplasts from schyzon, algal cells were treated with a hypotonic solution and then suspended and homogenized in a corn starch solution prior to a homogeneity-gradient centrifugation (Fig. 1A).^24),29) The isolated chloroplasts were spherical structures with a diameter of approximately 2 μm (Fig. 1B). The photochemical efficiency of photosystem II (PSII) in isolated chloroplasts was subsequently quantified using a Double-Modulation Fluorometer FL 3500 to determine the maximum quantum yield of PSII (Fv/Fm). To properly evaluate the PSII function, we treated whole cells and chloroplasts with 3-(3′, 4′-chlorophenyl)-1,1-dimethylurea (DCMU), which can inhibit the transfer of electrons from PSII. For both whole cells and isolated chloroplasts, the chlorophyll fluorescence yield decreased regardless of whether they were treated with DCMU (Fig. 2A). The decrease in chlorophyll fluorescence yield was smaller for isolated chloroplasts than for whole cells, indicating that the electron transport chain was less efficient in isolated chloroplasts than in whole cells. However, following the DCMU treatment, the decrease in the chlorophyll fluorescence yield was similar between isolated chloroplasts and whole cells and was much smaller than the decrease in the absence of DCMU, reflecting a significant decrease in the efficiency of the electron transport chain. Therefore, we concluded that the electron transport chain was functional, even in isolated chloroplasts. Furthermore, the Fv/Fm value determined according to the respective fluorescence decay curves did not differ significantly between isolated chloroplasts (average 0.32) and whole cells (average 0.39) (Fig. 2B). In addition, no morphological abnormalities or decreases in chloroplast fluorescence were observed after a long-term storage at 4℃ (Fig. 2C). On the basis of imaging PAM fluorometry, we analyzed the PSII activity of each isolated chloroplast during long-term storage. Stable Fv/Fm values were obtained even after 6 days of storage (Fig. 2D). These results indicate that chloroplasts isolated from schyzon can maintain a functional PSII for an extended period when stored at low temperatures, making them useful for the subsequent experiments.

Fig. 1

(Color online) Isolation of chloroplasts from schyzon. (A) The homogenized sample in a centrifuge tube was separated into two dark green layers after a homogeneity-gradient centrifugation: cell debris (upper layer) and isolated chloroplasts (lower layer). (B) Microscopy images of schyzon and isolated chloroplasts. Scale bar, 2 μm.

Fig. 2

Photosynthetic activities and morphological changes of isolated chloroplasts. (A) Normalized (Fm set to 1) fluorescence decay of whole cells and isolated chloroplasts treated with or without DCMU. Data are presented as the mean ± SD (n = 3 for whole cells and n = 4 for isolated chloroplasts). (B) Maximum quantum yield of PSII (Fv/Fm) of whole cells and isolated chloroplasts. Fv/Fm values were calculated according to the fluorescence decay of whole cells and isolated chloroplasts. Data are presented as the mean ± SD (n = 3 for whole cells and n = 4 for isolated chloroplasts), with significance determined by a two-sided Student’s t-test. (C) Fluorescence (Chl) and bright field images (scale bar, 10 μm) of chloroplasts and their magnified images (scale bar, 2.5 μm) during a 6 day storage at 4℃. Chloroplasts were visualized according to chlorophyll autofluorescence. (D) Mean Fv/Fm values during the 6 day storage at 4℃. Data are presented as the mean ± SD (n ≥ 150 for isolated chloroplasts), with significance determined by a one-way analysis of variance.

To investigate whether isolated chloroplasts are photosynthetically active in a mammalian cellular environment, we first co-cultivated CHO-K1 cells with isolated chloroplasts (1:100 ratio) for 2 days to enable CHO-K1 cells to incorporate isolated chloroplasts (Fig. 3A). During the 2 day co-cultivation, CHO-K1 cells had a higher growth rate than control CHO-K1 cells (i.e., cultivated without chloroplasts) (Fig. 3B). After the co-cultivation with chloroplasts, we performed a confocal microscopy imaging analysis. Approximately 20% of the CHO-K1 cells contained 1–3 chloroplasts at 0 day after the co-cultivation (Fig. 3B, C). When the co-cultured cells were incubated for 2 and 4 days after the co-cultivation without passaging, the number of incorporated chloroplasts per cell decreased (Fig. 3D), likely because of the subcellular digestion of chloroplasts in CHO-K1 cells or the random distribution to daughter cells after CHO-K1 cells divided. Interestingly, on day 0 after the co-cultivation, 1% of the CHO-K1 cells were revealed to be chloroplast-rich cells that had taken up seven or more chloroplasts (7–45 chloroplasts per cell) (Fig. 3D). The incorporated chloroplasts were circularly localized in the cytoplasm of CHO-K1 cells (Fig. 3E). Additionally, we detected mitochondria surrounding incorporated chloroplasts as well as subcellular space between chloroplasts and mitochondria (Fig. 3F). To more precisely analyze the subcellular localization of incorporated chloroplasts, we conducted a confocal super-resolution microscopy analysis using a DNA stain to visualize the nucleus (Fig. 4A). Some chloroplasts were localized near the nuclear outer membrane and were in contact with the nuclear outer membrane in three dimensions (Fig. 4A), but they did not penetrate the nucleus (Fig. 4B). Chloroplast DNA was detected in the center of incorporated chloroplats,³⁰⁾ suggesting that chloroplast DNA was maintained in CHO-K1 cells.

