Integrated Microalgae Analysis Photobioreactor for Rapid Strain

May 26, 2016 - A set of dark-field scattering spectra was measured using three objectives of different numerical apertures (NA) from a thin slice of a...
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Integrated Microalgae Analysis Photobioreactor for Rapid Strain Selection SoonGweon Hong,†,‡,¶ Minsun Song,†,‡,¶ Sungjun Kim,†,‡ Doyeon Bang,†,‡ Taewook Kang,†,§ Inhee Choi,∥ and Luke P. Lee*,†,‡,⊥,# †

Department of Bioengineering, ‡Berkeley Sensor and Actuator Center, ⊥Department of Electrical Engineering and Computer Sciences, and #Biophysics Graduate Program, University of California, Berkeley, California 94720, United States § Department of Chemical and Biomolecular Engineering, Sogang University, Seoul 121-742, Korea ∥ Department of Life Science, University of Seoul, Seoul 130-743, Korea S Supporting Information *

ABSTRACT: Algal photosynthesis is considered to be a sustainable, alternative, and renewable solution to generating green energy. For high-productivity algaculture in diverse local environments, a high-throughput screening method is needed to select algal strains from naturally available or genetically engineered strains. Herein, we present an integrated plasmonic photobioreactor for rapid, high-throughput screening of microalgae. Our 3D nanoplasmonic optical cavity-based photobioreactor permits the amplification of a selective wavelength favorable to photosynthesis in the cavity. The hemispheric plasmonic cavity allows intercellular interaction to be promoted in the optically favorable milieu and also permits effective visual examination of algal growth. Using Chlamydomonas reinhardtii, we demonstrated a 2-fold enhanced growth rate and a 1.5-fold lipid production rate with no distinctive lag phase. By facilitating growth and biomass conversion rates, the integrated microalgae analysis platform will serve as rapid microalgae screening platforms for biofuel applications. KEYWORDS: plasmonics, optical cavity, microalgae, bioreactor, rapid strain selection productive of valuable byproducts in a local environment11−16 implies the necessity for efficient screening methods of microalgae for individual local algal farming. Moreover, since the current genetic engineering approach for the enhancement of bioproductivity further expands strain diversity due to the imperfectness of engineering methods (i.e., lack of gene delivery methods specific for various subcellular compartments5), a screening procedure is critical in developing genetically superior strains. In this report, we describe a rapid screening platform enabling accelerated growth rates and biomass production of microalgae, which provides compelling advantages for the initial screening using a low-density algal population. A conventional microalgal screening from an algal pool (i.e., local sample and genetic mutation) involves a low-density culture in liquid-based media.15,17−19 This approach requires a long period over a few weeks and a large amount of media, thus limiting the throughput of screening. As an alternative, agarplate-based screening has been introduced to increase screening

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hotoautotrophic organisms have been playing a crucial role in capturing solar energy and converting it to chemical energy in the earth. Traditional agriculture of phototrophic plants has been vital for providing food throughout human history. Now, different types of photosynthetic byproducts are being sought from phototrophic species as a promising solution to our current energy crisis.1−8 Among phototrophic organisms, microalgae are being considered as the most promising long-term, sustainable sources of biomass and fuel energy. In addition to their rapid growth, the merits of microalgae stem from the fact that algal farming does not compete with agriculture, as it uses nonarable land and water sources that are not usable for arable crops.9 Because less than 4% of the earth’s surface area and 1% of global water10 are suitable for traditional agriculture, microalgae offer enormous potential for the generation of phototrophic byproducts if algal farming can be achieved in environments other than on arable land and water. However, current microalgal researchers have mainly focused on the initial feasibility of microalgal applications and investigated a minuscule portion of the huge biodiversity of microalgae available in nature. An estimation that noninvestigated microalgae strains will be potentially more © 2016 American Chemical Society

Received: February 1, 2016 Accepted: May 26, 2016 Published: May 26, 2016 5635

DOI: 10.1021/acsnano.6b00803 ACS Nano 2016, 10, 5635−5642

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Figure 1. Schematics of integrated microalgae analysis photobioreactor (iMAP). (a) Our iMAP in this study was arrayed with four types (PC1−PC4) of iMAP as a 96-well format. A zoom-in illustration shows the photonic cavity unit integrated with plasmonic nanoparticles on the inner wall. (b) Spectral conversion attained in the iMAP, favorable to photosynthesis pigment. (c) Illustration for optical focus of the favorably converted optical spectrum at the bottom center of the iMAP cavity. Red and yellow arrows indicate incident illumination and plasmonic scattering from the nanoparticles, respectively. Phototaxis and bounded movement of microalgae can gather microalgae at the position of the optical focus. (d,e) Photographs of accumulation of a flagellated green microalgae (Chlamydomonas reinhardtii) at the cavity bottom center. A few representative traces of single-cell swims are overlaid on the top view (e). After 3 h of ring illumination with the focus at the bottom center of the bare cavity, the population of microalgae was clearly accumulated. The diameter of the cavity was ca. 6.5 mm. (f) Pseudocolored scanning electron microscopy (SEM) image for the cryosectioned photonic cavity (PC3). The thickness of plasmonic nanoparticle layer was about 1 μm thick. The scale bar is 2 μm. (g,h) SEM and optical dark-field images for gold nanoparticle clusters embedded in a photonic cavity, respectively. The scale bar is 20 μm.

