Applied Microbiology and Biotechnology

Re-cultivation of Neochloris oleoabundans in exhausted autotrophic and mixotrophic media: the potential role of polyamines and free fatty acids --Manuscript Draft--

Manuscript Number: AMAB-D-15-00751R1 Full Title: Re-cultivation of Neochloris oleoabundans in exhausted autotrophic and mixotrophic media: the potential role of polyamines and free fatty acids Article Type: Original Article Section/Category: Applied microbial and cell physiology Corresponding Author: Simonetta Pancaldi

Italy Corresponding Author Secondary Information: Corresponding Author's Institution: Corresponding Author's Secondary Institution: First Author: Alessandra Sabia First Author Secondary Information: Order of Authors: Alessandra Sabia Costanza Baldisserotto Stefania Biondi Roberta Marchesini Paola Tedeschi Annalisa Maietti Martina Giovanardi Lorenzo Ferroni Simonetta Pancaldi Order of Authors Secondary Information: Funding Information: Abstract: Neochloris oleoabundans (Chlorophyta) is widely considered one of the most promising microalgae for biotechnological applications. However, the large-scale production of microalgae requires large amounts of water. In this perspective, the possibility of using exhausted growth media for the re-cultivation of N. oleoabundans was investigated in order to simultaneously make the cultivation more economically feasible and environmentally sustainable. Experiments were performed by testing the following media: autotrophic exhausted medium (E+) and mixotrophic exhausted medium after cultivation with glucose (EG+) of N. oleoabundans cells grown in a 20-L photobioreactor (PBR). Both exhausted media were replenished with the same amounts of nitrate and phosphate as the control brackish medium (C). Growth kinetics, nitrate and phosphate consumption, photosynthetic pigments content, photosynthetic efficiency, cell morphology, and lipid production were evaluated. Moreover, the free fatty acid (FFA) composition of exhausted media and the polyamine (PA) concentrations of both algae and media were analyzed in order to test if some molecules, released into the medium, could influence algal growth and metabolism. Results showed that N. oleoabundans can efficiently grow in both exhausted media, if appropriately replenished with the main nutrients (E+ and EG+), especially in E+ and to the same extent as in C medium. Growth promotion of N. oleoabundans was attributed

Powered by Editorial Manager® and ProduXion Manager® from Aries Systems Corporation to PAs and alteration of the photosynthetic apparatus to FFAs. Taken together, results show that recycling growth medium is a suitable solution to obtain good N. oleoabundans biomass concentrations, while providing a more sustainable ecological impact on water resources. Response to Reviewers: see attached file

Dear Editor,

we are very grateful that our MS was positively evaluated by the Reviewers.

We have modified the MS according with indications of Reviewers 1 and 2. Changes made in the MS were highlighted by using the track changes mode in MS Word.

Hoping that our revised MS can be now eligible for publication in Applied Microbiology and Biotechnology, I convey you my very best regards.

Simonetta Pancaldi

Associate Professor Cover Letter

UNIVERSITÀ DEGLI STUDI DI FERRARA DIPARTIMENTO di SCIENZE della VITA e BIOTECNOLOGIE Corso Ercole I d'Este, 32 - 44121 Ferrara (Italia)

Ferrara, March 26th, 2015

Dear Prof. Steinbüchel,

please find our manuscript entitled “Re-cultivation of Neochloris oleoabundans in exhausted

autotrophic and mixotrophic media: the potential role of polyamines and free fatty acids” by

Alessandra Sabia, Costanza Baldisserotto, Stefania Biondi, Roberta Marchesini, Paola Tedeschi,

Annalisa Maietti, Martina Giovanardi, Lorenzo Ferroni, Simonetta Pancaldi, which we are submitting

for publication in Applied Microbiology and Biotechnology.

 We declare that the paper has not been published previously, is not accepted for publication, has not been submitted previously to Applied Microbiology and Biotechnology and is not currently under consideration for publication elsewhere.

 All Authors approve the submission of the paper to Applied Microbiology and Biotechnology and no-one entitled to authorship has been excluded.

 The work does not violate the laws of the country (Italy) in which it was carried out, including those relating to conservation and animal welfare.

 The Authors declare that they have no conflict of interest.

 Colour figures are intended only for the online version.

Hoping that our work is eligible for publication in your Journal, I convey you our very best regards.