Fig. 3

Subcellular localization of incorporated chloroplasts in mammalian cultured cells. (A) Schematic illustration of the experiment conducted to examine the uptake of isolated chloroplasts by CHO-K1 cells as well as photosynthetic activities. (B) Cell proliferation rates for the normal cultivation (control) and co-cultivation with chloroplasts (co-culture) during 2 day co-cultivation. Data are presented as the mean ± SD (n = 3), with significance determined by a two-sided Student’s t-test (^*P ＜ 0.05, ^**P ＜ 0.01). (C) Fluorescence microscopy images and orthogonal view of the z-stack of CHO-K1 cells co-cultivated for 2 days with (left) and without (right) isolated chloroplasts. The cells were washed after the co-cultivation. CHO-K1 cell membranes were stained with PlasMem Bright Green (green). Chloroplasts were visualized according to chlorophyll autofluorescence (magenta). Scale bar, 20 μm. (D) Number of incorporated chloroplasts in a CHO-K1 cell at 0, 2, and 4 days after co-cultivation. The frequency distribution of the number of chloroplasts in each CHO-K1 cell at each time point was compared with the corresponding data for the 0 day by a two-sided Fisher’s exact test (^*P ＜ 0.01, n ≥ 468 CHO-K1 cells). (E) Fluorescence microscopy image of a CHO-K1 cell that incorporated more than forty isolated chloroplasts (chloroplast-rich animal cell) at 0 day after the co-cultivation. Scale bar, 20 μm. (F) Fluorescence microscopy image of mitochondria and chloroplasts in CHO-K1 cells. Mitochondria were stained with MitoBright LT Green (green) and nuclei were stained with Hoechst 33342 (blue). Chloroplasts were visualized according to chlorophyll autofluorescence (magenta). Scale bar, 20 μm.

Fig. 4

Three-dimensional (3D) analyses of incorporated chloroplasts in mammalian cells. (A) Super-resolution fluorescence microscopy image and orthogonal view of the z-stack of CHO-K1 cells that incorporated chloroplasts at 0 day after the co-cultivation. Cell membranes were stained with FM 1-43 (green), and nuclei were stained with DAPI (blue). Chloroplasts were visualized according to chlorophyll autofluorescence (magenta). Some chloroplasts maintain chloroplast DNA (white arrow). Scale bar, 5 μm. (B) 3D image of (A). A white arrow indicates the same chroloplast DNA as (A). Scale bar, 16 μm.

To examine the membrane structure in more detail, we analyzed chloroplasts incorporated into animal cells via electron microscopy. Because CHO-K1 cells with chloroplasts represented only 2%–20% of all cells (Fig. 3D), searching for cells with incorporated chloroplasts in the microscope field of view was a time-consuming and labor-intensive process. Considering the lack of intracellular organelles with chlorophyll in animal cells, electron microscopy images can be obtained efficiently if chlorophyll autofluorescence is used to detect cells with incorporated chloroplasts. Therefore, we performed a correlative light and electron microscopy (CLEM) analysis, in which fluorescence and pigments in a sample were detected using a light microscope, after which the same sample and the same area were examined using an electron microscope. The resulting CLEM images were superimposed and correlated to reveal the ultrastructure of the cell or organelle where the target molecule was localized.^31),32)

We first searched for CHO-K1 cells with incorporated chloroplasts on the basis of chlorophyll autofluorescence detected using a fluorescence microscope (Fig. 5A). The coordinates of the position of CHO-K1 cells with chloroplasts were recorded, and cells in the same position were embedded in resin and sectioned, then observed using a scanning electron microscope (Fig. 5B). Isolated chloroplasts are enclosed by a double envelope membrane and have multiple layers of thylakoid membranes, which do not form grana stacks in the chloroplasts of land plants (Fig. 5C). At 0 day after the co-cultivation, we detected two types of chloroplasts surrounded by a membrane in CHO-K1 cells; one maintained the layered structure of thylakoid membranes similar to that in an intact chloroplast (Fig. 5D), whereas the other contained partially deformed thylakoid membranes (Fig. 5E). At 2 days after the co-cultivation, we detected an increase in the space between thylakoid membranes in some chloroplasts, which also had enlarged plastoglobules³³⁾ (i.e., plastoquinone-containing lipoprotein particles present in chloroplasts under biotic stress conditions) (Fig. 5F, G). At 4 days after the co-cultivation, the layered structure of thylakoid membranes appeared to be degraded (Fig. 5H). Chloroplasts were incorporated into the subcellular vesicle and were surrounded by mitochondria (Fig. 5D, E, F), which was consistent with the fluorescence microscopy image (Fig. 3F). The incorporated chloroplast was located near the nucleus, but the chloroplast envelope did not appear to be connected to the outer nuclear membrane and did not enter the nucleoplasm (Fig. 5D).

