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NATURE PLANTS | www.nature.com/natureplants In the version of the Supplementary Information file originally published figure captions were omitted and several characters in the text were rendered incorrectly. ese errors have been corrected in this file 10 April 2015. Physical interaction between peroxisomes and chloroplasts elucidated by in situ laser analysis Kazusato Oikawa, Shigeru Matsunaga, Shoji Mano, Maki Kondo, Kenji Yamada, Makoto Hayashi, Takatoshi Kagawa, Akeo Kadota, Wataru Sakamoto, Shoichi Higashi, Masakatsu Watanabe, Toshiaki Mitsui, Akinori Shigemasa, Takanori Iino, Yoichiroh Hosokawa and Mikio Nishimura Nature Plants 1, 15035 (2015); published online 30 March 2015; corrected online 10 April 2015. CORRIGENDUM ARTICLES NATURE PLANTS DOI: 10.1038/NPLANTS2015.57 © 2015 Macmillan Publishers Limited. All rights reserved.

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In the version of the Supplementary Information file originally published figure captions were omitted and several characters in the text were rendered incorrectly. These errors have been corrected in this file 10 April 2015.

Physical interaction between peroxisomes and chloroplasts elucidated by in situ laser analysisKazusato Oikawa, Shigeru Matsunaga, Shoji Mano, Maki Kondo, Kenji Yamada, Makoto Hayashi, Takatoshi Kagawa, Akeo Kadota, Wataru Sakamoto, Shoichi Higashi, Masakatsu Watanabe, Toshiaki Mitsui, Akinori Shigemasa, Takanori Iino, Yoichiroh Hosokawa and Mikio Nishimura

Nature Plants 1, 15035 (2015); published online 30 March 2015; corrected online 10 April 2015.

CORRIGENDUM

ARTICLESNATURE PLANTS DOI: 10.1038/NPLANTS2015.57

© 2015 Macmillan Publishers Limited. All rights reserved.

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Physical interaction between peroxisomes and chloroplasts elucidated by in situ laser analysis Supplementary Information 1 Confocal laser scanning microscopy analysis White light (20 µmolm-2s-1) was used in most experiments, as this intensity was determined to be optimal for studying the relationship between peroxisomes and chloroplasts. Under these conditions, chloroplasts gathered at the cell surface, and the interaction between peroxisomes and chloroplasts was easily observed by confocal laser scanning microscopy (CLSM). Rosette leaves from 3-week-old Arabidopsis plants were gently evacuated into a syringe filled with deionised water21,49,51. Prior to observations of chloroplast relocation, leaf samples were placed in the dark for 30 minutes and then either retained in dark conditions or moved to either weak (20 µmolm-2s-1) or bright (100 µmolm-2s-1) white light for 2 hours at 23 °C. Images of the leaf palisade mesophyll cells were obtained using one-photon CLSM (LSM510, Carl Zeiss, Germany) through a 40× air objective lens (Carl Zeiss; EC Plan-Neofluar, NA 0.75). A 488 nm Ar/Kr laser was used to excite GFP and the fluorescence was detected through an emission filter BP505–550. A 543 nm He/Ne laser was used to excite RFP, and chlorophyll and their fluorescence was detected through emission filters BP560–615 and LP580, respectively. Images were obtained at 10, 20, and 30 second intervals depending on the experiment. All images were obtained as a single image. The images of F-actin and peroxisomes in Supplementary Fig. 11a under red light conditions were obtained using a different confocal system, according to the methodology of Kadota et al, 200952. Transverse sections (Supplementary Fig. 1a) were obtained by slicing the rosette leaves with razor blades and mounting them in tap water between a slide and cover slip. They were then analysed as described above. Peroxisomes and chloroplasts were obtained by bursting isolated protoplasts53 after exposing leaves to light for 2 hours. Fluorescence images (Figs. 3a,b,h and Supplementary Fig. 11g) were observed using a two-photon CLSM system (Olympus; FV300-Ix71-TP) in which a Ti:sapphire femtosecond laser oscillator (Spectra-Physics; Mai Tai, 950 nm, 80 femtosecond (fs), < 10 nJ/pulse, 80 MHz, Spectra-Physics, USA) was installed. Observations were made through a 60× water-immersion objective lens (Olympus; UPlanApo IR, NA 1.2). Light of 950 nm in wavelength was used to excite GFP and chlorophyll, and their fluorescence was individually detected through emission filters BP515–550 and LP590, respectively. Although the excitation wavelength was not tuned to the absorption maximum of chlorophyll, the autofluorescence signal from chloroplasts was obtained in the same way as described for one-photon CLSM. For time-lapse imaging, light irradiation was delivered using a LG-PS2 illuminator (Olympus) through band-pass filters (Asahi Spectra, Japan) for blue (450 ± 10 nm), green (530 ± 10 nm), red (650 ± 10 nm), or far-red (730 ± 10 nm) light. The period of time for which the effect of 2 hours light adaptation was retained in the dark was measured initially. It was