throughput.20−23 Although several operational benefits are attained, such as convenience in culturing, tracking, and retrieving individual colonies, rapid and homogeneous initial growth rate, and naked-eye examination, the solid plate culture cannot be ideal for microalgal screening because microalgae in nature grow as floating in water but not immobilized. Our integrated microalgae analysis photobioreactor (iMAP) can provide homogeneous and rapid growth of microalgae as liquidbased culture due to the unique optical properties (i.e., the enhanced excitation by the selective resonance wavelength and hemispheric geometry of plasmonic optical cavity), and the multiwall format (i.e., 96 and 384 wells) of iMAP can increase throughput of screening. The functional unit, the photonic cavity, is configured as a monolithic assembly of plasmonic nanostructures on a mesoscale hemispherical wall (Figure 1a). For rapid growth and screening of microalgae, the integrated plasmonic nanostructures and the optical geometry of the iMAP units are designed to amplify the incident light and convert it into a spectrum favorable for photosynthesis. The hemispherical cavity, as uniformly covered with the plasmonic nanostructures, can focus light selectively scattered from the individual plasmonic nanoparticles on the bottom center of the cavity. Because phototaxis and bounded movement in the cavity chamber permit a high density of microalgae at the bottom of the iMAP cavities, the favorably amplified optical energy can be effectively delivered to a population of microalgae in iMAP (Figure 1b−e).

named them as PC1, PC2, PC3, and PC4 from low to high concentration of gold ions (0.5, 1, 2, and 5 mM of HAuCl4, respectively). The overall geometry of those photonic cavities was identical by using the same volume of water solution and confinement in the ionic water lithography, but by applying various gold ion concentrations, the plasmonic nanoarchitecture could be varied on a thin layer underneath the cavity with different nanoparticle densities and optical properties (Figure 1f−h). As we aimed to generate optical characteristics favorable to photosynthesis, we tuned the optical scattering of the plasmonic nanoparticles to be resonant around 680 nm, while nanoparticle density could be varied from 500 to 3000 nm interparticle distances (details in Supporting Information). Also, for a control experiment, we fabricated a transparent cavity with the same geometry but no nanoplasmonic integration. We utilized this “bare” cavity to visualize the movement of individual microalgae and to decouple the effects of enhanced intercellular interactions and the favorably amplified light resulting from nanoplasmonic scattering. First, we characterized the spectral conversion and light focusing features of our photonic cavities through theoretical calculations and experiments. Because the gold nanoparticles embedded in the photonic cavity walls were 100−200 nm in diameter, as characterized through electron microscopy, we evaluated the spatial pattern of optical scattering from nanoparticles of this size. Our electromagnetic simulation indicated that plasmonic nanoparticles larger than 100 nm in diameter can generate an asymmetrical pattern of optical scattering that differs from ideal dipole scattering (i.e., a pattern that is perpendicular to the incident direction), as shown in Figure 2a. To experimentally examine the scattering asymmetry of the nanoparticles of photonic cavities, we compared scattering spectra through dark-field spectroscopy with different

RESULTS AND DISCUSSION In this study, we fabricated four types of photonic cavities (PCs) as arrayed in a 96-well plate by varying the concentration of gold ion in the process of ionic water lithography (see details in the Materials and Methods section). For those types, we 5636

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Figure 2. Characterization of optical properties of the photonic cavity of iMAP. (a) Spatial distribution of optical scattering generated from 150 nm gold nanosphere for the polar angle θ as a function of wavelength. This heatmap plot indicates a scattering profile different from an ideal dipole scattering. (b) Experimental demonstration for the asymmetric scattering pattern of the gold nanoparticle. A set of dark-field scattering spectra was measured using three objectives of different numerical apertures (NA) from a thin slice of a photonic cavity (PC2) as immersed in a refractive-index-matched liquid. (c) Heatmap plot for optical intensity profile inside the photonic cavity. A ray tracing was applied for 650 nm illumination with the scattering profile of the nanoparticles attained in (a). Main rays are shown as red lines in the plot. (d) Dark-field scattering spectra of the four types of photonic cavities. The absorption spectrum of photosynthetic pigments extracted from Chlamydomonas reinhardtii is overlaid as a black line. All spectra are normalized, and each maximal intensity is indicated in the legend. (e) Experimental measurement of optical power distribution along the z-axis of the cavity from the bottom center. The light from a 4000 K halogen bulb was illuminated vertically on the PC2 half-cut along the z-axis. Details of experimental setup are shown in Figure S2. (f) Optical characteristics of the four photonic cavities regarding spectral quality for photosynthetic pigments and optical power on the cellular population accumulated in the cavity center.

