Chloroplast-containing coacervate micro-droplets as a step towards photosynthetically active membrane-free protocells† †Electronic supplementary information (ESI) available: Details of experiments, microscopy data, supplementary videos, photosynthetic assay data, zeta measurements and schematic of sample holder for microscopy. See DOI: 10.1039/c8cc01129j

Encapsulation of structurally and functionally intact chloroplasts within coacervate micro-droplets is used to prepare photosynthetically active membrane-free protocells.

and an upper layer (5 mL, 10% Percoll-PEG-BSA). 3-4 mL of the resuspended chloroplasts were loaded on top of the Percoll density column, and the samples centrifuged at 10000 x g for 5 min. Intact chloroplasts accumulated at the 45-80 % interface, and were isolated and mixed with an equal volume of grinding medium, and then pelleted by centrifugation (60 s at 1500 x g) to remove the Percoll. The sediment was washed three times by re-suspending the intact chloroplasts in grinding buffer followed by centrifugation, and then stored in the grinding buffer at 4°C.
Estimation of chlorophyll content was carried out using the Arnon method. 2 In brief, 0.5 mL of the chloroplast suspension was diluted with 10 mL of 80 % v/v acetone-water solution and the absorbance measured at 645 nm and 663 nm using a Perkin Elmer Lambda750 UV/Vis spectrophotometer. The chlorophyll content was calculated using the equation: where, A 645 and A 663 refer to the absorbance at 645 nm and 663 nm, respectively, and V total = volume of the chloroplast solution + the volume of 80 % v/v acetone-water solution.

Sequestration of chloroplasts into PDDA/CMDX coacervate micro-droplets:
Positively-charged coacervate micro-droplet dispersions were prepared by mixing 0.125 mL of poly(diallyldimethylammonium chloride) (PDDA, 400 mM) and 0.45 mL of carboxymethyl-dextran (CMDX, 370 mM), and the volume was made up to 1 mL using grinding buffer. The coacervate phase was isolated by centrifugation and redispersed in the grinding buffer maintaining the volume of the dispersion. The chloroplast suspension (0.45 mg mL -1 chlorophyll, 37.5 µL) was added directly or after dilution to 0.5 mL with the grinding buffer to a coacervate micro-droplet dispersion in grinding buffer (0.5 mL). The dispersions were then placed on a vortexer for 20 minutes, and then observed using bright field and fluorescence microscopy. 3D stacks of fluorescence images were acquired using confocal microscopy to establish the inclusion of chloroplasts into the micro-droplets. To enable visualization of the coacervate phase using fluorescence/confocal microscopy, 5% of FITC tagged CMDX was added to the coacervate mixture.
Electrostatically mediated interactions between the coacervate micro-droplets and chloroplasts were imaged in a custom-built sample holder ( Supplementary Fig. S16). 10 µL of positively charged PDDA/CMDX coacervate micro-droplets (monomer mole ratio PDDA : CMDX = 0.3 : 1) or negatively charged PDDA/CMDX coacervate micro-droplets (monomer mole ratio PDDA : CMDX = 1 : 20) were introduced into one side of the sample holder and allowed to settle onto the PEGylated glass coverslip. After ca. 5 minutes, 10 µL of the chloroplast dispersion were injected from the other side of the sample chamber, and the coacervate droplets then imaged during mixing with the chloroplasts.
Photosynthetic activity of chloroplasts: Assays were undertaken on dispersions of chloroplast-containing PDDA/CMDX coacervate micro-droplets (16.8 or 22.4 µg mL -1 chloroplasts; 1 mL, 50 mM PDDA; 165 mM CMDX) in the presence of DPIP (22.5,30,45 or 60 µM) by aliquot analysis of samples after a series of brief exposures (10 s) to 40 kilolux of light (Osram Ultra-Vitalux, 300 W). The reaction dispersions were stored in the dark on an ice bath between light exposures. Aliquots (100 µL) were mixed with sodium chloride solution (200 µL, 2 M) to disassemble the coacervate phase to avoid scattering interference associated with the coacervate dispersions. Reduction of oxidized DPIP was monitored by UV-visible spectroscopy at intervals of 10 s using the time-dependent decrease in the absorption intensity measured at 620 nm. Contributions to the absorbance at 620 nm arising from the chloroplasts were corrected for by reference to the chloroplast absorption at 850 nm where DPIP showed no absorbance. All assays were carried out in glass vials treated with PEGylated silane.
Assays were also undertaken on buffered dispersions of free chloroplasts (30 µg mL -1 ), or in the presence of individual coacervate components (50 mM PDDA or 50 mM CMDX), by adding DPIP (25 µM) and exposing the chloroplast suspensions to brief exposures (10 s) to 40 kilolux of light. Reduction of DPIP was monitored by UV-vis spectroscopy at intervals of 10 s. Different concentrations of DPIP (7.5, 10, 22.5 or 60 µM) were investigated at a constant chloroplast concentration of 5.8 µg mL -1 .
Degradation/stability assays were carried out by batch analysis of samples stored in the dark at 4 °C and analyzed at 48 h intervals for residual photosynthetic activity. Activity assays were performed on buffered dispersions of chloroplasts-containing PDDA/CMDX coacervate micro-droplets (11.8 µg mL -1 chloroplasts; 1 mL, 50 mM PDDA; 165 mM CMDX) and on free chloroplasts (11.0 µg/mL) in the presence of DPIP (22.5 µM) by aliquot analysis of samples after three 30s exposures to 40 kilolux of light (Asahi Spectra Max-303; Visible bandwith). The reaction dispersions were stored on ice between exposures to the light.

