Photosynthetic physiology and biomass partitioning in ...

Algal Research 18 (2016) 51–60

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Photosynthetic physiology and biomass partitioning in the model diatom Phaeodactylum tricornutum grown in a sinusoidal light regime Denis Jallet, Michael A. Caballero, Alessandra A. Gallina, Matthew Youngblood, Graham Peers ⁎ Department of Biology, Colorado State University, Fort Collins, CO 80523, USA

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Article history: Received 12 February 2016 Received in revised form 11 April 2016 Accepted 15 May 2016 Available online xxxx Keywords: Phaeodactylum tricornutum Cultivation Biomass characterization Photosynthetic physiology

a b s t r a c t Photosynthetic microbes respond to changing light environments to balance photosynthetic process with light induced damage and photoinhibition. There have been very few characterizations of photosynthetic physiology or biomass partitioning during the day in mass culture. Understanding the constraints on photosynthetic efficiency and biomass accumulation are necessary for engineering superior strains or cultivation methods. We observed the photosynthetic physiology of nutrient replete Phaeodactylum tricornutum growing in light environments that mimic those found in rapidly mixing, outdoor, low biomass photobioreactors. We found little evidence for photoinhibition or non-photochemical quenching in situ, suggesting photosynthesis remains highly efficient throughout the day. Cells doubled their organic carbon from dawn to dusk and a small percentage – around 20% – of this carbon was allocated to carbohydrates or triacylglycerol. We conclude that the self-shading provided by dense culturing of P. tricornutum inhibits the induction of photodamage, and energy dissipation processes that would otherwise lower productivity in an outdoor photobioreactor. © 2016 The Authors. Published by Elsevier B.V. This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/).

1. Introduction The rotation of the earth causes sunlight levels to vary in predicable patterns of night and day with gradual increases and decreases in photosynthetically active radiation (PAR). Environmental variables like cloud cover, sunflecks, gradual vertical mixing in natural aquatic systems impose rapid changes in PAR that are superimposed on the aforementioned day/night cycles [1,2]. Turbulent mixing in turbid photobioreactors has similar effects. Understanding the physiological response to fluctuations in light fluxes is important for parameterizing models of photosynthetic productivity and for providing directions for improvements of overall biomass yields [3,4]. Maintaining high photosynthetic efficiency is essential for high biomass yields and for the production of valuable storage compounds such as biofuel precursors and lipid-based nutraceuticals. Stramenopile algae, including diatoms, are excellent candidates for the production of biofuel precursors and lipid-based nutraceuticals because they can accumulate carbon as triacylglycerols (TAGs), particularly during periods of nutrient deprivation [4]. However, little is known about the Abbreviations: AET, alternative electron transfer; chla, chlorophyll a; chlc1 + c2, chlorophyll c1 and c2; ePBR, environmental photobioreactor; FIRe, fluorescence induction and relaxation; NPQ, non-photochemical quenching; PAM, pulse amplitude modulated; PAR, photosynthetically active radiation; PSII, photosystem II; RLC, rapid light curve; ROS, reactive oxygen species; TAG, triacylglycerol; TN, total nitrogen; TOC, total organic carbon. ⁎ Corresponding author. E-mail address: [email protected] (G. Peers).

effects of a natural light regime on biomass accumulation, biomass partitioning and photosynthesis in diatoms. Net photosynthesis, the amount of carbon fixed after correction for respiration, increases linearly with light at low light fluxes but saturates at high light fluxes [5]. At low PAR fluxes, the capacity to perform carbon fixation reactions is greater than the supply of light energy, which results in “light-limited” rates of photosynthesis. However, the absorption of light energy that is in excess of photosynthetic capacity at high light fluxes can lead to reactive oxygen species (ROS) production such as singlet oxygen (1O2), hydrogen peroxide (H2O2), superoxide anions (O− 2 ) and hydroxyl radicals (OH•). These ROS may oxidize any surrounding protein, pigment or lipid, resulting in damage to the photosynthetic system [6,7]. The D1 protein of photosystem II (PSII) is a major target for photooxidative damage [8] and it has rapid turnover rates in algae such as the diatom Phaeodactylum tricornutum (hereafter Phaeodactylum) [9]. Following its damage, it is removed from PSII reaction centers and replaced by a newly synthesized, fully functional D1 subunit. Photoinhibition occurs when the rate of photodamage to D1 exceeds the rate of D1 removal/replacement [10,11]. This is more likely to happen under very strong irradiances or from other abiotic stresses [12]. Damage to D1 can be measured as an overall reduction in photosynthetic capability and also at the level of PSII maximal quantum yield (Fv/ Fm) [8]. Photoinhibition reduces photosynthetic productivity by not only reducing the efficiency of light energy transduction to organic carbon, but also in the energetic costs associated with repair to D1 and other damaged components of the cell.

http://dx.doi.org/10.1016/j.algal.2016.05.014 2211-9264/© 2016 The Authors. Published by Elsevier B.V. This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/).