and absorption magnitude of photosynthesis pigment at a specific wavelength, respectively) and as optical power exposed cells residing at the bottom center of the cavity (Figure 2f). Moreover, the hemispherical optical geometry of iMAP can facilitate intercellular proximity among microalgal cells and thus accelerate initial population growth in a homogeneous manner due to the enhanced intercellular interaction within the optical cavity geometry. Our single-cell tracking of Chlamydomonas reinhardtii, a flagellated green microalgae, indicates that the iMAP optical cavities enhance the local cell density and intercellular interactions, which could be advantageous for early stage cell growth as a shortened lag phase and homogeneous growth are commonly observed in agar plate culture.20 As shown in Figure 3a,b, the swimming patterns of the flagellated microalgae were apparently different between the flat chamber and the hemispherical cavity chamber. The individual swimming patterns of microalgae in the cavity chamber were consistently confined around the geometric center, whereas those of microalgae in the flat chamber were random and not bounded. We analyzed two aspects of these different swimming patterns: the spatial frequency of cell positioning and the effective radius of the swimming trace. As expected, the spatial frequency analysis demonstrated that the cavity chamber culture resulted in a higher frequency of positioning closer to the center of the cavity, but the cells in the flat chamber displayed a random spatial distribution of positioning

numerical apertures (NA) of 0.25, 0.4, and 0.6 as NA is related to the angular range of the scattering measurement (details in Supporting Information). Based on the three NAs, we characterized scattering regions of θ < sin−1 (0.25), sin−1 (0.25) < θ < sin−1 (0.4), and sin−1 (0.4) < θ < sin−1 (0.6) as NA = n sin(θ) (n or refractive index is 1). The results in Figure 2b address that the nanoparticles can scatter incident light especially at the angular region between 120 and 140° relative to the dark-field illumination angle, which is sin−1 (0.25) < θ < sin−1 (0.4). Combined with the asymmetry pattern of the nanoparticle scattering, the optical geometry of the cavity permits strong focusing of incident light at the bottom center of the cavity, as shown in the ray optics simulation of Figure 2c. All of the iMAP optical cavities produced optical resonance similar to the absorption peak of a photosynthesis pigment at approximately 680 nm with minor variations (Figure 2d). Using an in-house scattering measurement setup equipped with a low NA optical fiber scanner (Figure S2), we confirmed that the amplified optical field is generated at the bottom center of the cavity (Figure 2e); the four types of the photonic cavities exhibited nearly identical scattering distribution with absolute maxima proportional to the dark-field scattering intensities. Based on the measured optical properties, we could define two indices for the optical environments attained inside the four types of photonic cavities, as spectral quality (∫ IPCB·Qabsdλ, where IPCB and Qabs are scattering intensity of a photonic cavity 5637

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Figure 3. Microalgal accumulation and enhanced intercellular interaction in iMAP cavity culture. (a,b) Individual swimming traces in the flat chamber and iMAP, respectively. (c,d) Heatmap plots for spatial positioning frequency in the flat chamber and iMAP, respectively. (e) Histogram for effective radii of swimming traces of individual cells. Approximately 300 cells were counted for this histogram. (f,g) Top-view photographs of short-term (up to 3 h) and long-term (up to 4 days) culture of Chlamydomonas reinhardtii in the bare cavity. The microalgae were inoculated to the cavity vessel at 3000/μL. To mimic optical conditions in the iMAP cavity, a ring illumination was placed at the top of the cavity and focused at the bottom center. A stable accumulation of the microalgae at the bottom center was attained within 3 h and maintained along with the population growth. Another example (initial cell density of 6000/μL) is shown in Supporting Information Movie S1. (h) Correlation plot between cell number and green color intensity at the cavity center. Known concentrations of cells (x-axis) were inoculated in cavities with a volume of 200 μL. Those cells were accumulated at the bottom center after 4 h.

Figure 4. Characterization of microalgal culture in iMAP cavities. (a) Change of cell number and chlorophyll content from iMAP cavities and control chambers after 24 h culture. Chlamydomonas reinhardtii, which were in the exponential stage of growth, were inoculated into the various culture chambers, and the amounts of population and chlorophyll were measured by a hemocytometer and optical density, respectively. All cases were normalized by the flat chamber culture. The lighter and darker bars in the plot present cell number and chlorophyll, respectively. The red line is the comparison between cell number and chlorophyll amount. (b) Characterization of lag phase in low-density culture. Cells in the saturation stage were inoculated into various culture chambers at a low density (i.e., 1 per 5 μL in 200 μL volume). All standard deviations were calculated from five independent cultures. (c) Histogram of single-cell characterization on lipid induction. Lipid production of Chlamydomonas reinhardtii was induced by nitrogen starvation for 2 day. The insets are confocal microscopy images, where the green and red channels are chlorophyll and lipid, respectively.

chamber (Figure 3e). As shown in the analysis, the algal cells

frequency, which may be a cause of inconsistent initial growth of low density culture (Figure 3c,d). The analysis on the effective radius (the minimum radius enclosing the entire trace for individual cells) shows that, regardless of total swimming distance, the overall movement in the cavity chambers was bounded within smaller radii than the cases observed in the flat

can be stably situated at the bottom center of the cavity, thus enhancing intercellular interactions and possibly permitting more homogeneous growth especially in a low concentration culture. 5638