Fig. 5

(Color online) Ultrastructural analyses of isolated and incorporated chloroplasts in mammalian cells. (A) Fluorescence microscopy image of CHO-K1 cells. Cell membranes were stained with FM1-43 (green), nuclei were stained with DAPI (blue), and chloroplasts were visualized according to chlorophyll autofluorescence (magenta). The white arrow indicates a CHO cell with chloroplasts. Scale bar, 50 μm. (B) Electron microscopy image of the same area examined using a fluorescence microscope. The black arrow indicates a CHO cell with chloroplasts. (C) Electron microscopy image of an isolated chloroplast. Scale bar, 2 μm. (D, E) Electron microscopy images of incorporated chloroplasts in a CHO-K1 cell at 0 day after the co-cultivation. Scale bar, 2 μm. (D) Chloroplast localized near the outer nuclear membrane and surrounded by mitochondria. (E) Incorporated chloroplast surrounded by mitochondria in the cytoplasm. (F, G) Electron microscopy images of incorporated chloroplasts in a CHO-K1 cell at 2 days after the co-cultivation. Scale bar, 2 μm. (F) Four incorporated chloroplasts in subcellular vesicles. (G) Chloroplast in a subcellular vesicle reveals the expansion of the space between thylakoid membranes. (H) Electron microscopy image of degenerated chloroplasts in a CHO-K1 cell at 4 days after the co-cultivation. Scale bar, 3 μm. White arrowheads indicate chloroplasts. Nu and Mt show nuclei and mitochondria, respectively.

We conducted an imaging PAM analysis to measure the effective quantum yield (φII) of the chloroplasts in CHO-K1 cells at 0, 2, and 4 days after the co-cultivation (Fig. 6). We chose a CHO-K1 cell with incorporated chloroplasts using a microscope and irradiated it with continuous actinic red light (625 nm) at 2.8 μmol m^-2 s^-1, after which φII was calculated. There were no significant differences in φII between intact chloroplasts and chloroplasts in CHO-K1 cells at 0 and 2 days after the co-cultivation (Fig. 6). Considering these findings along with the observed maintenance of thylakoid membrane morphology in incorporated chloroplasts during the first 2 days after the co-cultivation, it is highly possible that the electron transport system in chloroplasts was functional in mammalian cells at least until day 2. However, the photosynthetic activity in incorporated chloroplasts decreased significantly at 4 days after the co-cultivation, implying the chloroplasts in CHO-K1 cells were not functioning normally (Fig. 6).

Fig. 6

(Color online) Measurement of photosynthetic quantum yield of isolated chloroplasts and incorporated chloroplasts in CHO-K1 cells at 0, 2 and 4 days after the co-cultivation via imaging PAM microscopy. The y-axis represents the effective quantum yield (φII) under continuous actinic light (2.8 μmol m^-2 s^-1). Data are presented as the mean ± SD; n = 8 (areas of interest in isolated chloroplasts: I.C.), n = 9 (areas of interest in isolated chloroplasts in CHO-K1 cells at 0 day), n = 6 (at 2 days), and n = 5 (at 4 days), with significance determined by a two-sided Student’s t-test (^*P ＜ 0.05).

In this study, we incorporated intact chloroplasts isolated from algae into mammalian cells and revealed the electron transport system remained active for at least 2 days. Although the maintenance of photosynthetic activity in isolated chloroplasts in vitro has been reported, our study is the first to measure the photosynthetic activity of chloroplasts in mammalian cells. The increased cell growth rate during the co-culture period (Fig. 3D) suggests that the chloroplasts may have served as a carbon source for animal cells. Some incorporated chloroplasts were localized around the host cell nucleus, implying that the subcellular vesicle containing chloroplasts was transported close to the nucleus. In addition, many mitochondria surrounded the subcellular vesicle containing chloroplasts. Although the exchange of substances between incorporated chloroplasts and nuclei or mitochondria remains undetermined, certain pathways and processes may be active, including the retrograde signaling pathway, which sends small molecular signals from chloroplasts to the nucleus,³⁴⁾ or redox transport via malate from chloroplasts to mitochondria.^35),36) Future studies should investigate the exchange of substances between chloroplasts and animal cell organelles as well as the use of substances produced by chloroplasts in animal cells. For example, kinetic analyses may be conducted and selected substances may be tracked by incorporating isotope-labeled chloroplasts.

Our electron microscopy analysis indicated that in incorporated chloroplasts at 2 days after the co-cultivation, the thylakoid membrane structure appeared to be collapsing from within.³⁷⁾ Additionally, on day 4 after the initiation of co-cultivation, the layered structure of thylakoid membranes seemed to be completely disordered, reflecting the structural and functional degradation of chloroplasts. This suggests that the significant decrease in φII can be attributed to the complete disintegration of thylakoid membranes, resulting in a non-functional electron transport chain at 4 days after the co-cultivation.