Simonetta Pancaldi

University of Ferrara Laboratory of Plant Cytophysiology Department of Life Sciences and Biotechnologies Corso Ercole I D’Este, 32 44121 Ferrara – Italy e-mail: [email protected]; Phone: 0039 0532 293786; Fax: 0039 0532 208561 Authors' Response to Reviewers' Comments Click here to download Authors' Response to Reviewers' Comments: Response to Reviewers comments.docx

List of changes and answers to Reviewers’ comments

Reviewer #1:

General question:

About the correlation between the amount and composition of intracellular fatty acid composition inside Neochloris oleoabundans cells and in exhausted media, we performed several in-depth analyses on lipid profile of autotrophic and mixotrophic cells. The results are very interesting and are the topic of an extensive study we have just submitted to an International journal for publication. We cannot reiterate these data in the revised MS, however we can confirm to the Reviewer that the FFA profile in the medium is consistent with that in cells.

Other comments:

1) Glucose concentration of EG media was determined by 1,5-dinitrosalicylic acid assay during the growth of Neochloris oleoabundans inside the PBR. At the end of the experiment, i.e. at the 8th day of growth, when the exhausted medium was harvested for our experiments, glucose was shown totally consumed. This aspect has been better specified in “Material and Methods” section, at page 6, line 19ff. About pH changes during the experiments, measurements were periodically performed in C, E+ and EG+ samples, but we did not introduce them in the previous MS. Results showed that pH of exhausted media remained almost stable, fluctuating around 9.5 and 8 during the whole experiment, without any evident trend. For this reason we think that growth promotion and metabolic changes observed in microalgae grown in E+ and EG+ media cannot be attributed to pH fluctuations. In the Revised MS we included this experiment (page 6 line 21ff, page 7, line 23, page 13 line 22 ff, page 18 line 1 ff, Figure S1 in “Supplemental Materials”).

2) OK. “Data treatment” has been changed to “Data analysis”.

3) In BM medium, the concentration of phosphate is 100 times lower than nitrate (0.2 gL-1 vs 0.002 gL-1 respectively). It is probably for this reason that in this work phosphate was consumed faster than nitrate. This aspect, as requested, has been explained in the “Discussion” section (page 18 line 8ff). However, it is also reported in previous works that FFAs induce an alteration in membrane permeability of microalgal cells, with the consequent rapid consumption of phosphate. This aspect was already discussed in the MS (page 20 line 2ff).

4) We thank the Reviewer for his/her positive comments on TEM images.

5) OK, as requested, the two section “extracellular FFA composition” and “polyamine content” have been moved before the “growth kinetics” section.

6) Figures 2 and 4 have been changed to higher resolutions (1200 dpi).

Reviewer #2:

1) About the number of repetition, we performed three replicates for each treatment (C, E+ and EG+ media, page 7, line 8). Moreover, both exhausted media derived from replicates of different cycles in PBR, stocked at -20°C before the experiments.

2) EG+ medium was not replenished with glucose, but only with nitrate and phosphate at the initial concentration of BM medium. For this reason, in this work we did not include a control in which fresh BM medium was supplied with glucose, but only a control composed by fresh BM medium, that was used as reference for both E+ and EG+ samples.

3) Regarding the micronutrient concentrations, their consumption and the linkage with the photosynthetic efficiency, according with the Reviewer we also think that a lack of these trace elements may contribute + + to the decrease of FV/FM in both E and EG samples. However, because of their low concentration even at time 0 in fresh BM medium, we were not able to measure the consumption of these minerals by microalgae during the experiments. The quantification of trace elements has been reported also in previous works to be very difficult because of the low concentrations in the microalgal media (Hadj- Romdhane et al., 2012. Bioresource Technol 123:366-374). Moreover, considering that we did not replenish the exhausted media with trace elements, it is probable that micronutrients were lacking in E+ and EG+ since the beginning of the experiments. For this reason, we believe that the gradual decrease in photosynthetic efficiency in cells grown in both exhausted media can be more probably attributed to the presence of inhibiting molecules rather than to the consumption of trace elements.

4) We thank the Reviewer for his/her suggestion to implement research with samples grown in fresh BM medium added with FFAs and PAs. We also agree that it would be very interesting to perform these analyses to provide a mechanistic physiological background to the potential role of these metabolites on algae metabolism. However, the main scope of this MS was to test recycle of media in an applied perspective, according to this Journal’s scope. As correctly pointed out by the Reviewer, the research will be extended in the future with basic analyses to improve the understanding of algae physiology. Clearly, the experimental efforts to reach this goal go well beyond the aim of the current MS and will be the subject of a new paper.