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the iMAP cultures appeared to enter the exponential stage even on the first day of culture. Also, the growth rates of the iMAP culture were enhanced by more than 2-fold during the exponential stage compared to the bare cavity and flat chamber cultures, such that our naked-eye examination could observe a clear population growth at day 1.5 for the photonic cavity, whereas the bare cavity needed a 2 times longer period. We also examined the influence of iMAP microenvironments on the lipid production, which is important for biofuel applications. When a nitrogen source (i.e., NH4+) is removed from the culture medium (i.e., tris-acetate-phosphate [TAP] medium), the algal metabolism and photosynthetic products result in the accumulation of lipid bodies in the cytoplasm.30,31 Commonly, small-sized lipid bodies can be observed in the 2−3 days of induction, and along with further induction, the lipid bodies easily become as big as a half of the cell volume.31 In our nitrogen-deprived induction in the iMAP cavities, we could attain earlier formation of a lipid body than what is commonly expected. The comparison on the second day induction shows that the total amount of lipids attained in the iMAP optical cavities was 1.5- and 1.2-fold higher than ones in the flat chamber and bare cavity, respectively (Figure 4c). Also, a small portion of population in iMAP induction was found to contain a large volume of lipid bodies in the cytoplasm (Figure 4c inset ③), which is rare in the flat chamber induction. Although further validation will be required, we consider the enhanced photosynthesis and intercellular interaction as the main origin of the rapid lipid induction. As demonstrated using a green microalgae Chlamydomonas reinhardtii, the increase of growth rate, the reduction of lag phase duration, and the increase of lipid production rate can be attained in the iMAP cavity culture, which are crucial factors for initial low-population screening.

Also, the confined movement at the cavity center allows a quick naked-eye examination on population size, similarly to agar plate culture.24−26 Figure 3f,g shows the possibility of naked-eye examination for the population change of Chlamydomonas reinhardtii freshly inoculated in the cavity. Within the initial 3 h, an increased local density at the bottom center was observed, and the extent of growth was also able to be distinguished over 4 days as the series of photographs present. In order to examine the correlation between cell number and naked-eye examination, we added various numbers of cells into the cavity chambers and analyzed the green color intensity from the photographs taken after 4 h. As shown in Figure 3h, a high-level correlation was attained between the total cell number and the color intensity. As discussed, the features of the iMAP can be valuable for accelerating the initial growth of microalgae, thus decreasing the time cost of microalgal R&D stages which require handling of a small number of microalgal cells for individual screenings and selections. Blue and green wavelength light is reported to be inhibitory biomass production and cell division through the dissipation of excess excitation energy as heat or low photosynthetic quantum yield.27 Also when only a very small number of cells are available in culture, growth inhibitory factors often cause elongated lag phase randomly, thus making early characterization difficult and statistically not profound (Figure S6). As the iMAP culture can attain a photosynthesisfavorable optical condition and enhanced intercellular interactions, a rapid characterization on population growth and biomass production rates can be accomplished in a low-density screening. In order to demonstrate the advantages of iMAP in strain screening, we characterized three aspects of algal culture productivity (growth rate, lag phase duration, and biofuel production rate) by culturing Chlamydomonas reinhardtii CC124 (a wild-type strain) at a low population density. These aspects are generally considered when screening microalgal strains for biofuel applications.28,29 First, to evaluate the microalgal growth rate in the iMAP, microalgae in the exponential phase (Figure S3) were inoculated in the four types of photonic cavities, and the changes of the cell number and chlorophyll content were characterized (Figure 4a). Compared to a cylindrical flat chamber and a bare cavity, the iMAP cultures significantly enhanced growth rate and chlorophyll content by up to 2-fold. Noticeably, the PC1, PC2, and PC3 induced more chlorophyll production than population growth, indicating that chlorophyll content was strongly influenced by spectral quality than by optical power. Otherwise, in the PC4 the chlorophyll increase was smaller than population increase, which possibly indicated that an overdose of light exposure can be caused by the high-density plasmonic nanostructures upon the normal culture illumination (100 μmol m−2 s−1). Yet, with a 100-fold decrease of illumination, the highest growth rate was attained from PC4 culture, possibly by providing enough optical energy. To evaluate the lag phase duration in iMAP culture, microalgae on the fifth day of the saturation stage were inoculated to the iMAP cavities and the control vessels with fresh media. Over the next 4 days, a typical S-shaped curve growth was attained from all cases by counting cells on a microscope; typically, the microalgal growth curve consists of a lag phase, an exponential phase, and a stationary phase. Compared to a flat chamber culture, the lag phases of iMAP culture and bare cavity culture were shortened by 2.5 and 1.5 days, respectively. As shown in Figure 4b, the growth curve of