A system for the long-term maintenance of chloroplasts within animal cells may be established on the basis of natural phenomena. For example, the photosymbiotic relationship of Paramecium bursaria depends on the temporary resistance of some algal symbionts in the digestive vacuole (DV) to lysosomes, which protects the algae from digestion.³⁸⁾ Subsequently, the algae are surrounded by a membrane derived from the DV membrane (i.e., perialgal vacuole membrane) in the acid phosphatase-inactive region within the lysosome.^38)-41) This process may be artificially recreated by producing listeriolysin O (LLO) from Listeria monocytogenes in chloroplasts to prevent digestion by phagosomes.⁴²⁾ LLO can disrupt the phagocytic vacuole to enable L. monocytogenes to enter the cytosol before being digested. A previous study showed LLO produced in Bacillus subtilis can protect against macrophage phagocytosis.⁴³⁾ It was unclear in our study whether the isolated chloroplasts incorporated in CHO-K1 cells were surrounded by membranes or internalized in organelles. This will need to be clarified in future studies. This study showed that photosystems in isolated chloroplasts remain active in vitro and even in cultured mammalian cells for several days. However, additional research is needed to determine the duration and efficiency of this photosynthetic activity. No genetic modifications were made to the host mammalian genome in this study to potentially increase the chloroplast capacity. If genes required to maintain or extend photosynthetic activities can be inserted into the CHO-K1 genome and expressed, with the encoded proteins transferred to chloroplasts, it is possible that the photosynthetic products will be compatible with host cell metabolism in mammalian cells. Indeed, carbon fixation has been achieved by modifying the metabolic pathways of non-photosynthetic organisms, such as yeast and Escherichia coli, through the synthesis and delivery of mega-base sized DNA fragments and genome editing.^44),45) In summary, we developed a top-down approach to introducing photosynthetic reactions into mammalian cells using intact chloroplasts, thereby providing the basis for future studies aimed at creating artificially photosynthetic animal cells in a relatively robust manner.

Adherent CHO-K1 cells (RCB0285) were cultured in Ham’s F12 (Wako, Cat. No. 087-08335) supplemented with 10% fetal bovine serum (FBS) at 37℃ in a 5% CO₂ atmosphere. Cells were routinely subcultured every 3 to 4 days by dilution to a cell density of approximately 2 × 10⁵ cells mL^-1 in the 100 mm dish. Cyanidioschyzon merolae strain 10D-14 cells were cultured in MA2 (Modified Allen) medium shaken at 130 rpm (TAITEC, NR-30) under continuous light (40 W/m²) at 42℃ (TOMY, CLH-301).⁴⁶⁾ Cells were maintained to keep at O.D.₇₅₀ = 0.5-3.0.

70 mL of cell culture (OD₇₅₀ = 1.0) aerated in 100 mL test tubes for 17 hours was centrifuged at 2500 rpm for 5 minutes (TOMY, MDX-310), and the supernatant was removed. The precipitate was suspended in 15 mL of 180 mM sucrose buffer (20 mM Tris-HCl, 5 mM EGTA, 5 mM MgCl₂・6H₂O, 5 mM KCl) shielded from light, and gently mixed at 40° for 1 hour. The supernatant was removed by centrifugation at 2,500 rpm for 5 minutes, 125 mL of 180 mM sucrose buffer and 1/2 tablet of cOmplete (Roche, Cat. No. 11836153001) were added, mixed with an equal volume of maize starch (20%, sterilized with ethanol) suspended in 180 mM sucrose solution, and homogenized in a Dounce Tissue Grider. The maize starch was removed by centrifugation, DNAaseI (Merck, Dn25-1g) was added to the supernatant and placed on ice for 1 h. In a 50 mL tube, Percoll mixed with 300 mM sucrose buffer (20 mM Tris-HCl, 5 mM EGTA, 5 mM MgCl₂・6H₂O, 5 mM KCl) created layers of 80% , 60%, 40% and 0%, the homogenized solution was placed on the 0% and centrifuged. The layer containing isolated chloroplasts was collected.

The CHO-K1 cells were seeded at a density of 2.4 × 10⁵ in a 35 mm glass-bottom dish (Iwaki) and pre-cultured for 12 hours. Before the co-cultivation, 4.8 × 10⁵ isolated chloroplasts were centrifuged at 2500 rpm for 5 minutes, and the supernatant was removed. The isolated chloroplasts were then resuspended in 1 mL of Ham’s F-12 with 10% FBS. The CHO-K1 cell culture supernatant was removed, and the chloroplast solution was resuspended and seeded onto the culture medium containing CHO-K1 cells. The dish was gently shaken to ensure uniform liquid distribution. Co-cultivation was then conducted in an incubator at 37℃ with 5% CO₂ for 2 days. After the 2 day co-cultivation, cells were washed with PBS (－) and the culture medium was replaced with fresh Ham’s F-12 (10% FBS). The cells were then cultured for more 4 days with daily washing and medium replacement without passaging.