Minor remarks: Sentence in the Abstract has been changed as requested, specifying that exhausted media which allowed to obtain good biomass concentration of N. oleoabundans were replenished with the main nutrients (page 2, line 7ff). Authors' Response to Reviewers' Comments Click here to download Authors' Response to Reviewers' Comments: Sabia et al. Revised MS marked.doc

Re-cultivation of Neochloris oleoabundans in exhausted autotrophic and mixotrophic media:

the potential role of polyamines and free fatty acids

Alessandra Sabia1, Costanza Baldisserotto1, Stefania Biondi2, Roberta Marchesini1, Paola

Tedeschi3, Annalisa Maietti3, Martina Giovanardi1, Lorenzo Ferroni1, Simonetta Pancaldi1

1Department of Life Sciences and Biotechnology, University of Ferrara, Corso Ercole I d'Este 32,

44121 Ferrara, Italy.

2Department of Biological, Geological and Environmental Sciences, University of Bologna, via

Irnerio 42, 40126 Bologna, Italy.

3Department of Chemical and Pharmaceutical Science, University of Ferrara, via Fossato di

Mortara 17, 44121 Ferrara, Italy.

*Corresponding Author. Tel: +39 0532 293786; Fax: +39 0532 208561; E-mail address:

[email protected].

ABSTRACT

Neochloris oleoabundans (Chlorophyta) is widely considered one of the most promising microalgae Formatted: Font: Italic

for biotechnological applications. However, the large-scale production of microalgae requires large

amounts of water. In this perspective, the possibility of using exhausted growth media for the re-

cultivation of N. oleoabundans was investigated in order to simultaneously make the cultivation

more economically feasible and environmentally sustainable. Experiments were performed by

testing the following media: autotrophic exhausted medium (E+) and mixotrophic exhausted

medium after cultivation with glucose (EG+) of N. oleoabundans cells grown in a 20-L

1 photobioreactor (PBR). Both exhausted media were replenished with the same amounts of nitrate and phosphate as the control brackish medium (C). Growth kinetics, nitrate and phosphate consumption, photosynthetic pigments content, photosynthetic efficiency, cell morphology, and lipid production were evaluated. Moreover, the free fatty acid (FFA) composition of exhausted media and the polyamine (PA) concentrations of both algae and media were analyzed in order to test if some molecules, released into the medium, could influence algal growth and metabolism.

Results showed that N. oleoabundans can efficiently grow in both exhausted media, if appropriately replenished with the main nutrients (E+ and EG+), especially in E+, and to the same extent as in C medium. Growth promotion of N. oleoabundans was attributed to PAs and alteration of the photosynthetic apparatus to FFAs. Taken together, results show that recycling growth medium is a suitable solution to obtain good N. oleoabundans biomass concentrations, while providing a more sustainable ecological impact on water resources.

Keywords:

Neochloris oleoabundans

Recycling culture media

Photosynthetic apparatus

Biomass production

Polyamines

Free fatty acids

Introduction

Over the last decades, microalgae have received much attention as promising high-potential

feedstock for biodiesel (Chisti 2007; Li et al. 2008; Borowitzka and Moheimani 2013) and as an

attractive raw material for the production of a wide range of high value bioproducts (Molina Grima

et al. 2003; Mata et al. 2010). They are photosynthetic microorganisms with numerous key

features, which potentially make them a more appreciable renewable source than terrestrial plants

(Richmond 2004; Hu et al 2008; Ndimba et al. 2013). For instance, these microorganisms have

faster growth rates, higher photosynthetic efficiencies, higher rates of carbon dioxide fixation, and

higher biomass productivities compared to plants (Richmond 2004; Gouveia et al. 2009; Harun et

al. 2010; Mata et al. 2010). Moreover, microalgae can thrive in non-arable lands being able to grow

in waters having different salinity levels and chemical composition (Smith et al. 2010; Borowitzka

and Moheimani 2013).

The environmental and economical impact of large-scale production of microalgae has been widely discussed, with the aim of making the process increasingly sustainable (Sheehan 2009; Solomon

2010). In this perspective, microalgal cultivation systems are crucial factors to be taken into account

(Smith et al. 2010; Stephens et al. 2010; Borowitzka and Moheimani 2013). A wide variety of systems has been described by Mata and co-workers (2010) and recently by Borowitzka and

Moheimani (2013). Microalgae can be cultivated in open pond reactors or closed photobioreactors

(PBRs), depending on algal strain, issue of research, type of desired products and environmental conditions (Chisti 2007; Mata et al. 2010; Borowitzka and Moheimani 2013). Although open ponds are the most commonly used culture systems, due to their low cost of production and operation, recently closed PBRs are receiving much attention for the possibility of producing valuable compounds (Pulz 2001; Grobbelaar 2009; Smith et al. 2010). However, algal cultures require large amounts of water in both culture systems (Lam and Lee 2012; Borowitzka and Moheimani 2013).