CONCLUSION In conclusion, we present an integrated microalgae analysis platform that can generate the increased growth rate, the reduction of lag phase duration, and increased lipid production rate. The photonic cavities of iMAP provide optical environments favorable to microalgal photosynthesis; as well, the hemispherical geometry enhances intercellular interaction of microalgae so that homogeneous and enhanced growth were attained in an early stage of low-population culture. The demonstration using Chlamydomonas reinhardtii shows that our iMAP can enhance metabolic activities of microalga, resulting in rapid growth and biofuel production with no distinctive lag phase. The rapid, enhanced phenotype expression attained in the iMAP culture will accelerate microalgae strain screening and also potentially play a critical role as an effective bioreactor component. MATERIALS AND METHODS Materials. Wild-type Chlamydomonas reinhardtii CC-124 (mt+) was purchased from Chlamydomonas Resource Center (University of Minnesota). TAP medium was purchased from Phyto Technology Laboratories. Silicone elastomer/silicone elastomer curing agent (Sylgard 184 silicone elastomer kit) and 96-well strip holders were purchased from Dow Corning Corporation (Midland, MI) and Corning (USA), respectively. Nile red and BODIPY 505/515 (4,4difluoro-1,3,5,7-tetramethyl-4-bora-3a,4a-diaza-s-indacene) were purchased from Invitrogen and stored as a stock solution of 0.2 mg mL−1 in acetone and 1 mg mL−1 in ethanol, respectively. All chemicals including gold(III) chloride trihydrate (HAuCl 4·3H2O) were purchased from SigmaAldrich Chemical Co. unless otherwise indicated, and autoclaved distilled water was used in all experiments. 5639

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ACS Nano Ionic Water Lithography. At the interface between different media that are immiscible with each other, the byproducts of chemical reactions can be unique, differently from those of volume reactions, in which one type of molecule reacts with different molecules in other interfacing media. As an application of interfacial chemistry, we are currently developing a new fabrication process termed ionic water lithography. By utilizing interfacial reactions during the polymerization process of a prepolymer, a monolithic array of metal nanoparticle can be created at the surface of the polymer. The detailed procedure for ionic water lithography is as follows. By dropping ionic water onto the prepolymer, a bowl-shaped geometry is created due to the combined effects of gravity, surface tension, density differences, and repulsion between the immiscible water and prepolymer. The curvature of the bowl-shaped geometry is determined by the surface tension, the volume of the ionic water droplets, and the density difference between the two immiscible media. Simultaneously, metal nanoparticles are formed and align with the surface of the bowl-shaped geometry. The formation and arrangement of these metal nanoparticles are driven by the diffusion of cations into the prepolymer as well as by charge interactions, the reducing potential of the curing agent, thermodynamics, and surface energy. The size and density of the metal nanoparticles on the surface of the bowl-shaped geometry are controlled by the concentration of ionic water, the ratio of prepolymer base to curing agent, and the temperature. Fabrication of Multiwell Photonic Cavity Bioreactor Plates. The 96-well platform-based photonic cavities were fabricated with ionic water lithography (Figure 1a). Polydimethylsiloxane (PDMS) prepolymer was prepared by thoroughly mixing base and curing agent Sylgard 184 silicone elastomer kit in a 10:1 ratio, and four different concentrations of gold chloride (HAuCl4) solutions (0.2, 1, 2, and 5 mM) were prepared from a 100 mM stock solution. Then, ca. 40 g of PDMS prepolymer was poured onto a 96-well plate frame (96 Well StripWell, Corning, USA) and degassed for approximately 30 min to remove all of the bubbles. Next, 200 μL of the gold chloride solution was dropped into each of the wells. As a result, the four types of photonic cavities were fabricated, and bare cavities were also prepared with DI water, whose density adjusted with glycerol as an experimental control. All of the reactions were performed at room temperature overnight. Then, the fabricated photonic cavities were rinsed with DI water to remove unreacted ions. Prior to all cell culture experiments, the cavities were autoclaved (121 °C, 20 min) and incubated with TAP medium at 4 °C. Characterization of Gold Nanoparticles in the Photonic Cavity. To characterize the size of the gold nanoparticles and thickness of the nanoparticle-infiltrated PDMS layer, we froze and cracked the fabricated photonic cavities and imaged Pt-coated pieces of them through a scanning electron microscope (FEI Quanta 3D FEG). Gold nanoparticles were embedded underneath the cavity surface so that SEM could not image the nanoparticles. In order to expose the plane holding the nanoparticles, we peeled off the nanoparticle-infiltrated layer by rapidly repeating freezing and thawing. Difference of thermal expansions enabled a clear separation of the nanoparticle-infiltrated layer from PDMS bulk. Also, for characterization of the thickness of the nanoparticle-infiltrated PDMS layer, a cross-sectioned photonic cavity was prepared by immersing a photonic cavity into liquid nitrogen and cracking normally to the cavity inner surface using a knife edge. Then, 2 nm of a platinum layer was deposited on the samples before electron microscopy imaging. On the SEM imaging, peeled PDMS bulk held a lot of dimple patterns, and the peeled PDMS layer matching PDMS bulk held an array of gold nanoparticle clusters that consist of 100−200 nm of gold nanoparticles. Also, the cross-sectional SEM image revealed that the thickness of the nanoparticle-infiltrated PDMS layer is 1−2 μm as we distinguished through energy-dispersive X-ray spectroscopy. Simulation Method for Electromagnetism and Ray Optics. Using commercial software (a 3D RF module of COMSOL 3.5a), an electromagnetic field generating around 100−200 nm gold nanospheres were calculated upon a spatially homogeneous illumination of 500−1000 nm. The calculation result was imported into MATLAB (a mathematical software) and postprocessed as far-field intensity