To measure the growth curve during the 2 day co-cultivation, CHO-K1 and the co-culture sample with chloroplasts were plated in a 24 well plate, with each well containing 2 mL of volume at a concentration of 2.4 × 10⁵ cells/mL. Cells were harvested every day, and cell concentrations were counted using a LUNA II automatic cell counter (Logos Biosystems).

We used a dual-modulation fluorometer (Photon Systems Instruments) to assess the decay of chlorophyll fluorescence yield.⁴⁷⁾ Before measurements, the cells were kept in the dark for 10 minutes. In experiments involving the inhibition of electron transfer from Q_A to Q_B, a key component of PSII, DCMU was introduced into the sample precisely two minutes before the measurement process. The baseline fluorescence intensity was labelled Fo. After the flash pulse irradiation, which induced a reduction in Q_A, the fluorescence intensity increased rapidly to reach a maximum, referred to as Fm. Post reaching this maximum, the fluorescence decreased gradually, reflecting the ensuing electron transfer processes. Fv is the difference between Fo and Fm.

A BX53 upright microscope (Olympus) was used for brightfield imaging of chloroplasts and schyzon. A FV1200 inverted confocal microscope (Olympus) was used to image chloroplasts. The chlorophyll autofluorescence was excited with a 559 nm diode laser. To image membranes of CHO-K1 cells, Plasmem Bright Green (Dojindo) was added to the culture medium of CHO-K1 cells and incubated at 37℃ with 5% CO₂ for 5 minutes. To image mitochondria and nuclei in living CHO-K1 cells, MitoBright LT Green (Dojindo) and Hoechst 33342 (Dojindo) were added to the cell culture and incubated at 37℃ with 5% CO₂ for 30 minutes. After incubation, the dish was examined using an inverted fluorescence microscope (IX81; Olympus) equipped with a laser (405 nm for Hoechst 33342, 488 nm for Plasmem Bright Green or MitoBright LT Green and 561 nm for chlorophyll autofluorescence), a confocal scanning unit (CSU-X1; Yokogawa), and an Andor Neo 5.5 sCMOS camera (Oxford Instruments). Z-stack images were captured at 0.2 μm intervals. The images and Z-stacks were processed with ImageJ.

To image incorporated chloroplasts in CHO-K1 cells at super-resolution, a FV3000 inverted confocal microscope with FV-OSR (Olympus) was used. Before the imaging, the CHO-K1 cells were fixed with the 4% paraformaldehyde (PFA) for 15 minutes and washed twice with 1 mL of PBS (－). The nuclei and the cell membrane were stained with DAPI (Nacalai tesque) and FM1-43 Dye (Invitrogen). Following staining, a 405 nm laser was visualized for DAPI, a 488 nm laser for FM1-43, and a 561 nm laser for chlorophyll autofluorescence. Z-stack images were captured at 0.2 μm intervals. The 2D and 3D images were processed with FV31S-SW software (Olympus).

Electron microscopic observation was performed following previous report with some modifications.³¹⁾ The CHO-K1 cells at 0, 2, and 4 days after co-cultivation, which were plated on a 13 mm diameter coverslip with grids (Matsunami Glass Ind., Ltd.) were pre-fixed for 30 min at 37℃ in prewarmed PBS (－) containing 4% PFA. Nuclei and cell membranes were stained with DAPI (Nacalai tesque) and FM1-43 Dye (Invitrogen), respectively. Following the fluorescence observation, the cells were kept in PBS (－) containing 4% PFA and 2% glutaraldehyde (GA) for several days at 4℃. Isolated chloroplasts were pre-fixed for 2 hours in 20 mM sodium cacodylate buffer (pH7.2) containing 1% GA. Then, the cells and chloroplasts were post-fixed in 20 mM (for isolated chloroplast) or 50 mM (for CHO-K1 cells) sodium cacodylate buffer containing 1% osmium tetroxide (Nissin-EM) for 30 min, followed by dehydration in a graded ethanol series (25, 50, 75, 90, and 100%) for 10 min each at 23℃. The cells and chloroplasts were embedded in 100% Epon812 epoxy resin (TAAB) at 60℃ for 72 h. Embedded cells were sliced using a Leica EM UC7 ultramicrotome (Leica Microsystems) with a diamond knife (Ultra 35°; Diatome). The diamond notch knife (SYNTEK) was used to identify locations of cells.⁴⁸⁾ A series of 100 nm-thin serial sections were put on a silicon wafer. The sliced samples were sequentially stained with 0.4% uranyl acetate for 12 min and lead citrate (Sigma-Aldrich) for 3 min. After osmium coating for 3 sec using an osmium coater (HPC-1SW; Vacuum Device Co., Ltd.), electron microscopic observation was performed using a field-emission scanning electron microscope (SU8220; Hitachi High-Tech) equipped with a highly sensitive backscatter-electron detector (YAG-BSE; Hitachi High-Tech) at an accelerating voltage of 5 kV.