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judged that the effect of light was lost within 30 minutes of dark adaptation, based upon the appearance of the spherical form of the peroxisomes (Supplementary Fig. 7). Leaf sections were prepared as described above and kept under weak light (20 µmolm-2s-1) for 90 minutes to induce weak light responses in chloroplasts at the cell surface. They were then moved into the dark for 30 minutes to inactivate peroxisomes before irradiation with different light spectra (Fig. 3a). Images were obtained at 20 second intervals for 600 seconds so as not to cause damage to the cell. Supplementary Information 2 Adhesion analysis between peroxisomes and chloroplasts using an amplified femtosecond laser Amplified femtosecond laser pulses from a regeneratively amplified Ti:sapphire femtosecond laser system (Cyber Laser, Ifrite SP-1, 780 ± 5 nm, 230 fs, < 1 mJ/pulse, 125 Hz) were focused onto leaf palisade mesophyll cells through a 100× oil-immersion objective lens (Olympus; Plan N, NA 1.25) on a CLSM (Olympus, FV300-Ix71). A single shot of the amplified pulse was detected with a mechanical shutter (gate time: 8 ms) and delivered to the sample. The laser pulse was collimated by dual convex lenses before the microscope, and the laser focal point was tuned to the plane of the image. The diameter of the laser focal point, which is consistent with the beam waist, was estimated by the etching pattern on a carbon-doped polymer film and determined to be less than 1 µm. A leaf section and water were put in a square chamber with an area of 1×1 cm, which was fabricated in a rubber film with a thickness of 100 µm and sandwiched by cover glasses with thickness of 100 µm. The sample chamber was mounted on the microscope stage and the laser pulse was focused into the mesophyll cell of the leaf section. Shock and stress waves are generated at the central area of the laser focal point, where the multi-photon and excited state absorptions of water was induced efficiently. The time duration of the pulse was lengthened slightly in the passage between optics and sample because of slight wavelength variation of the reflective index in the spectrum of the laser pulse (780 ± 5 nm); however, this is generally a minor problem because, for a 230 fs pulse, the extent of the lengthening is expected to be less than 10 fs. After the pulse passed the objective lens, a half-wavelength (λ/2) plate and dual polarisers were used to tune the laser pulse energy to between 0 and 75 nJ/pulse. The threshold energy for cavitation bubble generation in water (Fth

W), which was estimated in the sample chamber containing pure water without the leaf section, was approximately 5 nJ/pulse, corresponding to 0.6 J/cm2 as a unit of laser fluence. This is in rough agreement with the fluence reported as a threshold of optical breakdown24, suggesting that the lengthening of pulse duration and optical aberrations caused by light propagation through the optical system were not critical. When the laser is focused into the mesophyll cell, it is difficult to estimate the pulse energy at the laser focal point, because the laser focus is disturbed by light refraction, scattering, and diffraction by organelles and cell walls. To solve this problem, we made a reliable assumption that the light energy absorbed at the laser focal point was the same as at the

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threshold conditions of cavitation bubble generation (the impulsive force generation) because the main component of light absorption and force generation is water molecules, even in mesophyll cells. Namely, the pulse energy at the laser focal point when the laser is focused into the mesophyll cell was estimated from the threshold of the cavitation bubble generation. The threshold energy detected by the power meter (Fth

Cell) was about two times larger than FthW, although the energy

slightly depends on the sample and the setting conditions. The pulse energy at the laser focal point in the mesophyll cell was compensated with a linear factor of Fth

W / FthCell. The x-y position was

controlled by a motorised microscope stage (Sigma Koki; BIOS-102T) coupled to a joy stick controller. The z position was controlled by mechanically shifting the microscope stage from the image plane. The accuracy of the x-y-z position, based on the mechanical precision of the microscope stage, was approximately 1 µm. The process of detachment was monitored by taking fluorescence and transmission images with a CLSM in which a 488 nm diode-pumped solid-state (DPSS) laser was installed. GFP in the peroxisomes was excited using the DPSS laser, and green fluorescence was detected through a band-pass filter (530–570 nm). Chloroplasts and cell walls with high light scattering abilities were monitored by measuring the transmission rate of the DPSS laser. Fluorescence and transmission images were merged (Fig. 2b). Images were obtained at approximately 1 second intervals. Since IR cut filters (> 750 nm) were additionally installed in front of the detectors for the fluorescence and transmission images, image acquisition was possible even during femtosecond laser shooting. Experiments to detach the peroxisome from the chloroplast were performed on 80 individual peroxisomes in order to estimate the probability of detachment. The distance between the peroxisome and the laser focal point was set at 5 µm in all experiments. If the distance was longer than 5 µm, it was difficult to obtain statistically significant data because of the small amount of free space around the chloroplast. At the lowest level of detachment probability (20%, light conditions using 25 nJ laser irradiation; Fig. 2c), the number of detached peroxisomes was 16, whereas, at the highest probability of detachment (91%, dark conditions using 75 nJ laser irradiation; Fig. 2c), 73 peroxisomes were detached. The relationship between Z-score and the number of detached peroxisomes is shown in Supplementary Figure 4. Across the range of detachment probabilities (20–91%: green zone in Supplementary Fig. 4), when the observed numbers of detached peroxisomes were slightly deviated from the estimated values, the Z-score rarely changed enough to affect the estimates indicated in Figure 2g. However, when the detachment experiments were performed using pulse energies lower or higher than those of the reported experimental conditions, it was difficult to obtain a reliable estimation because of large deviation of the Z-scores. In ref. 19, a method using laser-induced impulsive force to estimate adhesion strength was described. This study broke the adhesions between leukocyte–endothelial cells and epithelial cells, which are over 10 µm in diameter. The peroxisome is, in comparison, extremely small (about 1 µm in diameter) and, therefore, the current experiments were performed