distribution in a nanoparticle-centered spherical coordinate. One of the representative results shown in Figure 2a is a heatmap plot of wavelength versus polar angle (θ) from a zenith direction. For this, a circular oblique illumination attained by the dark-field condenser was considered as incident light. For ray tracing on the photonic cavity, an in-house program was applied to trace a bundle of rays. In the calculation, PDMS and cavity volume were set with refractive indexes of 1.4 and 1.33, respectively, and the cavity surface was a scattering boundary with characteristics attained from electromagnetic simulation. Representative rays were overlaid over the optical intensity heatmap as red lines. Algal Strain and Culture Conditions. Green microalgae Chlamydomonas reinhardtii (CC-124) were cultivated in 250 mL flasks in TAP medium with adjusted pH = 7.0 ± 0.1. The algae were grown at 28 °C under continuous illumination of 100 μmol m−2 s−1 provided by white halogen lamps (4700 K). The cells were maintained continuously in the mid-exponential growth phase before experiments. Cells from the fifth day of the saturation phase were used for lag phase characterization and curvature effect experiments, and cells from midexponential growth phase were all used in the other experiment. In the case of the lipid induction experiment, the culture medium of cells in the third day of stationary phase was changed from TAP to nitrogendepleted media (TAP-N; pH 7) after centrifugation (2500 rpm, 3 min). The time point of cells that were used in each experiment are indicated in Figure S3. Algal Pigment Characterization. In order to characterize absorption spectra of algal pigments, pigments of Chlamydomonas were extracted by ethanol and the absorption spectra were measured by spectroscopic analysis. In detail, the suspension of Chlamydomonas cells in the ethanol was placed in the ice bath and sonicated (40 kHz) for 20 min as a 2 min break at every 5 min sonication in a sonicator (S4000, Misonix). After sonication, the suspension was centrifuged at 5000 rpm for 10 min, and the supernatant containing algal pigments was collected. For the absorption spectra measurement, the sample was prepared as follows. Punched PDMS with a 10 in. diameter was attached on the glass slide (3 × 1 in.), and the extracted algal pigment solution was placed within. In this experiment, PDMS with a thickness of 4 mm was used as a solution boundary to secure enough path length for absorption. The sample was covered by a cover slide (18 × 18 mm, No. 1) and mounted on the inverted microscope (same setting with the above-mentioned dark-field spectroscope except with a dark-field condenser). Then, the transmission spectra were measured with a 10× objective lens and converted to absorption spectra. Cell Population/Chlorophyll Quantification. We assessed the amount of cells by counting the number in a specific volume in a conventional or a homemade hemocytometer or by measuring chlorophyll optical density. In detail, algae population was counted in a hemocytometer by sampling 100 μL of algae suspension from the cavity and adding 25 μL of 4% formaldehyde to immobilize the algae. Each counting was conducted twice, and three samples were taken for each experiment. Also, a spectrophotometric method was used to determine the quantity of chlorophyll in algal samples. Fifty microliters of Chlamydomonas reinhardtii cells sampled from photonic cavities was mixed with 150 μL of 100% ethanol for chlorophyll extraction and incubated for 30 min in the dark at room temperature. The total volume of 200 μL was then analyzed by a spectrophotometric method (absorbance at 450 nm) in a microplate reader (Thermomax, Molecular Devices). Lipid Body Quantification. Nile red staining, which is a general fluorescent lipophilic staining method, was applied to determine the quantity of lipid bodies in algal samples. Chlamydomonas reinhardtii cells were stained with a 2 μg mL−1 solution of Nile red and incubated for 30 min in the dark at room temperature. Nile red fluorescence in neutral lipids was observed with a fluorescence microscope (Biorevo, BZ-9000, Keyence) equipped with a 20× objective lens, using a TRITC filter for 545−565 nm excitation. By being bound to neutral lipids, Nile red emits a yellow-gold fluorescence (λmax = 580 nm). Fluorescence intensity of the stained lipid body was analyzed with microscope software (BZ-H1AE, Keyence). 5640

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ACS Nano Lipid Body Imaging. For confocal imaging, cells from the photonic cavity at the second day of lipid induction stage were stained with BODIPY 505/515 at a final concentration of 10 μg mL−1 (from a stock of 1 mg/mL in ethanol), followed by 30 min incubation in the dark at room temperature. Stained cells were then fixed with 1% formaldehyde solution for 30 min, followed by rising with the TAP media. After being placed on the microscope slide, algae images were captured using a laser scanning confocal microscope (LSM710, Carl Zeiss Inc.) with a 100× oil immersion objective lens. For detection of BODIPY 505/515 fluorescence from neutral lipids, the 488 nm laser was used and emission was collected between 475 and 610 nm. Chlorophyll fluorescence was also captured using a 633 nm excitation laser, and emission was collected between 620 and 700 nm. For threedimensional imaging, Z stacks through an entire cell were acquired at 0.25 μm intervals, and each image was computationally projected using a ZEN software (Zeiss). Chlorophyll and lipid body images were also merged and pseudocolored (green, chlorophyll; red, lipid body) using ZEN software (Zeiss). Lag Phase Characterization in iMAP Cavities. In order to assess the advantage of iMAP cavity on the early stage of culture regarding lag phase, we inoculated CC-124 as single-cell dissociation in PC3 (fabricated with 2 mM HAuCl4), bare cavity, and flat chamber as a low concentration of one cell per 5 μL. As the total volume of each culture chamber was 200 μL, approximately 40 cells (±3.9) were placed in individual chambers, and a calculation shows that the intercellular distance can be 1.71 mm in the flat chamber. To minimize evaporation from the small volume along the culture duration, we covered the top of culture chamber with a thin PDMS film, gaspermeable but blocking water vapor evaporation. All the culture chambers were exposed with illumination of 100 μmol m−2 s−1 of the solar-like spectrum at 28 °C. At every 12 h for 5 days, five chambers of all cases were sampled and immediately treated with 1% formaldehyde to fix the population amount. The 100 μL volume of each sample was imaged using a fluorescence microscope (Biorevo, BZ-9000, Keyence) and counted for chlorophyll autofluorescent cells.