The photosynthetic quantum yield of chloroplasts was measured using imaging PAM microscopy (WALZ) at 25℃. The culture medium was removed from the glass bottom dish using an aspirator, leaving only the medium in the depressed area at the center of the dish. A cover glass was then placed in the center of the dish and the dish was inverted and set on the stage. The cells were observed using a 63 × objective lens. Cells that contained chloroplasts were captured within the field of view, and the area of interest (AOI) was set using Imaging WinGigE software (WALZ). After a 5-minute dark period under 625 nm pulse-modulated measuring light irradiation, the fluorescence (F) and maximum fluorescence (Fm′) levels of AOIs were measured by irradiating the cells with continuous actinic light at 2.8 μmol m^-2 s^-1 and saturation pulsed light. Additionally, fluorescence intensity (F) and quantum yield of photosystem II (φII) images for the whole field of view were acquired. The φII image was obtained using the following equation:

The blanks (Fn and Fm′n, respectively) for F and Fm′ of CHO-K1 cells without chloroplasts were measured and excluded from the data for chloroplasts incorporated into CHO-K1 cells. Therefore, the φII of the chloroplasts in CHO-K1 cells was calculated using the following equation:

We also measured samples of isolated chloroplasts only to obtain the Fv/Fm and φII. To do this, we dropped 5 μL of isolated chloroplasts, which had been light-shielded in sucrose solution at 4℃, onto a glass slide.

We thank Edanz ( https://jp.edanz.com/ac) for editing a draft of this manuscript.

R.A., Y.I., and S. Matsunaga designed the experiments. R.A., Y.I., Y.O., M.S., N.T.-K., K.T., K.S., H.M., B.G., S. Maruyama, T.T., and K.S. conducted the experiments and analyzed data. R.A. and S. Matsunaga wrote the manuscript. All authors contributed through discussions and reviewed the manuscript.

This work was supported by grants from JST CREST (JPMJCR20S6), MEXT/JSPS KAKENHI (JP20H03297 and JP22H00415) to S. Matsunaga, (JP21K06101) to T.T., (JP22H02651 and JP23H04961) to K.S., (JP22H04926) to K.T., (JP23H04962) to S. Maruyama, and JST SPRING (JPMJSP2108) to R.A.

The authors have no competing interests to declare.

Edited by Tsuneyoshi KUROIWA, M.J.A.

Correspondence should be addressed to: S. Matsunaga, Department of Integrated Biosciences, Graduate School of Frontier Sciences, The University of Tokyo, 5-1-5 Kashiwanoha, Kashiwa, Chiba 277-8562, Japan (e-mail: [email protected]).