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using a high-magnification (100×) objective lens instead of a 10× objective lens, as in ref. 19. By altering the objective lens, not only high-resolution observation of the peroxisome but also force loading in the small space was achieved. Although other objective lenses with different magnifications were used in preliminary experiments, only usage of the 100× objective lens enabled the collection of statistically robust evidence concerning peroxisome detachment. Supplementary Information 3 Calibration of femtosecond laser-induced impulsive force by atomic force microscopy (AFM) To confirm the magnitude of the impulsive force, the head of an AFM (Pacific Nanotechnology; Nano-R2) was mounted on the microscope stage instead of the sample. A tipless silicon AFM cantilever (NanoWorld; TL-NCH) with a glass water shield plate was attached to the AFM head. The space between the water shield plate and a glass substrate was filled with sterile water, into which the femtosecond laser was focused. The difference between the top and bottom voltages of the quadrant photo diode (QPD) was directly monitored using an oscilloscope (Tektronix; DP4104). The difference in voltage was converted to the position shift with a linear coefficient of 5.3 mV/nm, which was estimated by moving the cantilever on the glass substrate using a piezoelectric motor. The threshold of the impulsive force generation in the leaf palisade mesophyll cells (20 nJ/pulse) was about four times larger than that in distilled water (5 nJ/pulse) because the laser focus inside the mesophyll cell is disturbed by reflection, scattering, and diffraction by organelles and cell walls. Oscillations of the cantilever are induced by actions of the impulsive force, as shown in the top schematics of Supplementary Figure 5a. When a quantity of water molecules at the laser focal point (Of) is excited by a multi-photon absorption of the infrared femtosecond laser pulse (Supplementary Fig. 5a (1)), a transient vapour bubble (a cavitation bubble) is generated at Of by a pressure decrease, caused by the shock wave emission, and a temperature increase due to non-radiative relaxation of the excited states. The cantilever is pushed by a positive force resulting from a stress wave caused by the expansion of the cavitation bubble (Supplementary Fig. 5a (2)). The cantilever is then pulled back to Of by a negative force caused by a second stress wave resulting from the contraction of the cavitation bubble (Supplementary Fig. 5a (3)). In addition, the cantilever is moved by a residual jet flow following the collapse of the cavitation bubble (Supplementary Fig. 5a (4)). Previously19,20, we

applied a very simple model to our data: a single impulse (I·δ(t)) was used to estimate the total impulsive force, expressed as a unit of impulse [N-s]. In the present analysis, this model was improved to estimate the impulsive force as units of force [N] and pressure [N/m2], thus enabling comparisons with previous reports addressing protein-protein binding37-39. The positive and negative forces were hereby divided into two impulses, I1 and I2, corresponding to integrals of the positive and negative forces (middle graph, Supplementary Fig. 5a). The positive and negative impulses loaded onto the AFM cantilever are given as I1

AFM and I2AFM, respectively, and thus the motion

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equation of Z shift of the cantilever is expressed as: ( ) ( )TtItIkZZZm Δ−⋅+⋅=++ δδα AFM

2AFM12 !!! [S1]

where m is the equivalent mass of the cantilever system, Z!α is the viscous damping force, k is the spring constant at the fundamental vibrational mode, and ΔT is the time interval between I1

AFM and I2

AFM. The experimental data were analysed using this equation. Supplementary Figs. 6a-c shows the pulse energy dependencies of the bending movements of the AFM cantilever (red lines) and also the fitted results from Eq. [S1] (blue lines). The data were acquired by changing the position of Of in the direction of the optical axis (Z0 in Supplementary Fig. 5b); the distance between Of and the top of cantilever in the XY plane was fixed at 10 µm (X0 = 10 µm in Supplementary Fig. 5b). The data wave was reproduced well by a least-squares fitting of Eq. [S1], although the first sift around 0 µs was not reproduced by the previous analysis19, which assumed a single impulse without the right side second term of Eq. [S1]. The fitting parameters of impulses (I1

AFM and I2AFM) and the time

interval (ΔT) as a function of Z0 position are shown in Supplementary Figs. 6d-g. From the geometrical relationship between Of and the cantilever (Supplementary Fig. 5b), assuming that the

stress wave propagates spherically from Of, the impulse at the small fraction Δs on the AFM cantilever in the direction of optical axis is given by:

( ){ } ( )s

ZyZxXZyxX

ZIyxI or

Z Δ⋅+

⋅++

⋅+++

⋅−=Δ20

220

20

232

022

0

3021

'

1

''

1

''

'4

),(π

, [S2]

⎟⎟⎠

⎞⎜⎜⎝

+⎟⎟⎠

⎞⎜⎜⎝

⎛ −=⎟⎟

⎞⎜⎜⎝

zZX

ZX

0

0

0

0

cossinsincos

''

θθ

θθ

,

where θ is the angle of the cantilever (7°)19. The total impulses loaded on the cantilever (I1AFM and