(4) Malcata, F. X. Microalgae and Biofuels: A Promising Partnership? Trends Biotechnol. 2011, 29, 542−549. (5) Markou, G.; Nerantzis, E. Microalgae for High-Value Compounds and Biofuels Production: A Review with Focus on Cultivation under Stress Conditions. Biotechnol. Adv. 2013, 31, 1532− 1542. (6) Schenk, P.; Thomas-Hall, S.; Stephens, E.; Marx, U.; Mussgnug, J.; Posten, C.; Kruse, O.; Hankamer, B. Second Generation Biofuels: High-Efficiency Microalgae for Biodiesel Production. BioEnergy Res. 2008, 1, 20−43. (7) Huang, G.; Chen, F.; Wei, D.; Zhang, X.; Chen, G. Biodiesel Production by Microalgal Biotechnology. Appl. Energy 2010, 87, 38− 46. (8) Spolaore, P.; Joannis-Cassan, C.; Duran, E.; Isambert, A. Commercial Applications of Microalgae. J. Biosci. Bioeng. 2006, 101, 87−96. (9) Pittman, J. K.; Dean, A. P.; Osundeko, O. The Potential of Sustainable Algal Biofuel Production Using Wastewater Resources. Bioresour. Technol. 2011, 102, 17−25. (10) Stephens, E.; Ross, I. L.; Mussgnug, J. H.; Wagner, L. D.; Borowitzka, M. A.; Posten, C.; Kruse, O.; Hankamer, B. Future Prospects of Microalgal Biofuel Production Systems. Trends Plant Sci. 2010, 15, 554−564. (11) Yang, H.-L.; Lu, C.-K.; Chen, S.-F.; Chen, Y.-M.; Chen, Y.-M. Isolation and Characterization of Taiwanese Heterotrophic Microalgae: Screening of Strains for Docosahexaenoic Acid (Dha) Production. Mar. Biotechnol. 2010, 12, 173−185. (12) Zhou, W.; Li, Y.; Min, M.; Hu, B.; Chen, P.; Ruan, R. Local Bioprospecting for High-Lipid Producing Microalgal Strains to Be Grown on Concentrated Municipal Wastewater for Biofuel Production. Bioresour. Technol. 2011, 102, 6909−6919. (13) Duong, V. T.; Li, Y.; Nowak, E.; Schenk, P. M. Microalgae Isolation and Selection for Prospective Biodiesel Production. Energies (Basel, Switz.) 2012, 5, 1835−1849. (14) Zhou, W.; Hu, B.; Li, Y.; Min, M.; Mohr, M.; Du, Z.; Chen, P.; Ruan, R. Mass Cultivation of Microalgae on Animal Wastewater: A Sequential Two-Stage Cultivation Process for Energy Crop and Omega-3-Rich Animal Feed Production. Appl. Biochem. Biotechnol. 2012, 168, 348−363. (15) Lee, K.; Eisterhold, M. L.; Rindi, F.; Palanisami, S.; Nam, P. K. Isolation and Screening of Microalgae from Natural Habitats in the Midwestern United States of America for Biomass and Biodiesel Sources. J. Nat. Sci., Biol. Med. 2014, 5, 333−339. (16) Larkum, A. W.; Ross, I. L.; Kruse, O.; Hankamer, B. Selection, Breeding and Engineering of Microalgae for Bioenergy and Biofuel Production. Trends Biotechnol. 2012, 30, 198−205. (17) Doan, T. T. Y.; Sivaloganathan, B.; Obbard, J. P. Screening of Marine Microalgae for Biodiesel Feedstock. Biomass Bioenergy 2011, 35, 2534−2544. (18) Sydney, E. B.; da Silva, T. E.; Tokarski, A.; Novak, A. C.; de Carvalho, J. C.; Woiciecohwski, A. L.; Larroche, C.; Soccol, C. R. Screening of Microalgae with Potential for Biodiesel Production and Nutrient Removal from Treated Domestic Sewage. Appl. Energy 2011, 88, 3291−3294. (19) Cole, J. K.; Hutchison, J. R.; Renslow, R. S.; Kim, Y.-M.; Chrisler, W. B.; Engelmann, H. E.; Dohnalkova, A. C.; Hu, D.; Metz, T. O.; Fredrickson, J. K.; Lindemann, S. R. Phototrophic Biofilm Assembly in Microbial-Mat-Derived Unicyanobacterial Consortia: Model Systems for the Study of Autotroph-Heterotroph Interactions. Front. Microbiol. 2014, 5, 109. (20) Pereira, H.; Barreira, L.; Mozes, A.; Florindo, C.; Polo, C.; Duarte, C. V.; Custodio, L.; Varela, J. Microplate-Based High Throughput Screening Procedure for the Isolation of Lipid-Rich Marine Microalgae. Biotechnol. Biofuels 2011, 4, 61. (21) Chan, A.; Andersen, R.; Le Blanc, M.; Harrison, P. Algal Plating as a Tool for Investigating Allelopathy among Marine Microalgae. Mar. Biol. (Heidelberg, Ger.) 1980, 59, 7−13. (22) Volk, R.-B. Screening of Microalgal Culture Media for the Presence of Algicidal Compounds and Isolation and Identification of