• 1)  Nishihara, A., Tsukatani, Y., Azai, C. and Nobu, M. K. (2024) Illuminating the coevolution of photosynthesis and Bacteria. Proc. Natl. Acad. Sci. U.S.A. 121, e2322120121.
• 2)  Sato, N. (2021) Are cyanobacteria an ancestor of chloroplasts or just one of the gene donors for plants and algae? Genes 12, 823.
• 3)  Miyagishima, S. (2023) Taming the perils of photosynthesis by eukaryotes: constraints on endosymbiotic evolution in aquatic ecosystems. Commun. Biol. 6, 1150.
• 4)  Miller, T.E., Beneyton, T., Schwander, T., Diehl, C., Girault, M., McLean, R., Chotel, T. et al. (2020) Light-powered CO₂ fixation in a chloroplast mimic with natural and synthetic parts. Science 368, 649-654.
• 5)  Matsunaga, S. (2018) Planimal cells: Artificial photosynthetic animal cells inspired by endosymbiosis and photosynthetic animals. Cytologia 83, 3-6.
• 6)  Jackson, H.O., Taunt, H.N., Mordaka, P.M., Smith, A.G. and Purton, S. (2021) The algal chloroplast as a testbed for synthetic biology designs aimed at radically rewiring plant metabolism. Front. Plant. Sci. 12, 708370.
• 7)  Chen, P., Liu, X., Gu, C., Zhong, P., Song, N., Li, M. et al. (2022) A plant-derived natural photosynthetic system for improving cell anabolism. Nature 612, 546-554.
• 8)  Arnon, D.I., Allen, M.B. and Whatley, F.R. (1954) Photosynthesis by isolated chloroplasts. Nature 174, 394-396.
• 9)  Martin, W., Stoebe, B., Goremykin, V., Hansmann, S., Hasegawa, M. and Kowallik, K.V. (1998) Gene transfer to the nucleus and the evolution of chloroplasts. Nature 393, 162-165.
• 10)  Burns, J.A., Kerney, R. and Duhamel, S. (2020) Heterotrophic carbon fixation in a salamander-alga symbiosis. Front. Microbiol. 11, 1815.
• 11)  Aoki, R. and Matsunaga, S. (2021) A photosynthetic animal: A sacoglossan sea slug that steals chloroplasts. Cytologia 86, 103-107.
• 12)  Maeda, T., Takahashi, S., Yoshida, T., Shimamura, S., Takaki, Y., Nagai, Y. et al. (2021) Chloroplast acquisition without the gene transfer in kleptoplastic sea slugs, Plakobranchus ocellatus. eLife 10, e60176.
• 13)  Okabe, Y. and Matsunaga, S. (2022) Natural and artificial photosymbiosis in vertebrates. Cytologia 87, 69-72.
• 14)  Bonnett, H.T. and Eriksson, T. (1974) Transfer of algal chloroplasts into protoplasts of higher plants. Planta 120, 71-79.
• 15)  Vasil, I.K. and Giles, K.L. (1975) Induced transfer of higher plant chloroplasts into fungal protoplasts. Science 190, 680.
• 16)  Fowke, L.C., Gresshoff, P.M. and Marchant, H.J. (1979) Transfer of organelles of the alga Chlamydomonas reinhardii into carrot cells by protoplast fusion. Planta 144, 341-347.
• 17)  Wang, H., Wu, M. A. and Woo, Y. J. (2019b) Photosynthetic symbiotic therapy. Aging 11, 843-844.
• 18)  Cohen, J.E., Goldstone, A.B, Paulsen, M.J., Shudo, Y., Steele, A.N., Edwards, B.B. et al. (2017) An innovative biologic system for photon-powered myocardium in the ischemic heart. Sci. Adv. 3, e1603078.
• 19)  Özugur, S., Chávez, M.N., Sanchez-Gonzalez, R., Kunz, L., Nickelsen, J., Straka, H. et al. (2021) Green oxygen power plants in the brain rescue neuronal activity. iScience 24, 103158.
• 20)  Miyagishima, S., Wei, J.L., Nozaki, H. and Hirooka, S. (2017) Cyanidiales: Evolution and habitats. In Cyanidioschyzon merolae: A New Model Eukaryote for Cell and Organelle Biology (eds. Kuroiwa, T., Miyagishima, S., Matsunaga, S., Sato, N., Nozaki, H., Tanaka, K. et al.). Springer, Singapore, pp. 3-15.
• 21)  Miyagishima, S.Y. and Tanaka, K. (2021) The unicellular red alga Cyanidioschyzon merolae--The simplest model of a photosynthetic eukaryote. Plant Cell Physiol. 62, 926-941.
• 22)  Ohta, N. (2003) Complete sequence and analysis of the plastid genome of the unicellular red alga Cyanidioschyzon merolae. DNA Res. 10, 67-77.
• 23)  Reinbothe, C., Bakkouri, ME., Buhr, F., Muraki, N., Nomata, J., Kurisu, G. et al. (2010) Chlorophyll biosynthesis: spotlight on protochlorophyllide reduction. Trends Plant Sci. 15, 614-624.
• 24)  Takayama, Y., Inui, Y., Sekiguchi, Y., Kobayashi, A., Oroguchi, T., Yamamoto, M. et al. (2015) Coherent X-ray diffraction imaging of chloroplasts from Cyanidioschyzon merolae by using X-ray free electron laser. Plant Cell. Physiol. 56, 1272-1286.
• 25)  Kildegaard, H.F., Baycin-Hizal, D., Lewis, N.E. and Betenbaugh, M.J. (2013) The emerging CHO systems biology era: harnessing the ‘omics revolution for biotechnology. Curr. Opin. Biotech. 24, 1102-1107.
• 26)  Wurm, M.J. and Wurm, F.M. (2021) Naming CHO cells for bio-manufacturing: Genome plasticity and variant phenotypes of cell populations in bioreactors question the relevance of old names. Biotech. J. 16, 2100165.
• 27)  Takahashi, S., Ozawa, S., Sonoike, K., Sasaki, K. and Nishihara, M. (2020) Morphological and cytological observations of corolla green spots reveal the presence of functional chloroplasts in Japanese gentian. PLoS One 15, e0237173.