I2AFM) were calculated from the area integration of ΔIz on the cantilever and fitted on the Z0

dependencies, as shown by the solid lines of Supplementary Figs. 6d-f. The Z0 dependencies were reproduced well only by tuning impulses at Of (I1 and I2). The fitted results show that these

assumptions are reliable. On the other hand, ΔT scarcely depended on the Z position and increased with laser pulse energy (Supplementary Fig. 6g). This result is reliable because ΔT is not a factor that depends on the geometrical relationship between Of and the cantilever. The cavitation bubble generation was observed using a CMOS high-speed camera (Photron; FASTCAM-APX RS 250K) under the present experimental conditions18, and representative images obtained using a pulse energy of 70 nJ/pulse are shown in Supplementary Figure 6i. The cavitation bubble was observed in only one frame with a gate time of 4 µs, which is the resolution limit of the high-speed camera. This

indicates that the lifetime of the cavitation bubble must be a little shorter than 4 µs. ΔT estimated using the AFM method is in agreement with this time scale. This result supports the reliability of the present analysis using AFM and the assumptions illustrated in the top schematic of Supplementary Fig. 5a. The maximum force achieved is during the generation of the positive force caused by the expansion of the cavitation bubble (bottom schemes of Supplementary Fig. 5a), as the

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maximum absolute value of the negative force is smaller than that of the positive force. Therefore, in this analysis, the maximum force (F0) at Of was estimated as:

TIF Δ= 10 [S3]

assuming the expansion and collapse times of the cavitation bubble (Texpansion and Tcollapse in

Supplementary Fig. 5a) are nearly equal to ΔT. The F0 was used as an index of force loaded on the peroxisome. Using the results shown in Supplementary Figs. 6d-g, the maximum force, F0, applied by pulses 30, 50, and 70 nJ was calculated (Supplementary Fig. 6h). The force increased linearly with the laser pulse energy [PE] and thus the relationship can be approximated as a linear function:

[ ] [ ] 6.35nJ0.28µN0 += PEF [S4]

Generally the light absorption of the femtosecond laser pulse, strongly concerned with the force generation, is not simply proportional to the input pulse energy PE. When a laser pulse with a wavelength of 780 nm is absorbed by water, whose maximum absorption wavelength is about 200 nm, the absorption during irradiation with a wavelength of 800 nm would be mainly dominated by two kinds of processes: multiphoton absorption (n-photon abospriton) and excited state absorption (plasma absorption [23]). When the pulse energy is not so high, such as the observation conditions of two-photon microscopy (typically PE < 1nJ/pulse, corresponding to < 100 mW@100 MHz), the multiphoton absorption is the main process. In these conditions, it is known that the absorbance is proportional to the nth powers of the pulse energy like the dotted blue line in supplementary Figure 6h. On the other hand, when the laser pulse energy is high enough, such as during the present experimental conditions inducing local explosion of water (typically the PE is over 10 times larger than PE on the former condition.), the multiphoton absorption from the steady state to the excited state saturates at the front of the femtosecond pulse, which arrives earlier to the water, and the latter pulse energy is absorbed by the excited state absorption from the lower excited state to the higher excited state. Since the excited absorption is a one-photon process, the light absorption increases linearly with the laser pulse energy on the condition that the multiphoton absorption saturates. Therefore, when the laser pulse energy is high enough, light absorption of the water is considered to be proportional to the laser pulse energy PE. The force F0 generated by the absorbed light energy increases linearly with PE in a simple model assumption as illustrated in supplementary figure 5a. Therefore it is reasonable that the F0 is proportional to the laser pulse energy PE in the region of the

present experiment, though F0 (PE = 0) ≠ 0. To apply the equation [S4], not only the PE in the mesophyll cell but also the slight difference of PE by the AFM measurement was compensated with by Fth

W, which was estimated by the experimental conditions for pure water in the sample chamber without a mesophyll cell, by the same procedure mentioned in the supplementary information 2. The nature of the relationship between PE and F0 is conclusively shown by the close fit of Eq. [S4] (green arrows in Supplementary Fig. 6h) to the experimental data. Since the sizes of Of (<1 µm in diameter) and the laser focal point are much smaller than 5 µm, we assume that the stress wave

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propagates spherically from Of. Hence, the pressure loaded on a peroxisome 5 µm from Of [P5µm] is estimated using:

( )]fN/nm[18.3

m54]µN[

42

020

20

m5 FFrFP ===

µππµ [S5]