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D. Jallet et al. / Algal Research 18 (2016) 51–60

In order to avoid energetically costly damage to the cell, eukaryotic algae have evolved a series of defense mechanisms. These defense mechanisms are controlled at the physiological level and at the level of genetic regulation [13]. For instance, Phaeodactylum has a high capacity for the Non-Photochemical Quenching (NPQ) of light energy as heat when shifted from low light to high light [14,15]. This process requires xanthophyll pigment cycling [14,16] and the LHCX (Light Harvesting Complex X) proteins [17,18]. Transcripts associated with NPQ are rapidly induced in response to a sudden increase in irradiance [19]. Another photoprotective strategy employed by algae consists in mobilizing Alternative Electron Transport (AET) pathways that remove excess reductant from thylakoid membranes [20]. AET – including the Mehler reaction – can consume up to 50% of the electrons released by PSII in diatoms [21]. While these processes are essential for avoiding photoinhibition, they also represent significant energy losses. Most physiological characterizations of Phaeodactylum and other stramenopiles like Nannochloropsis have been carried out in either constant light or in day/night cycles where the transitions between light and dark are a simple step function for light intensity [22–24]. However, these studies may not capture the dynamics of photosynthetic efficiency associated with growth in natural light regimes. A key aspect for future biofuel production is to characterize algal performance in industrially relevant conditions [3]. Unfortunately, large-scale experiments in outdoor systems are time consuming and they require resources that are not readily available to the research community at large. Recently developed photobioreactors can re-create fluctuating light environments in controlled laboratory conditions [25– 27] and these have been used to study the effects of a sinusoidal light regime and/or changing temperatures on the growth and biomass production of Nannochloropsis [27] and Chlamydomonas [28]. Here we report our characterization of Phaeodactylum photophysiology and the partitioning of carbon over the course of a sinusoidal light cycle in conditions that mimic a low-biomass photobioreactor. We found a linear accumulation of total cellular carbohydrate and TAG over the course of the light period and very little evidence for photoinhibition throughout the day. Our results suggest Phaeodactylum efficiently balances high rates of photosynthesis with photoprotective strategies to maintain high biomass production during rapid and regular fluctuations in light intensity. 2. Materials and methods 2.1. Culture conditions and sampling design P. tricornutum CCAP 1055/1 was grown axenically in silicon-free Instant Ocean artificial seawater (salinity 35‰). Nutrients were added according to the stoichiometry described in [29] but at 2.3 fold higher concentrations to avoid nutrient limitation during our experiment. Cultures were maintained in early exponential growth phase by serial dilutions. Day/night cycle experiments were carried out in Phenometrics ePBR photobioreactors (see Lucker et al. [25] for a full description and Fig. S1 for an annotated photograph) containing 500 mL medium. We replaced the supplied polycarbonate vessels with custom glass vessels (see Fig. S1). Cultures were grown at 18 °C, with bubbling (1 L air L−1 culture min−1) and with stirring (500 rpm). We measured pH values ranging between 8.0 at dawn and 8.8 at dusk on a representative culture. Light was supplied in a 12 h:12 h light:dark cycle. The white LED lights (see Fig. 2 from Lucker et al. [25] for light spectrum) were programmed to provide a sinusoidal light regime, reaching a maximal intensity of 2000 μmol photons m− 2 s− 1 6 h after dawn according to the following equation: Incident Light Flux(t) = Amax * sin (2* π* f * t).where t = s after dawn, Amax = 2000 μmol photons m− 2 s−1, f = 1/(2*DayLength[s]) = 1.16 × 10−5 s−1. Light intensity was measured on top and at the bottom of the water column over a representative sampling day, by inserting a 4π light

meter probe (Walz Universal Light Meter). See Fig. 1 for light distribution. Cells were acclimated to these conditions for approximately 5 generations (5 day:night cycles), diluted into fresh medium and grown for 2 more days before data collection. Time series data were collected during one diurnal cycle starting at dawn. Samples were harvested between 0 h and 0.5 h after dawn (dawn), between 3 h and 3.5 h after dawn (mid-morning), between 6 h and 6.5 h after dawn (mid-day), between 9 h and 9.5 h after dawn (mid-afternoon) and between 12 h and 12.5 h after dawn (dusk). 2nd dawn samples were collected 24 h after the first harvest.

2.2. Cell counts Cell counts and cell specific chlorophyll autofluorescence were measured using a BD Accuri C6 flow cytometer. Culture samples were diluted (100 μL culture in 900 μL medium) and passed through 30 μm filters to remove debris. Flow cytometry was carried out at a flow rate of 35 μL min−1 and a core size of 16 μm. Phaeodactylum cells were quantified after gating from forward scatter and relative fluorescence emission (excitation at 488 nm, emission detection at N 670 nm).

2.3. Total organic carbon (TOC) and Total nitrogen (TN) content of cells Cellular TOC was measured using a Shimadzu TOC-L Laboratory Total Organic Carbon Analyzer. Samples were prepared based on manufacturer's instructions (Shimadzu Application News no.049), with some modifications. Cells (1.0–2.0 × 107 cells, determined as described above) were collected from the culture and immediately frozen at −80 °C. At the same time, media blanks were collected by filtering an equal volume of culture through 0.2 μm pore size nylon filters. The filtrate was frozen at −80 °C. Samples were thawed on the day of analysis and 4 mL sample was diluted with de-ionized water (18.2 MΩ) to a final volume of 40 mL in acid-washed TOC glass vials. An injection volume of 100 μL was used and only replicate measurements with 2 successive values with b10% difference were used to calculated carbon content. The total amount of carbon (TC) in cells was calculated as: TCcells = TCsample − TCblank. 50 mM HCl was added to purge inorganic carbon from the solution and inorganic carbon in cells was calculated as: ICcells = ICsample − ICblank. Finally TOC in cells was calculated by: TOCcells = TCcells − ICcells. TN was determined from: TNcells = TNsample − TNblank. Standard curves were based off of potassium hydrogen phthalate, sodium carbonate/bicarbonate and potassium nitrate for TC, IC, and TN, respectively.