• 28)  Sakamoto, T., Takatani, N., Sonoike, K., Jimbo, H., Nishiyama, Y. and Omata, T. (2021) Dissection of the mechanisms of growth inhibition resulting from loss of the PII protein in the cyanobacterium Synechococcus elongatus PCC 7942. Plant Cell Physiol. 62, 721-731.
• 29)  Miyagishima, S., Itoh, R., Aita, S., Kuroiwa, H. and Kuroiwa, T. (1999) Isolation of dividing chloroplasts with intact plastid-dividing rings from a synchronous culture of the unicellular red alga Cyanidioschyzon merolae. Planta 209, 371-375.
• 30)  Suzuki, K., Ehara, T., Osafune, T., Kuroiwa, H., Kawano, S. and Kuroiwa, T. (1994) Behavior of mitochondria, chloroplasts and their nuclei during the mitotic cycle in the ultramicroalga Cyanidioschyzon merolae. Eur. J. Cell Biol. 63, 280-288.
• 31)  Jahn, K.A., Barton, D.A., Kobayashi, K., Ratinac, K.R., Overall, R.L. and Braet, F. (2012) Correlative microscopy: providing new understanding in the biomedical and plant sciences. Micron 43, 565-582.
• 32)  Yoshinaga, N., Miyamoto, T., Odahara, M., Takeda-Kamiya, N., Toyooka, K., Nara, S. et al. (2024) Design of an artificial peptide inspired by transmembrane mitochondrial protein for escorting exogenous DNA into the mitochondria to restore their functions by simultaneous multiple gene expression. Adv. Funct. Mat. 34, 202306070.
• 33)  Bréhélin, C. and Kessler, F. (2008) The Plastoglobule: A bag full of lipid biochemistry tricks. Photochem. Photobiol. 84, 1388-1394.
• 34)  Mullineaux, P.M., Exposito-Rodriguez, M., Laissue, P.P., Smirnoff, N. and Park, E. (2020) Spatial chloroplast-to-nucleus signalling involving plastid-nuclear complexes and stromules. Philos. Trans. R. Soc. Lond. B. Biol. Sci. 375, 20190405.
• 35)  Noctor, G. and Foyer, C.H. (2016) Intracellular redox compartmentation and ROS-related communication in regulation and signaling. Plant Physiol. 171, 1581-1592.
• 36)  Zhao, Y., Luo, L., Xu, J., Xin, P., Guo, H., Wu, J. et al. (2018) Malate transported from chloroplast to mitochondrion triggers production of ROS and PCD in Arabidopsis thaliana. Cell Res. 28, 448-461.
• 37)  Minamikawa, T., Toyooka, K., Okamoto, T., Hara-Nishimura, I. and Nishimura, M. (2001) Degradation of ribulose-bisphosphate carboxylase by vacuolar enzymes of senescing French bean leaves: Immunocytochemical and ultrastructural observations. Protoplasma 218, 144-153.
• 38)  Kodama, Y. and Fujishima, M. (2005) Symbiotic Chlorella sp. of the ciliate Paramecium bursaria do not prevent acidification and lysosomal fusion of host digestive vacuoles during infection. Protoplasma 225, 191-203.
• 39)  Karakashian, S.J. and Rudzinska, M.A. (1981) Inhibition of lysosomal fusion with symbiont-containing vacuoles in Paramecium bursaria. Exp. Cell Res. 131, 387-393.
• 40)  Gu, F., Chen, L., Ni, B. and Zhang, X. (2002) A comparative study on the electron microscopic enzymo-cytochemistry of Paramecium bursaria from light and dark cultures. Eur. J. Protistol. 38, 267-278.
• 41)  Kodama, Y. and Fujishima, M. (2010) Secondary symbiosis between Paramecium and Chlorella Cells. Int. Rev. Cell Mol. Biol. 279, 33-77.
• 42)  Agapakis, C.M., Niederholtmeyer, H., Noche, R.R., Lieberman, T.D., Megason, S.G., Way, J.C. et al. (2011) Towards a synthetic chloroplast. PLoS One 6, e18877.
• 43)  Madsen, C.S., Makela, A.V., Greeson, E.M., Hardy, J.W. and Contag, C.H. (2022) Engineered endosymbionts that alter mammalian cell surface marker, cytokine and chemokine expression. Commun. Biol. 5, 888.
• 44)  Gleizer, S., Ben-Nissan, R., Bar-On, Y.M., Antonovsky, N., Noor, E., Zohar, Y. et al. (2019) Conversion of Escherichia coli to generate all biomass carbon from CO₂. Cell 179, 1255-1263.e12.
• 45)  Gassler, T., Sauer, M., Gasser, B., Egermeier, M., Troyer, C., Causon, T. et al. (2020) The industrial yeast Pichia pastoris is converted from a heterotroph into an autotroph capable of growth on CO₂. Nat. Biotech. 38, 210-216.
• 46)  Toda, K., Takano, H., Miyagishima, S., Kuroiwa, H. and Kuroiwa, T. (1998) Characterization of a chloroplast isoform of serine acetyltransferase from the thermo-acidophilic red alga Cyanidioschyzon merolae. Biochim. Biophys. Acta 1403, 72-84.
• 47)  Cser, K. and Vass, I. (2007) Radiative and non-radiative charge recombination pathways in Photosystem II studied by thermoluminescence and chlorophyll fluorescence in the cyanobacterium Synechocystis 6803. Biochim. Biophys. Acta 1767, 233-243.
• 48)  Goto, Y., Takeda-Kamiya, N., Yamaguchi, K., Yamazaki, M. and Toyooka, K. (2024) Effective alignment method using a diamond notch knife for correlative array tomography. Microscopy (Oxf.) 73, 446-450.

Chinese hamster ovary

pulse amplitude modulation

Cyanidioschyzon merolae