which corresponds to Eq. [1] in the main text. The pressure is also indicated in Supplementary Figure 6h. As specified in the bottom part of Supplementary Fig. 5a, the main force loaded on the peroxisome is from the stress wave rather than the shock wave. The effect of the shock wave is strongly dependent on the acoustic mismatching between cytoplasm and cell membrane, which is quite small, but the effect of the stress wave within a very short time approximates to the action on a rigid body as well as on the AFM cantilever. The detachment of a peroxisome is induced at an early stage of the arrival of the stress wave (at the maximum value of the positive force), as shown in the bottom section of Supplementary Figure 5a. Therefore, we consider that the effects of reflection and scattering of the stress wave at the boundary between peroxisome and chloroplast have negligible effects on peroxisome detachment. We conducted experiments using an agarose solution to confirm that propagation of the impulsive force (shock and stress waver) is very little affected when the viscosity of water is slightly increased19. In contrast to these experimental conditions, Brownian motion of small objects in the cytoplasm of leaf palisade mesophyll cells was the same as that in water, indicating that the viscosity of cytoplasm is very similar to that of water. The practical evidence is presented in Supplementary Movie 18, in which a chloroplast in the mesophyll cell was manipulated by optical tweezers. It is known that the force to load a micro-object on the optical tweezers is pN order [37] which is a little larger than the force of Brownian motion as described in the third paragraph of the discussion section. On the other hand, as shown in supplementary figure 14, the force loaded on the peroxisome by the femtosecond laser impulsive force is over 10 nN, which is over 10,000 times larger than the driving force (optical pressure) of the optical tweezers. Conclusively, under the condition of such strong manipulation by the impulsive force, free motion of peroxisome would not be affected by the viscosity of cytoplasm. Supplementary Information 4 Inhibitor assays 3-(3,4-Dichlorophenyl)-1,1-dimethylurea (DCMU; 20 mM; Sigma Chemical) and 2,5-dibromo-3- isopropyl-6-methyl-p-benzoquinone (DBMIB; 30 mM; Sigma Chemical) were used to inhibit the photosynthetic transport of electrons through photosystem II (PSII). Dicyclohexylcarbodiimide (DCCD; 10 mM; Wako Chemicals) was used to inhibit both the formation of a proton gradient and ATP synthesis. To determine the proper concentration of inhibitors, we first examined the inhibitory effect of several different concentrations on the interaction between peroxisomes and chloroplasts. Leaf sections were immersed in inhibitors after gentle evacuation from a syringe filled with deionised water, and then kept under light conditions

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(20 µmolm-2s-1) for 1.5 hours before observations were made. To investigate the effect of disrupting actin filaments (F-actin) on the interactions between peroxisomes, mitochondria, and chloroplasts, leaf sections were prepared, as described above, and then immersed in 20 µM latrunculin B (LatB; Sigma Chemical) or cytochalacine D (CD; Sigma Chemical) for 1.5 hours before observation. We also examined the effect of higher concentrations, such as 100 µM, of LatB and CD. The inhibitory effect was confirmed by the complete disappearance of F-actin from the leaf palisade mesophyll cells of transgenic plants expressing both GFP-mTalin or GFP-fABD2 and RHP-PTS1 (Supplementary Fig. 11). Supplementary Information 5 Fig. 5d. Model for light-dependent interaction between organelles In the dark (left-hand panel), both peroxisomes (P) and mitochondria (M) are spherical and flow into the cytosol and away from the chloroplasts (C) and F-actin. This suggests that the interaction between peroxisomes, mitochondria, and chloroplasts is weakened or almost lost under these conditions (0), as is the interaction between these organelles and actin filaments (F-actin) (0). The mobility of organelles is greatly reduced, and fluctuated randomly with no autonomous motion, which would be mainly due to Brownian motion. In the light (right-hand panel), most peroxisomes show an enhanced interaction with chloroplasts under blue (B) and, more particularly, red (R) light. This is regulated by photosynthesis and would involve ATP. Peroxisome mobility is also increased during photosynthesis, as up to 30% of peroxisomes move to another chloroplasts. The interaction between peroxisomes and F-actin is also increased, causing peroxisomes to move along the F-actin to reach the chloroplasts. However, the direct interaction between peroxisomes, mitochondria, and chloroplasts is independent of F-actin. This may facilitate the interactions between the three organelles, promoting more efficient metabolite exchange and preventing the movement of peroxisomes and mitochondria from the chloroplasts. The light-dependent interactions between the three organelles and subsequent metabolite exchange would proceed via steps (0) to (3) on chloroplasts, and are regulated by photosynthesis. Supplementary Information 6 Measurement of activities of glycolate oxidase and catalase The activities of glycolate oxidase were measured following to the methodology described in Rojas et al, 201254 with minor modification. The reaction was started by addition of sodium glycolate to the assay buffer including protein extraction at 20 min and the absorbance at 440nm were recorded for every 5min. The activities of catalase were measured following to the methodology described in Shibata et al, 201355. Total proteins were extracted from the rosette leaves from 3-week-old Arabidopsis thaliana (L.) Heynh. (Columbia) following adaptation to dark or light for 3 hours after dark adaptation for 8

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hours. Supplementary Information 7 Analysis of chloroplast and peroxisome relocation Chloroplast relocation was analysed in detached leaves incubated on agar plates. Weak light (20 µmol m-2 s-1) was used to measure the accumulation response and strong light (100 µmol m-2 s-1) to measure the avoidance response21,35,49. The numbers of peroxisomes and chloroplasts at each of the top and bottom surfaces of the leaf palisade mesophyll cells were counted. Supplementary References 51. Kagawa T, et al. Arabidopsis NPL1: a phototropin homolog controlling the chloroplast

high-light avoidance response. Science 291, 2138-2141 (2001). 52. Kadota A, et al. Short actin-based mechanism for light-directed chloroplast movement in

Arabidopsis. Proc. Natl. Acad. Sci. USA 106, 13106-13111 (2009). 53. Yoo SD, et al. Arabidopsis mesophyll protoplasts: a versatile cell system for transient gene

expression analysis. Nat. Protoc. 2, 1565-1572 (2007). 54. Rojas CM, et al. Glycolate oxidase modulates reactive oxygen species-mediated signal

transduction during nonhost resistance in Nicotiana benthamiana and Arabidopsis. Plant cell 24, 336-352 (2012).

55. Shibata M, et al. Highly oxidized peroxisomes are selectively degraded via autophagy in Arabidopsis. Plant cell 25, 4967-4983 (2013).