Fig. 1. Sine light regime measured at the top (open diamonds) and at the bottom (grey squares) of a 23 cm deep culture of Phaeodactylum. Illumination was provided from an LED bank situated at the top. Black/white bars above the graph indicate night/day respectively.


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2.4. Chlorophyll a (chla) and chlorophyll c1 + c2 (chlc1 + c2) extraction and quantification

standard curve of tripalmitin using ImageJ, version 1.48 (http:// imageJ.nih.gov/ij/).

Phaeodactylum cells were collected by centrifugation at 10,000 ×g, 18 °C, for 15 min. The supernatant was carefully discarded and the pellet was frozen at −20 °C. Pellets were thawed, resuspended in 1 mL methanol, vortexed for 15 s, incubated in the dark for 15 min and centrifuged at 20,000 ×g, 18 °C, for 10 min. The pigment-containing supernatants were transferred to 1 cm path-length acrylic cuvettes. Absorbance spectra were recorded using a Cary 60 UV–Vis Agilent spectrophotometer in scan mode (300–800 nm, data interval 1 nm, scan rate of 600 nm min−1). Chlorophyll concentrations were determined based on the published extinction coefficients for chlorophyll a (chla) and chlorophyll c (chlc1 + c2) in methanol, according the equations presented in [30].

2.7. Total carbohydrates quantification

Phaeodactylum cells (approximately 5.0 × 107 cells) were collected by filtration through 0.8 μm pore size polycarbonate membranes and kept at −80 °C. Cells were subsequently resuspended in 3 mL chilled (4 °C) protein extraction solution made of 10 mM sodium phosphate buffer pH 7.4, 1 mM ethylenediaminetetraacetic acid (EDTA), 1 mM phenylmethylsulfonyl fluoride (PMSF) and 0.2% (v/v) Tween 20 [31]. Cells were lysed by sonication with a Qsonica probe sonicator (3 × 30 s, 50% power, samples kept 30 s on ice between each event). Cell debris were removed by centrifugation at 10,000 × g, 4 °C, for 10 min. Soluble proteins in the supernatant were transferred to a 1.5 mL tube and then kept on ice. Protein concentration was determined with the Pierce BCA Protein Assay kit with bovine serum albumin standards.

Phaeodactylum cells (1.0 × 108 cells) were harvested by centrifugation at 3000 × g, 4 °C, for 15 min. The supernatant was discarded and the pellet was frozen at − 80 °C. Total carbohydrates quantification was performed according to [35], with some modifications. Pellets were resuspended in 1.0 mL de-ionized water (18.2 MΩ). Samples were transferred to glass vials and then 250 μL 72% (w/w) sulfuric acid was added for hydrolysis. Glass vials were capped with 20 mm lyophilization stoppers and vortexed thoroughly. After 1 h of incubation at room temperature, samples were diluted using 6 mL de-ionized water. Glass vials were sealed with 20 mm tear-off aluminium caps and heated for 1 h at 121 °C at 0.83 bar pressure. Samples were cooled to room temperature and then 2 mL was transferred to 50 mL conical tubes and calcium carbonate was used to neutralize the pH to 6–8. This solution was filtered through 0.2 μm pore size nylon filters to remove precipitates. 500 μL of the filtrate was transferred to a glass vial along with 250 μL of 0.5 M NaOH, 250 μL of 3 mg mL−1 3-Methyl-2hydrazone hydrochloride benzothiazolinone hydrate (MBTH) and 250 μL of 1 mg mL−1 dithiothreitol. Vials were capped, vortexed thoroughly and incubated at 80 °C for 15 min. 500 μL of a 0.5% (w/v) ammonium iron(III) sulfate, 0.5% (w/v) sulfamic acid and 0.25 M HCl solution was added. Samples were cooled at room temperature for 15 min, diluted with 1250 μL de-ionized water and transferred to 1 cm acrylic cuvettes. Absorbance at 620 nm was measured using a Cary 60 UV–Vis Agilent spectrophotometer in single read mode. Total carbohydrates concentrations in samples were calculated relative to a standard curve of glucose. Values are reported as pg glucose equivalents per cell.

2.6. TAGs extraction and quantification

2.8. Chlorophyll fluorometery — estimating in situ physiology

Cultures (~5.0 × 107 cells) were harvested by filtration onto a 0.8 μm pore size polycarbonate membrane. Cells were resuspended into 2 mL medium and then pelleted by centrifugation at 20,000 × g, 18 °C, for 15 min. The supernatant was discarded, the pellet was frozen at − 80 °C. Pellets were thawed and lyophilized using a Savant Speed Vac SC110 concentrator (set to high speed) connected to a refrigerated Savant RVT100 Vapor Trap. Lyophilized pellets were ground into a fine power with stainless steel beads in a Qiagen Tissue Lyser II (1/30 frequency for 1 min). Lipid extractions were performed as described in [32], with some modifications. Ground cells were resuspended in 3 mL 95% n-hexane, transferred to 5.5 mL, screw-capped, glass vials and further homogenized by sonication on ice with a Qsonica probe sonicator (3 × 30 s, 70% power). Samples were then heated to 65 °C for 1 h 30 min and then cooled to room temperature for 1 h. Debris were removed by gravity filtration through 30 μm filters and the filtrate was evaporated to dryness under nitrogen. Neutral lipids were isolated by Solid Phase Extraction (SPE) based on the protocol described in [33], with some modifications. First, dried material was resuspended in 300 μL 95% n-hexane. SPE was performed on Bond Elut NH2 columns, following the manufacturer's instructions. TAGs were eluted with 3 mL of a 80:20:1 n-hexane:diethyl ether: acetic acid mixture. The elutate was evaporated under nitrogen and frozen at −80 °C. Quantification of TAGs was based on the thin layer chromatography method developed by [34]. Samples were resuspended in 200–700 μL nhexane and 10 μL of lipid sample was deposited onto HPTLC Silica gel 60F254 plates. The mobile phase was a 70:30:1 n-hexane:diethyl ether:acetic acid. After migration, plates were dipped in a 0.05% (w/v) primulin, 80:20 acetone:water solution for 1 min. The plates were then air-dried for 10 min. Images of lipid bands were captured with a Biorad E/Z Gel Doc EZ imager (UV illumination, exposure time 15– 30 s). TAG content was quantified by band intensity relative to a