Acoustic patterning of chloroplast-containing PDDA/CMDX coacervates micro-droplets:
The chloroplast-containing coacervate micro-droplet array was prepared in a custom-built acoustic trapping device with a square arrangement of four piezoelectric transducers (Noliac, NCE 51, L15 x W2 x T1 mm). The opposing transducer pairs were wired in parallel, driven by two signal generators (Agilent 33220a-001), and each connected to an oscilloscope (Agilent DSOX2014A). A glass coverslip was attached with adhesive to the bottom of the device. The two-orthogonal acoustic standing waves were generated from opposing transducer pairs operating at 6.703/6.717 MHz (10 V). The device chamber was filled with 800 µL of buffer and then 75 µL of a mixture containing freshly prepared coacervate micro-droplets and chloroplasts (50 mM PDDA; 165 mM CMDX (5% FITC-CMDX); 9.4 µg/mL chloroplasts) was added to it and mixed gently with a pipette. After ca. 10 min, the supernatant containing very small coacervate microdroplets and uncaptured chloroplasts was exchanged with buffer. 3D stacks of fluorescent microscopy images of the chloroplast-containing coacervate protocell arrays were recorded to determine the number distribution of chloroplasts per coacervate micro-droplet in the patterned arrays.
Determination of DPIP partition coefficient in PDDA/CMDX coacervates: 1 mL of a PDDA/CMDX coacervate dispersion (50 mM PDDA, 165 mM CMDX) was centrifuged, and 100 µL of the supernatant replaced with DPIP solution (0.3 mM) and vortexed for 1 min to redisperse the coacervate phase in the presence of DPIP. The coacervate dispersion was then allowed to age for a few hours before analyzing the supernatant and bulk coacervate phase for the amount of DPIP partitioned between the two phases using UV-visible spectroscopy. Aliquots of the supernatant (200 µL) and coacervate phase (20 µL) were mixed with 200 µL of phosphate buffer (0.1 M, pH 7.5) and 200 µL of sodium chloride solution (1.5 M) for the UV-Vis measurements.

PEGylation of coverslips and vials.
The vials and cover slips were treated with piranha solution (3 parts sulfuric acid and 1 part hydrogen peroxide) for about 1 hour, washed thoroughly with water and blow dried. A solution of PEGsilane in toluene (2% v/v) was used for surface functionalization. After about 1 hour, the vials and cover slips were rinsed with ethanol and blow dried.
Zeta potential measurements. All measurements were performed using a Malvern Zetasizer Nano-ZS instrument equipped with an internal Peltier temperature controller. The measurements were carried out in a disposable zeta cuvette at 25 °C.

Supplementary videos
Supplementary video 1. Optical microscopy video showing electrostatically-mediated capture of a single negatively charged chloroplast by a positively charged PDDA/CMDX coacervate microdroplet. The video is shown in real time.

Supplementary video 2.
Optical microscopy video showing the gradual re-structuring of the surface of a positively charged PDDA/CMDX coacervate micro-droplet on contact with a negatively charged chloroplast exhibiting conformal wetting behaviour. The video is shown in real time.

Supplementary video 3.
Optical microscopy video showing the absence of interaction between a negatively charged chloroplast and a negatively charged PDDA/CMDX coacervate microdroplet. Even after multiple collisions of the chloroplast with the coacervate micro-droplet there was no interaction or attachment observed. The video is shown in real time.                 S17. Sample holder used for optical and fluorescence microscopy imaging. Two cover slips were adhered to one side of a microscope slide with a channel or space between them, which was then enclosed by fixing a PEGylated cover slip on top using a UV-responsive glue. The coacervate micro-droplet dispersions were introduced into the channel from one of the open sides, and the sample holder was inverted to allow the coacervates to settle onto the PEGylated coverslip. The chloroplast dispersion was then introduced from the other side of the chamber and the interaction between the two dispersions observed using optical and fluorescence microscopy.