Legends to Supplementary Movies Supplementary Movie 1. Spherical peroxisomes in dark-adapted leaf palisade mesophyll cells.

Time-lapse images were collected every 10 seconds following dark adaptation of leaves for 2 hours.

Most peroxisomes (green) show non-directinal and random motion around chloroplasts (magenta).

The images are stacked as twenty-five-speed.

Supplementary Movie 2. Peroxisomes in light-adapted leaf palisade mesophyll cells.

Time-lapse images were collected every 10 seconds following light adaptation of leaves for 2 hours.

Some peroxisomes (green) retain their interactions with a chloroplast (magenta), but others move to

different chloroplasts.

Supplementary Movie 3. Peroxisomes in dark-adapted leaf palisade mesophyll cells.

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Time-lapse images were collected every 10 seconds following dark adaptation of leaves for 12

hours. Most spherical peroxisomes (green) are distant from chloroplasts (magenta) and immobile.

Supplementary Movie 4. Femtosecond laser analysis of dark-adapted leaf palisade mesophyll

cells. Time-lapse images of peroxisomes (green) and chloroplasts (grey) from dark-adapted leaf

palisade mesophyll cells were collected every second during femtosecond laser analysis. The blue

arrows during time-lapse imaging show the time point immediately before the femtosecond laser

shot. Peroxisomes in dark-adapted cells disappear (traced with white arrow).

Supplementary Movie 5. Femtosecond laser analyses of light-adapted leaf palisade mesophyll

cells. Time-lapse images of peroxisomes (green) and chloroplasts (grey) from light-adapted leaf

palisade mesophyll cells were collected every second during femtosecond laser analysis. The blue

arrows during time-lapse imaging show the time point immediately before the femtosecond laser

shot. Peroxisomes in light-adapted cells retain their interactions with chloroplasts.

Supplementary Movie 6. High-speed imaging of peroxisomes detached after the femtosecond

laser shooting. The movie was made using a high-speed CMOS camera (Hamamatsu Photonics;

ORCA-Flash 2.5) with a frame rate of 20 fps and 1× speed. Fluorescence excitation was induced

using a mercury lamp through a band-path filter (455–485 nm). Sequential images taken

immediately after the laser shot, indicating the relationships between peroxisomes, chloroplasts, and

the laser focal point, are shown in Supplementary Fig. 3.

Supplementary Movie 7. Two-photon excitation laser scanning microscope analysis of

peroxisomes and chloroplasts in the dark. Time-lapse images of peroxisomes (green) and

chloroplasts (magenta) were collected every 20 seconds in the dark. The first (0 seconds) and last

(600 seconds) merged images in this movie are shown in Fig. 3a (D). Note that peroxisomes were

spherical and flowed among chloroplasts during this experiment. Scale bar: 10 µm.

Supplementary Movie 8. Two-photon excitation laser scanning microscope analysis under red

light irradiation. Time-lapse images of dark-adapted leaf palisade mesophyll cells were collected

under red light irradiation, as described in Supplementary Movie 7. The first and last merged

images are shown in Fig. 3a (R). Note that elliptic peroxisomes interact directly with chloroplasts

(white arrows) and move actively.

Supplementary Movie 9. Time-lapse analysis of the white sectors in var2 leaves. Time-lapse

images of peroxisomes (green) in leaf palisade mesophyll cells in the white sectors of var2 plants

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were collected every 10 seconds after light adaptation for 2 hours. The spherical peroxisomes

mainly appear in the white sectors. They are inactive and show weak interactions with chloroplasts.

The chlorophyll fluorescence was so low that it is difficult to determine the signal in this movie.

Supplementary Movie 10. Time-lapse analysis of green sectors in var2 leaves. Time-lapse

images of peroxisomes (green) and chloroplasts (magenta) in leaf palisade mesophyll cells of green

sectors in var2 plants were collected every 10 seconds following light adaptation for 2 hours.

Peroxisomes interact actively with chloroplasts and some of them move to different chloroplasts.

White circles indicate mobile peroxisomes.

Supplementary Movie 11. Interactions between F-actin and peroxisomes in dark-adapted leaf

palisade mesophyll cells. Time-lapse images of peroxisomes (magenta) and F-actin (green) were

collected every 10 seconds following dark adaptation for 2 hours. The interactions between

peroxisomes and F-actin are either weak or lost.

Supplementary Movie 12. Interactions between F-actin and peroxisomes in white

light-adapted leaf palisade mesophyll cells. Time-lapse images were collected every 10 seconds

following light adaptation for 2 hours. There is a strong interaction between peroxisomes (magenta)

and F-actin (green), and the peroxisomes change their morphology in accordance with F-actin

movement.

Supplementary Movie 13. Two-photon excitation laser scanning microscope analysis of

LatB-treated leaf palisade mesophyll cells during red light irradiation. Time-lapse images of

peroxisomes (green) and chloroplasts (magenta) were collected every 20 seconds. During red light

irradiation, dark-adapted peroxisomes gradually become elongated along the chloroplasts, but no

peroxisome movement is observed. Note that only peroxisome movement is shown because of a

problem with the filter during red light irradiation.

Supplementary Movie 14. Time-lapse analysis of three organelles in the dark. Time-lapse

images of organelles in dark-adapted leaf palisade mesophyll cells were collected every 10 seconds.