Cells were collected from the ePBR photobioreactors and directly transferred into 125 mL Erlenmeyer flasks. Sub-samples (200 μL) were diluted with 800 μL cooled (18 °C) f/2 medium and transferred to a 1 cm quartz cuvette. Chla fluorescence parameters – including the maximum quantum yield of PSII (Fv/Fm) and the functional absorption cross-section of PSII (σPSII, Å2 quantum−1) – were immediately recorded using a Satlantic FIRe Fluorometer (excitation at 450 nm, emission measured at 678 nm) [36]. σPSII was calculated from raw data using Fireworx (https://sourceforge.net/projects/fireworx/). Total time from sampling to measurement was 15 s. Remaining cells were placed in a Percival Intellus incubator and maintained at 18 °C under 20 μmol photons m−2 s−1 white light for 30 min, which allows for progressive NPQ recovery [37,38]. Chlorophyll fluorescence parameters were measured as at time zero. In situ NPQ was calculated as described in [38]: NPQ = (Fm30 − Fm)/Fm, where Fm is the maximal fluorescence level measured in the sample immediately after harvesting from the ePBR and Fm30 is the maximal fluorescence level measured in the sample after 30 min under low light.

2.5. Protein extraction and quantification

2.9. Combined pulse amplitude modulated (PAM) fluorometry and oxygen evolution measurements for ex situ rapid light curves (RLCs) We used a Walz DUAL-PAM 100 fluorometer equipped with a Walz, ED-101US/MD cuvette holder and a FireSting OXROB10 probe connected to a FiresSting Optical Oxygen Meter for the simultaneous measurement of chlorophyll fluorescence and oxygen evolution/consumption. A 1 cm diameter, cylindrical quartz cuvette was sealed with a custom plastic stopper that held the oxygen probe in the cell suspension (Fig. S2). This system permitted rapid string of the cell suspension and control of temperature using a recirculating Fisher Scientific Isotemp water bath. All steps were performed at 18 °C. Rapid Light Curves (RLCs) were performed based on [39] with some modifications.

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~ 1.00 × 108 Phaeodactylum cells were centrifuged at 3000 × g for 10 min. The supernatant was carefully discarded. Cells were gently resuspended in f/2 medium supplemented with 20 mM HEPES buffer pH 7.5 and 5 mM sodium bicarbonate (from a freshly prepared 1 M stock solution). 1.5 mL of cells equal to chla concentrations of 4 μg mL−1 was transferred to the cuvette. Dark respiration rates were recorded for 5 min before turning on the PAM fluorometer actinic red light (635 nm) at a fluence of 5 μmol photons m − 2 s− 1 . This low light pre-treatment relaxes NPQ [37]. After 15 min the actinic light was turned off, a red measuring light (620 nm) was turned on and a saturating pulse (600 ms, 10,000 μmol photons m − 2 s− 1 , 635 nm) was applied to estimate the PSII maximum photochemical yield (Fv/Fm). Consequently, cells were illuminated for 1 min steps at increasing intensities (5, 18, 33, 91, 168, 227, 390, 620, 971, 1513, 2325 and 3507 μmol photons m− 2 s− 1 ). Chla fluorescence parameters including Fv/Fm, NPQ and 1-qL were calculated as described elsewhere [40,41]. Gross oxygen evolution rates were calculated based on chla content, as the sum of net oxygen evolution rates and absolute values of dark respiration rates. The maximal photosynthesis rate (Pmax), the irradiance at saturation (Ek) and the light limited slope (α) was calculated from oxygen traces using the equations of [42].

3. Results & discussion 3.1. Light regime over a day-night cycle We programmed incident light to change as a sinusoidal wave over the course of the day. Light also varied from the surface of the illuminated culture to the bottom (Fig. 1). The cells experienced a complex light environment due to the day/ night cycle and also due to rapid vertical mixing from the constant stirring and sparging [25]. Our experiments started with approximately 2 × 106 Phaeodactylum cells mL−1, a denser population than in natural habitats even during diatom blooms [43]. This led to self-shading effects and attenuation of light intensity through the 23 cm depth of the culture (Fig. 1, Fig. S1). Light attenuation with depth is also observed in largescale photobioreactors or open ponds [44]. Additionally, the accumulation of biomass and cell pigments throughout the day increased the attenuation of light in the photobioreactor leading to a slightly asymmetrical pattern of light at the base of the culture (Fig. 1). Therefore our conditions mimicked light environments like those experienced by cells in a low density, well-mixed large-scale photobioreactor where cells are mixed turbulently through a broad gradient of light. Our conditions did not mimic conditions found in very dense cultures where cells will move rapidly between light and dark environments.