ASSOCIATED CONTENT S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsnano.6b00803. Additional experimental methods (PDF) Movie S1 (AVI)

AUTHOR INFORMATION Corresponding Author

*E-mail: [email protected]. Author Contributions ¶

S.H. and M.S. contributed equally to this work.

Notes

The authors declare no competing financial interest.

ACKNOWLEDGMENTS This work was supported by Korea CCS R&D Center (KCRC) grant funded by the Korea government (Ministry of Science, ICT & Future Planning) (Grant No. 2015M1A8A1053539). REFERENCES (1) Mata, T. M.; Martins, A. A.; Caetano, N. S. Microalgae for Biodiesel Production and Other Applications: A Review. Renewable Sustainable Energy Rev. 2010, 14, 217−232. (2) Ahmad, A. L.; Yasin, N. H. M.; Derek, C. J. C.; Lim, J. K. Microalgae as a Sustainable Energy Source for Biodiesel Production: A Review. Renewable Sustainable Energy Rev. 2011, 15, 584−593. (3) Sayre, R. Microalgae: The Potential for Carbon Capture. BioScience 2010, 60, 722−727. 5641

DOI: 10.1021/acsnano.6b00803 ACS Nano 2016, 10, 5635−5642

Article

ACS Nano Two Bioactive Metabolites, Excreted by the Cyanobacteria Nostoc Insulare and Nodularia Harveyana. J. Appl. Phycol. 2005, 17, 339−347. (23) Mutanda, T.; Ramesh, D.; Karthikeyan, S.; Kumari, S.; Anandraj, A.; Bux, F. Bioprospecting for Hyper-Lipid Producing Microalgal Strains for Sustainable Biofuel Production. Bioresour. Technol. 2011, 102, 57−70. (24) Tulin, F.; Cross, F. R. A Microbial Avenue to Cell Cycle Control in the Plant Superkingdom. Plant Cell 2014, 26, 4019−4038. (25) Bogen, C.; Al-Dilaimi, A.; Albersmeier, A.; Wichmann, J.; Grundmann, M.; Rupp, O.; Lauersen, K. J.; Blifernez-Klassen, O.; Kalinowski, J.; Goesmann, A.; Mussgnug, J. H.; Kruse, O. Reconstruction of the Lipid Metabolism for the Microalga Monoraphidium Neglectum from Its Genome Sequence Reveals Characteristics Suitable for Biofuel Production. BMC Genomics 2013, 14, 926. (26) Wakao, S.; Chin, B. L.; Ledford, H. K.; Dent, R. M.; Casero, D.; Pellegrini, M.; Merchant, S. S.; Niyogi, K. K. Phosphoprotein Sak1 Is a Regulator of Acclimation to Singlet Oxygen in Chlamydomonas Reinhardtii. eLife 2014, 3, e02286. (27) Schulze, P. S. C.; Barreira, L. A.; Pereira, H. G. C.; Perales, J. A.; Varela, J. C. S. Light Emitting Diodes (Leds) Applied to Microalgal Production. Trends Biotechnol. 2014, 32, 422−430. (28) Gendy, T. S.; El-Temtamy, S. A. Commercialization Potential Aspects of Microalgae for Biofuel Production: An Overview. Egypt. J. Pet. 2013, 22, 43−51. (29) Hannon, M.; Gimpel, J.; Tran, M.; Rasala, B.; Mayfield, S. Biofuels from Algae: Challenges and Potential. Biofuels 2010, 1, 763− 784. (30) Libessart, N.; Maddelein, M. L.; van den Koornhuyse, N.; Decq, A.; Delrue, B.; Mouille, G.; D’Hulst, C.; Ball, S. Storage, Photosynthesis, and Growth: The Conditional Nature of Mutations Affecting Starch Synthesis and Structure in Chlamydomonas. Plant Cell 1995, 7, 1117−1127. (31) Wang, Z. T.; Ullrich, N.; Joo, S.; Waffenschmidt, S.; Goodenough, U. Algal Lipid Bodies: Stress Induction, Purification, and Biochemical Characterization in Wild-Type and Starchless Chlamydomonas Reinhardtii. Eukaryotic Cell 2009, 8, 1856−1868.

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DOI: 10.1021/acsnano.6b00803 ACS Nano 2016, 10, 5635−5642