Peroxisomes (magenta) and mitochondria (green) are spherical and some flow around the

chloroplasts (grey). Note that the interaction between organelles is almost lost.

Supplementary Movie 15. Time-lapse analysis of three organelles in the light. Time-lapse

images were collected every 10 seconds in light-adapted leaf palisade mesophyll cells. Both

peroxisomes (magenta) and mitochondria (green) are elongated and move actively along

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chloroplasts (grey), producing three-organelle complexes.

Supplementary Movie 16. Effect of an actin-depolymerising drug (LatB) on the

three-organelle complex. Time-lapse images of light-adapted leaf palisade mesophyll cells treated

with LatB were collected every 10 seconds. Both peroxisomes (magenta) and mitochondria (green)

retain their interactions with chloroplasts (grey), producing three-organelle complexes even in cells

with disrupted F-actin. Immobile peroxisomes or mitochondria are seen.

Supplementary Movie 17. Time-lapse analysis of peroxisomes and chloroplasts under strong

light. Time-lapse images were collected every 10 seconds in strong light-adapted leaf palisade

mesophyll cells. Peroxisomes (green) interact tightly with two chloroplasts (magenta).

Supplementary Movie 18. Manipulation of chloroplast in mesophyll cell of Arabidopsis by

optical tweezers. A CW Nd3+: YAG laser (1064 nm, 100 mW) was focused on a chloroplast into a

mesophyll cell of Arabidopsis through a 100× objective lens (NA 1.0). The chloroplast trapped by

the optical pressure was fixed at the laser focal point, which is center of the image. Even when

whole the cells was moved by controlling the microscope stage, the trapped chloroplast was not

moved from the laser focal point.

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Supplementary Figure 1. Direct interactions between peroxisomes and chloroplasts under light conditions. (a) Peroxisomes (green) and chloroplasts (magenta) in transverse sections of light-adapted leaf palisade mesophyll cells. Arrows indicate direct interactions between peroxisomes and chloroplasts. Scale bar: 10 µm. (b) Direct interactions between peroxisomes and chloroplasts in burst-protoplast cells from leaf tissue following adaptation to white light (20 µmole m-2 s-1) for 2 hours. Scale bar: 5 µm. (c) Peroxisomes and chloroplasts after 12 hours of dark adaptation. Scale bar: 10 µm. (d) Total displacement of peroxisomes from initial interaction with the chloroplast. (e) Velocity of peroxisomes (N = 50, selected from mobile peroxisomes). (f) The activities of glycolate oxidase and catalase in dark are set to 100%. Error bars: s.d., **P < 0.05, Student’s t-test.

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Supplementary Figure 2. Distribution of peroxisomes and chloroplasts in leaf palisade mesophyll cells. (a) Peroxisomes (green) and chloroplasts (magenta) under weak (WL) and strong (SL) light conditions. WL: 20 µmole m-2 s-1 white light; SL: 100 µmole m-2 s-1 white light. Scale bar: 10 µm. (b) Numbers of peroxisomes (grey bar) and chloroplasts (white bar) in the top and bottom regions of leaf palisade mesophyll cells. W-T: weak-top; W-B: weak-bottom; S-T: strong-top; and S-B: strong-bottom. The ratio of peroxisome number to chloroplast number is shown as a black line. (c) The numbers of peroxisome and chloroplast. Results are representative of three independent experiments.

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Supplementary Figure 3. Movement of a peroxisome immediately after femtosecond laser shooting. Sequential images were extracted from Supplementary Movie 4 (a). Magenta dots indicate the laser focal point. Arrows indicate the peroxisome, which is moving away from the chloroplast. Scale bar: 10 µm

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Supplementary Figure 4. Relationship between Z-score and the number of detached peroxisomes. The relationship was estimated from measurements obtained from tests on 80 peroxisomes. Left and right arrows indicate lowest and highest probabilities of detachment probability (numbers on arrows) observed in the present experimental condition. The Z-score was evaluated over the green zone.

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Supplementary Figure 5. Laser-induced impulsive force loaded on an atomic force microscopy (AFM) cantilever and peroxisome. (a) Schematic presentation of action of the forces acting on an AFM cantilever (top schematics) and a peroxisome (bottom schematics) over the time evolution (centre graph). (b) Geometrical relationship between the laser focal point (Of) and the AFM cantilever. (c) Geometrical relationship between Of and the peroxisome.

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Supplementary Figure 6. Measurement of the laser-induced impulsive force by bending movements of the AFM cantilever. (a-c) Oscillations of an AFM cantilever induced by the impulsive force as a function of distance between Of and the cantilever in the optical axis (Z0) with pulse energies of 30 (a), 50 (b), and 70 (c) nJ. The red and blue lines indicate the empirical data and fitted results using Eq. [S1], respectively. The observed values of Z = 10, 0, and -10 µm in (b) correspond with those shown in Fig. 2e. (d-f) The Z0-position dependence of positive (I1

AFM: red dot) and negative (I2AFM: blue dot) impulses that were estimated from the

oscillations of the AFM cantilever (Eq. [S1]) with pulse energies of 30 (d), 50 (e), and 70 (f) nJ. The red and blue lines are the Z0-position dependence of the total impulse calculated using Εq. [S2]. (g) The Z0-position time interval (ΔT) between I1

AFM and I2 AFM with pulse energies of 30 (blue dots), 50 (green dots), and 70 (red

dots) nJ. The averages are indicated as broken lines with the same colour as the dots. (h) The laser pulse energy dependences of the maximum force at Of (F0: left axis) and pressure at 5 µm from Of (P5µm: right axis) calculated using Εqs. [S3] and [S5], respectively. The red dots show the calculation value based on Figs. 2d-g. The result of least-square fitting (Εq. [S4]) is shown as the red line. Green arrows indicate laser irradiation conditions when the peroxisome was detached from the chloroplast. (i) High-speed images of cavitation bubble when the laser with a pulse energy of 70 nJ was focused into water.