2.10. SDS-page and Western blots

3.2. Growth and carbon partitioning over a day-night cycle

Total protein extracts (see Section 2.5) were thawed and 50 μL sample was transferred to a 1.5 mL Eppendorf tube with 50 μL denaturing sample buffer containing 125 mM Tris HCl buffer pH 6.8, 20% (v/v) glycerol, 4% (w/v) SDS and 0.0025% (w/v) Bromophenol Blue. The solution was heated at 65 °C for 5 min and subsequently cooled to room temperature. All proceeding steps were carried out at room temperature. Samples corresponding to 0.75 μg total protein were loaded into a Novex 10–20% Tris-Glycine Gel. Gel electrophoresis was carried out in a XCell SureLock Mini-Cell System, following the manufacturer's instructions and employing a Tris-Glycine SDS Running Buffer containing 25 mM Tris Base pH 8.3, 192 mM Glycine and 0.1% (w/v) SDS. After migration, proteins were transferred to Invitrolon PVDF Membranes in a XCell II Blot Module, following manufacturer's instructions with a transfer buffer containing 12 mM Tris pH 8.3, 96 mM glycine and 20% (v/v) methanol. Membranes were blocked overnight at 4 °C in TBS buffer with 5% (w/v) non-fat dry milk. Antibodies recognizing the D1 protein of PSII (Agrisera, #AS0584) and the beta subunit of ATP synthase, which recognized both the mitochondrial and chloroplastic forms (ATPB, Agrisera, #AS0585), were diluted 1:10,000 and 1:50,000, respectfully, in TBS with 0.5% non-fat dry milk. The membranes were challenged for 1 h. Membranes were washed three times for 5 min in TBS with 0.5% non-fat dry milk at room temperature. Membranes were then challenged with secondary antibody (horseradish peroxidase conjugated donkey anti-rabbit, Pierce, #31,458), diluted 1:50,000 in TBS with 0.5% non-dry fat milk for 1 h. Cross-reacting bands were visualized with SuperSignal West Femto Maximum Sensitivity Substrate (ThermoScientific), following the manufacturer's instructions. Relative band image intensities were quantified with ImageJ software, version 1.48 (http://imageJ. nih.gov/ij/). D1 protein abundance was calculated after normalization to ATPB content and relative to 2nd Dawn.

Phaeodactylum grew at an exponential rate of 0.83 ± 0.06 day− 1 (n = 6). We observed a general trend for Phaeodactylum cells to divide approximately once per day, with cell division beginning in the afternoon and continuing during the night (Fig. 2) as has been reported previously under square wave light/dark cycles [23,24,45]. Cell division coincided with the peak of cellular carbon and nitrogen contents (Fig. 2, Fig. 3A & B). Cells accumulated Total Organic Carbon (TOC) throughout the light phase (Fig. 3A), increasing from 4.18 ± 1.12 pg C cell− 1 at dawn to 9.89 ± 1.47 pg C cell−1 at dusk (n = 5–6, p b 0.05). TOC per cell linearly increased during the day, displaying similar values to those measured elsewhere [23,46]. Cells also accumulated Total Nitrogen (TN), from 1.48 ± 0.34 pg N cell−1 (n = 6) at dawn to 2.10 ± 0.33 pg N cell− 1 (n = 6) at dusk (Fig. 3B). We directly measured the cellular abundance of carbohydrates, proteins as well as TAGs and could calculate the quantity of C stored in these various pools at different timepoints (Fig. 3, Table 1). However, we did not measure the abundance of other key cellular components – such as structural lipids [47] or nucleic acids – and therefore did not perform a complete C balance analysis.

2.11. Statistical analysis of data A repeated measures one-way ANOVA (p = 0.05) followed by a Tukey HSD test was used to test for significant differences between time points (SigmaPlot). All values are reported as averages ± the standard deviation.

Fig. 2. Change in cell density over a day/night cycle. Data show the results from 6 individual experiments. Black/white bars above the graph indicate night/day respectively. Each symbol type represents data from a single culture.


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Fig. 3. Accumulation of organic carbon, total nitrogen, and biomass partitioning over a day/night cycle. A) Total Organic Carbon (TOC) B) Total Nitrogen (TN) C) Total carbohydrates (pg glucose equivalent cell−1) D) Triacylglycerols (pg tripalmitin equivalent cell−1) E) Proteins and F) Chlorophyll a (chla). Data are the averages of n = 6 (A, B), n = 4 (C, E, F) or n = 3 (D) biological replicates. Error bars show 1 standard deviation. Different letters indicate statistically different groups. Black/white bars above the graphs indicate night/day respectively.

Cellular carbohydrate content steadily rose after mid-morning, ranging from 0.78 ± 0.04 pg glucose equivalents cell−1 at mid-morning to 2.53 ± 0.25 pg glucose equivalents cell− 1 at dusk (Fig. 3D, n = 4, p b 0.05). But, carbohydrates represented a small proportion of TOC, from around 5% at dawn to 10% at dusk (Table 1). Diatoms use β-1,3 glucans of the chrysolaminarin family to store part of the chemical energy and reducing power generated by photosynthesis [48–50]. Diatoms also incorporate various hexoses and pentoses directly into their cell walls [51]. While we did not separate storage vs. structural carbohydrates using our methodology, we hypothesize that the ratio of storage:structural carbohydrates increased from dawn to dusk to store photosynthate for metabolism at night [49,50]. Our observations of carbohydrate are lower than previously observed for this species [23,34, 52]. This may be due to our utilization of an improved MBTH-based vs.

phenol-sulfuric acid or anthrone based assays which can overestimate carbohydrate content [35]. Diatoms can also store some of the reducing power and chemical energy generated by photosynthesis in TAGs, particularly under nutrient deprivation [34,47,52]. The cellular content of TAGs followed a similar pattern to carbohydrates (Fig. 3D). It increased during the day, from 0.04 ± 0.02 pg tripalmitin equivalents cell−1 at dawn to 1.01 ± 0.36 pg tripalmitin equivalents cell−1 in the middle of the afternoon (n = 3, p b 0.05). TAGs represented 0.7% of TOC at dawn and 7% of TOC at dusk respectively (Table 1). Phaeodactylum cells accumulated TAGs and carbohydrates during the day not only to fuel cell divisions but also metabolism at night. Algae can consume 1–22% of their cellular biomass at night [53]. Cell density increased by an average of 150% over the course of the night in

Table 1 Carbon distribution across several biomass components over a day/night cycle. Carbon (C) represents 40% of glucose (C6H12O6) mass, 76% of tripalmitin (C51H9806) mass and C is 44% of protein mass in diatoms [34] (n = 3–4, ±standard deviation, different letters indicate statistically different averages between timepoints). Timepoint

Total C (pg·cell−1)

C in carbohydrates (pg·cell−1)

C in TAGs (pg·cell−1)

C in proteins (pg·cell−1)

Dawn Mid-morning Mid-day Mid-afternoon Dusk 2nd Dawn

4.18 ± 1.25a 6.16 ± 1.95ab 7.38 ± 1.12b 9.77 ± 0.28c 9.89 ± 1.61c 5.72 ± 0.92ab

0.32 ± 0.03a 0.31 ± 0.01a 0.52 ± 0.07b 0.75 ± 0.12c 1.01 ± 0.1d 0.30 ± 0.07a

0.03 ± 0.02a 0.12 ± 0.07ab 0.50 ± 0.25bc 0.77 ± 0.27c 0.65 ± 0.35c 0.02 ± 0.00a

3.56 ± 0.38a 3.94 ± 0.12ab 4.51 ± 0.33ab 4.98 ± 0.14b 4.50 ± 0.15ab 3.65 ± 0.13a

The quantity of carbon present in each pool was calculated based on data presented in Fig. 3.

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our experiments. If there was an even distribution of cellular compounds between cells without respiratory consumption then we would expect cellular TAGs and carbohydrates to decrease by ~ 33% from dusk to 2nd dawn. TAGs were clearly expended in excess of 33% over the course of the night (Fig. 3D) and transcripts associated with β-oxidation reactions are known to increase during the night [23]. These observations are consistent with the role of TAGs as energy and carbon storage molecules. Total carbohydrates per cell also decreased more than would be expected from cell division alone, from 2.53 ± 0.25 pg cell−1 at dusk to 0.74 ± 0.18 pg cell−1 at 2nd dawn (Fig. 3C). This drop could not entirely be explained by an even redistribution between daughter cells after division, suggesting that glycolysis consumed some of the chrysolaminarin accumulated during the day [50]. Similar observations have been made for carbohydrate accumulation and consumption in Spirulina platensis (now Arthrospira platensis) growing in outdoor tubular photobioreactors [54]. Proteins constituted a major C sink, representing N 45% of TOC at all sampled timepoints (Table.1). This has also been observed in other carbon budgets of Phaeodactylum [34,52]. The protein content of the cells only changed by ~ 20% throughout the day and reached a maximum value of 11.07 ± 0.85 pg cell−1 in the middle of the afternoon (Fig. 3E, n = 4). Total proteins per cell dropped from 10.00 ± 0.63 pg cell−1 at dusk to 8.10 ± 0.29 pg cell−1 at 2nd dawn (Fig. 3). However, assuming an even redistribution between daughter cells after division, some proteins had to be synthesized during the night to reach such values at 2nd dawn. The drop in cellular carbon associated with carbohydrate and TAG may be enough to explain the redistribution of carbon skeletons from these cellular pools to protein. We did not measure polar lipids in this study but it is possible that they represent a large portion of the carbon budget [47]. Galactolipids, which are integral components of the thylakoid membranes, are consumed in the dark in other algae [55,56]. Our results suggest that dark catabolism in diatoms may represent a complicated interplay between lipid, carbohydrate and protein breakdown and biosynthesis. 3.3. Changes in photosynthetic parameters in situ and ex situ over a day/ night cycle Our short-term assays of photosynthetic capabilities indicated that oxygen evolution rates reached saturation at approximately 200 μmol photons m−2 s−1 (Table 2). Therefore, light absorption rates were in excess of photosynthetic assimilation capacity for the majority of the day (Fig. 1). These conditions are believed to typically induce photoinhibition [6,31,57]. However, we were surprised to observe minimal indications of photoinhibition in our experiments. We observed a small but significant drop in Fv/Fm from dawn to mid-day and this was followed by a recovery by dusk (Fig. 4A, n = 4, p b 0.05). The maximum rate of oxygen evolution normalized to cell, Pmax , was measured ex situ. It increased from 87 ± 12 nmol O2 cell−1 h−1 at dawn to 141 ± 21 nmol O2 cell−1 h−1 at dusk (Table 2, n = 4, p b 0.05). However, the chlorophyll normalized Pmax, the minimum irradiance at saturation Ek, and α, the light limited slope of photosynthesis vs. irradiance, did not change significantly throughout the day (Table 2). The efficiency of photosynthesis on a per chlorophyll basis remained unchanged throughout the day. Cellular chlorophyll content increased during the day, leading