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Supplementary Figure 7. Morphological changes of light-adapted peroxisomes in the dark. (a) Time-lapse images of peroxisomes (green) and chloroplasts (magenta) were collected every 5 minutes in the dark following adaptation to white light (20 µmole m-2 s-1) for 2 hours (L-D). Images taken after 0, 10, 20, and 30 minutes are shown in (a). The number of peroxisomes corresponds to the numbers in (c–e). Larger images of the inset (broken line) in (a) are shown in (b). Scale bars: 10 µm. (c–e) Long-axis length (c), circularity (d), and interaction length (e) were measured for five randomly selected peroxisomes from (a). Average values for the five peroxisomes are shown as magenta closed circles and lines.

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Supplementary Figure 8. Enlarged-tracking images of peroxisomes in the dark and red light. Tracking of peroxisomes were generated using image data, as shown in Fig. 3a with a little modification of enlarged images. Position changes of peroxisomes are tracked as lines with arrowheads and marked at start (s) and final position (f). Most peroxisomes show random motion at the same position in the dark, although a few peroxisomes flow randomly over short distances. Under red light irradiation, most peroxisomes show active movement.

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Supplementary Figure 9. Peroxisome-chloroplast interactions in cells of photoreceptor mutants. (a) Peroxisomes (green) and chloroplasts (magenta) in phyAphyB (pApB), cry1cry2 (c1c2), and phot1phot2 (p1p2) plants following dark adaptation (D), 20 µmole m-2 s-1 white light (W), 30 µmole m-2 s-1 blue light (B), or 30 µmole m-2 s-1 red light (R) for 2 hours. Scale bar: 10 µm. (b–e) Long-axis length (b), circularity (c), interaction length (d), and percentage of mobile peroxisome (e). Ler: Landsberg erecta; Ws: Wassilewskija; Col: Columbia. Results are representative of three independent experiments. Error bars: s.d., *P < 0.01.

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Supplementary Figure 10. Photosynthetic regulation of the peroxisome-chloroplast interaction. (a) Peroxisomes (green) and chloroplasts (magenta) in cells treated with photosynthesis inhibitors. Lower panels show enlarged images. Scale bar: 10 µm. Time-lapse images (collected every 20 seconds) of peroxisomes in the white (b) and green sectors (c) of var2 leaves. Arrowheads: immobile peroxisomes; circles: mobile peroxisomes. (d,e) Peroxisomes and chloroplasts in the photorespiratory mutants ped2 (d) and shmt1 (e). Magnified images are shown as insets. Scale bar: 10 µm.

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Supplementary Figure 11. Effect of Latrunculin B on the peroxisomes-chloroplast interaction. (a) Peroxisomes (magenta) and actin filament (F-actin; green) under red light conditions. Arrows indicate interactions between peroxisomes and F-actin. Scale bar: 10 µm. (b) EM images of a peroxisome (P) and a chloroplast. Arrow indicates the contact site. Scale bar: 1 µm. (c) Peroxisomes (green) and chloroplasts (magenta) in 100 µM Latrunculin B or 100 µM Cytochalasin D treated cells. Asterisks indicate chloroplasts with elongated peroxisomes after treatment with each inhibitor (c-e). (d,e) Peroxisomes (magenta) and GFP-mTalin (green) (d) or GFP-fABD2 (green) (e) in LatB-treated cells. (f) Enlarged image of F-actin around chloroplasts. (g) Time-lapse images of LatB-treated cells (as in Fig. 3a). Spherical peroxisomes (0 minutes) and elongated peroxisomes with immobile (10 minutes). Scale bars: 10 µm.

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Supplementary Figure 12. Light-dependent morphological changes in mitochondria. Mitochondria (green) and chloroplasts (magenta) in cells kept in the dark or after 2 hours of adaptation to white light (20 µmol m-2 s-1). Note mitochondria are elongated with increased interactions with chloroplasts in the light but spherical and with reduced interactions with chloroplasts in the dark. Scale bar: 10 µm.

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Supplementary Figure 13. Interactions between organelles under strong light conditions. (a) Time-lapse images of peroxisomes (green) and chloroplasts (magenta) taken every 20 seconds following adaptation to strong light (100 µmol m-2 s-1). Most peroxisomes are situated between two chloroplasts. (b) Three-organelle complexes under strong light. A high number of peroxisomes (magenta) and mitochondria (green) occur between two chloroplasts (grey), resulting in the formation of three-organelle complexes (asterisks). (c) The numbers of peroxisomes (P), mitochondria (M), and chloroplasts (C) observed under each condition. Peroxisome to chloroplast (P/C) and mitochondria to chloroplast (M/C) ratios are expressed as a percentage.

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Supplementary Figure 14. Schematic presentation of concentration of the impulsive force in the peroxisome.

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