to the increase in photosynthesis capability on a per cell basis (Fig. 3F, Fig. S3). NPQ is an important photo-protective process that dissipates excess absorbed light energy as heat [58]. The small reduction in PSII maximal quantum yield observed at mid-day (Fig. 4A) correlated with a slight induction of NPQ (0.32 ± 0.11, Fig. 4B, n = 6). We also observed slight decreases of the functional absorption cross-section serving PSII photochemistry (σPSII, Fig. 4C) and a slight increase in the Fo/σPSII parameter, which is proposed to estimate the relative concentration of active PSII centers [59] (Fig. 4D). NPQ reduces the effective transfer of light energy to the reaction center of PSII and it has been previously shown to negatively correlate with σPSII in Phaeodactylum [60,61]. The maximal NPQ values in the ex situ RLCs were higher than those estimated by our in situ methods (Fig. 5). However, we did not observe the large NPQ values (up to 8) that have been measured when Phaeodactylum cells are grown in light/dark cycles of 5 min/50 min [62] or those measured after a single shift from low to high irradiances [14,15]. NPQ appeared to have a small contribution to photoprotection in Phaeodactylum cells in our conditions, but none of our observations suggested significant photoinhibition which suggests that other photoprotective strategies may have been employed. AET pathways can be used to consume some of the excess reducing power generated by photosynthesis, thus decreasing the probability of ROS formation [20,58]. AET rates were not quantified here. However, modeling of Phaeodactylum photophysiology under a sinusoidal light regime predicted that AET could represent an important proportion of the total electron transport in thylakoids [26,63]. The proportion of electrons consumed by AET, measured as light dependent oxygen consumption, increased with increasing growth irradiance in the centric diatom Thalassiosira pseudonana, suggesting AET can be an important component for maintaining redox balance in excess light [21]. PSII cyclic electron flow [63], a plastid terminal oxidase, the transfer of reduced carbon compounds to the mitochondria for oxidation [64], or the Mehler reaction could be the mechanism of AET in diatoms. It appears likely that AET significantly contributed to photoprotection in our experiments given the low NPQ values in situ. Chlorophyll fluorometry derived estimates of plastoquinone oxidation in the dark [36] suggested no change in oxidation capacity throughout the day (Fig. S4). However, we note that this does not estimate the rate of AET in illuminated cells. We simultaneously measured oxygen evolution and chlorophyll fluorescence parameters using a RLC protocol to assess if photosynthetic capability changed throughout the day. Fig. 5 presents data from our observations of photophysiology at mid-morning and mid-afternoon. In both cases, gross oxygen evolution rates varied almost identically as a function of light intensity (Fig. 5A). The proportion of closed PSII reaction centers, represented by the fluorescence-derived parameter 1-qL [41], mirrored the pattern of oxygen evolution with irradiance. This suggests that saturation of photosynthesis coincided with a reduced electron transport chain [5]. Interestingly, the 1-qL and photosynthesis vs. irradiance curves appeared identical between all timepoints (Fig. S5). The only significant differences observed in chla fluorescence patterns was that cells were capable of higher maximal NPQ in the morning compared to the afternoon samples (NPQmax = 2.09 ± 0.12, vs. 1.49 ± 0.05, Fig. 5B). NPQ induction occurred at light intensities that were sufficient to saturate photosynthesis (Fig. 5A & C). These high ex situ NPQ

Table 2 Photosynthetic performance parameters observed over a day. Timepoint

Pmax (nmol O2 μg chla−1 h−1) Ek (μmol photons m−2 s−1) α

Dawn Mid-morning Mid-day Mid-afternoon Dusk

470 ± 20ab 520 ± 70ab 560 ± 50a 500 ± 30ab 440 ± 50b

183 ± 21a 228 ± 47a 246 ± 12a 218 ± 20a 177 ± 61a

3.1 × 10−3 ± 1 × 10–3a 2.3 × 10−3 ± 5 × 10–4a 2.3 × 10−3 ± 3 × 10–4a 2.3 × 10−3 ± 2 × 10–4a 2.6 × 10−3 ± 3 × 10−4a

Pmax (nmol O2 cell−1 h−1) Dark respiration rate (nmol O2 cell−1 h−1) 87 ± 12a 104 ± 14ab 126 ± 18bc 135 ± 19c 130 ± 28bc

16.0 ± 0.8a 18.3 ± 1.3ab 18.5 ± 1.2ab 19.5 ± 1.3b 18.5 ± 1.9ab

Maximum photosynthesis rate (Pmax), irradiance at saturation (Ek) and light-limited slope (α; nmol O2 μg chla−1 h−1 μmolphotons−1 m2 s1). The dark respiration rates are reported normalized to cell (n = 4, ± standard deviation, different letters indicate statistically different averages between timepoints).


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Fig. 5. Changes in photosynthetic and photoprotective capabilities ex situ. Fluorescence parameters were recorded with a Pulse Amplitude Modulated (PAM) fluorometer and oxygen evolution/consumption was simultaneously measured with a FireSting oxygen meter. Increasing actinic light intensities were applied to obtain Rapid Light Curves (RLCs). As an example, only RLCs associated with mid-morning (closed squares) and mid-afternoon (open triangles) samples are shown here (RLCs for other timepoints are shown in Fig. S5). A) Gross oxygen evolution rates (μmol O2 μg chla−1 h−1) B) NonPhotochemical Quenching (NPQ) C) 1-qL, a parameter that estimates plastoquinone pool reduction state [41]. Data are the averages of n = 4 independent cultures. Error bars show 1 standard deviation. When not shown, error bars are smaller than symbols.

Fig. 4. Changes in chlorophyll a fluorescence parameters in situ over a day. A) Maximum quantum yield of photosystem II (Fv/Fm) B) Non-photochemical quenching (NPQ) C) Effective absorption cross-section serving PSII photochemistry (σPSII, Å2 quantum−1) and D) The ratio of initial fluorescence during a saturating flash (F0) to σPSII. Data are the averages of n = 4 biological replicates, error bars show 1 standard deviation. When not shown, error bars are smaller than symbols. Different letters indicate statistically different groups.

values were observed after several minutes in excess light and the rapidly changing light environment encountered in situ may not be adequate to induce significant NPQ, as mentioned above. High light fluxes induce photodamage to the D1 protein of photosystem II [9,31]. Phaeodactylum cells avoid net photodamage by mobilizing the photoprotective mechanisms outlined above, by upregulating enzymes associated with oxidative stress defense [19], or by increasing repair rates of damaged PSII [9,38,65]. We did not measure ROS scavenging activities in our sinusoidal light/dark cycles but hypothesize

that the antioxidant system played an important role in preventing net photodamage in excess light [2]. D1 damage correlates with light intensity [66] and D1 protein synthesis has been shown to increase as a function of light intensity in plants and algae [67]. A previous study on Phaeodactylum found that the steady state levels of D1 decreased after a shift from low to excess light but inferred major increases in synthesis rates with the use of protein synthesis inhibitors [31]. We did not observe any significant variation in the steady state levels of D1 protein throughout 1 day/night cycle (Fig. 6A, Fig. 6C). We used the β subunit of ATP synthase (ATP-B) as a loading control which also did not change throughout the cycle. We note that our methodology does not capture the dynamics of D1 removal or synthesis. However, our results coupled with those outlined above suggest that the PSII repair cycle efficiently counterbalances photoinhibition in dense cultures.

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ponds or tubular photobioreactors [68–70]. In open ponds, almost no decrease of Fv/Fm was observed around solar noon and NPQ induction levels were small, reminiscent of our results [68,69]. Mid-day photoinhibition was observed at low cell densities in tubular photobioreactors, but increasing Phaeodactylum cell densities ameliorated this [68–70]. The low degree of photoinhibition found in outdoor cultures is likely due to the rapid mixing of cultures between excess light and sub-saturating intensities and the net time spent in excess light is not sufficient to induce NPQ or induce significant damage to PSII. This supports earlier suggestions that cell density and mixing regime are important considerations for maximizing productivity [28,71]. 4. Conclusions We used bench-top photobioreactors to characterize Phaeodactylum biomass partitioning and photophysiology under a sinusoidal day/night cycle. Our conditions mimicked those encountered in large-scale photobioreactors or open ponds [25] with relatively low cell densities and rapid mixing. We can infer two main conclusions about Phaeodactylum growth in mass culture. 1) Carbon partitioning between biomass components changes significantly over a day/night cycle. For example, TAGs and carbohydrates were more abundant at the end of the day compared to the morning. An opposite trend was observed for proteins. These daily patterns should be important considerations for timing of harvest, for the extraction of a biochemical compound of interest, for designing metabolic shunts or for measuring daily productivity. 2) Phaeodactylum evaded significant photoinhibition with minimal NPQ. This suggests that engineering NPQ dynamics in algae may not increase yields, despite its promise in other systems [72]. Our work represents an early step in understanding the physiology of photosynthetic microbes in mass culture. Extending this approach to include other environmental stresses experienced in mass culture will improve our ability to engineer biological and mechanical systems to improve biomass or bioproduct yields. Acknowledgements

Fig. 6. Changes in the abundance of the D1 protein over a day/night cycle. Gel electrophoresis/Western blots were performed loading 0.75 μg total proteins per well unless otherwise specified. A) Immunoblot detection of ATP-B and PsbA1 (D1 protein of PSII) with the final 3 lanes showing a dilution of samples from 2nd dawn. B) Results from comparative densitometry showing the relative abundance of ATP-B compared to 2nd Dawn C) Results from comparative densitometry showing the abundance of the D1 protein relative to 2nd Dawn, after normalization to ATP-B content. Data are the averages of n = 3 biological replicates. Error bars show one standard deviation. Different letters indicate statistically different groups. Black/white bars above the graphs indicate night/day respectively.

We would like to thank David Xing, Michael Cantrell, Bjoern Andersson and Alexander Hughes for their critical reading of the manuscript. Comments from one anonymous reviewer and Douglas Campbell greatly improved this manuscript. This project was funded by the US Department of Energy Office of Science, grant DOE-BER-DESC0008595. DJ was supported by Fondation Bettencourt Schueller. MAC was supported by NSF-GRFP. Appendix A. Supplementary data Supplementary data to this article can be found online at http://dx. doi.org/10.1016/j.algal.2016.05.014. References

3.4. Comparison with previous studies under natural light regimes Several previous studies have investigated Phaeodactylum photophysiology under a sinusoidal light regime, using either benchtop or outdoor cultures. Past bench-top studies employed 3 cm-deep rectangular photobioreactors for culture maintenance, to reduce selfshading effects. These studies observed higher levels of NPQ capability and modeling of photophysiology suggested a high degree of AET in excess light [26,63]. This scenario represents what could occur in shallow, non-turbid natural waters. When fluctuations of light intensity with a period of 30 min were applied to mimic vertical mixing in natural systems, much smaller NPQ values were observed close to those reported in our work [26]. The outdoor studies were conducted either in open

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