3 Considering that water is becoming a scarce natural resource, water demand represents an important factor when performing large-scale cultivation (Batan et al. 2013; Farooq et al. 2014). In order to reduce the high production costs and, at the same time, to make microalgal production more environmentally sustainable, recycling culture medium has been proposed as a possible solution

(Yang et al. 2011; Hadj-Romdhane et al. 2012; Farooq et al. 2014), especially in large-scale culture systems (Borowitzka 2005; Lam and Lee 2012; Zhu et al. 2013). However, the feasibility of recycling growth medium to re-cultivate microalgae has been tested only in a few cases (Lívanský et al. 1996; Rodolfi et al. 2003; Hadj-Romdhane et al. 2013; Zhu et al. 2013). It is reported that the recycling medium could have negative effects on biomass productivity, due to the release of inhibitory secondary metabolites (Richmond 2004). Harmful metabolites are, in fact, released during microalgal growth under physiological stress (Ikawa 2004; Moheimani and Borowitzka

2006). Free fatty acids (FFAs), and substances derived from the photooxidation of unsaturated fatty acids, are the most common metabolites with inhibitory effects on microalgae (Ikawa 2004; Wu et al. 2006; Bosma et al. 2008; Stephen et al. 2010).

In the present work, the suitability of recycling growth media for the re-cultivation of Neochloris oleoabundans (Chlorophyta) was investigated. This species was chosen because it is considered one Formatted: Font: Italic of the most promising oil-rich microalgae, due to its capability to accumulate lipids when grown under nitrogen starvation (Tornabene et al. 1983; Chisti 2007; Li et al. 2008; Pruvost et al. 2009) or mixotrophically, in the presence of glucose, or carbon-rich wastes, as organic carbon sources

(Giovanardi et al. 2013; Baldisserotto et al. 2014). Present results clearly indicate that N. oleoabundans can efficiently grow in its exhausted growth medium, if suitably replenished with the main nutrients. Based on this encouraging result, and with the aim of gaining further knowledge on the morpho-physiological aspects and biotechnological applications of this algal strain, the following features were analyzed: (1) growth kinetics in parallel to nitrate and phosphate consumption; (2) photosynthetic pigment content; (3) PSII maximum quantum yield; (4) cell morphology, with special attention to intracellular lipid accumulation, (5) FFA accumulation in

4 recycled growth media, and (6) polyamine (PA) concentration both inside cells and in the corresponding culture media. PAs were determined in order to understand if these plant growth regulators could be responsible for the growth promotion of N. oleoabundans in recycled growth medium. It is known, in fact, that PAs, together with other plant growth regulators, have stimulatory effects on algal growth and metabolism, and are involved in mitigating various types of biotic and abiotic stress (Tate et al. 2013).

Materials and Methods

Algal strains and culture condition

The strain used in this study was the Chlorophyta N. oleoabundans UTEX 1185 (syn. Ettlia Formatted: Font: Italic oleoabundans) (Sphaeropleales, Neochloridaceae), obtained from the Culture Collection of Algae Formatted: Font: Italic of the University of Texas (UTEX, Austin, Texas, USA; www.utex.org).

N. oleoabundans was cultivated in axenic liquid brackish medium (BM) (Baldisserotto et al. 2012) in a coaxial 20-L capacity PBR (M2M Engineering, Grazzanise, Caserta, Italy). Algae were cultivated autotrophically in BM for 25 days or mixotrophically, by addition of 2.5 g L-1 glucose, for 8 days, according to previously described protocols (Baldisserotto et al. 2014; Giovanardi et al.

2014). For autotrophic cultivation, cells were inoculated into the PBR to obtain an initial cell density of about 0.6 x 106 cells mL-1, while for mixotrophic cultivation initial cell density was higher (10 x 106 cells mL-1). Culture conditions in the PBR were: 24 ± 1°C; sterile air injection at

-2 -1 the bottom of the PBR, with 0.5/3.5 h bubbling/static cycles; irradiance 65 µmolphotons m s of

PAR (16:8 h light:dark photoperiod). Light was supplied with inner cool–white fluorescent Philips tubes.

Algal growth and morphology in the PBR were monitored as described below.

Preparation of growth media

Experiments were performed by testing the following media:

- C: freshly prepared BM medium (control);

- E: autotrophic exhausted medium;

- EG: mixotrophic exhausted medium after cultivation with glucose;

- E+: autotrophic exhausted medium replenished with nitrate and phosphate concentrations as for

BM medium;

- EG+: mixotrophic exhausted medium after cultivation with glucose and replenished with nutrients as in E+.

In order to obtain the exhausted media for the recycling experiment, N. oleoabundans was grown inside the PBR under the culture conditions described in the previous section. For E+ medium preparation, about 500 mL of autotrophic algal culture in the stationary phase of growth (at the 15th day of cultivation in the PBR) were centrifuged at 2,000 g for 10 min in order to separate the medium from the algae, thus obtaining an autotrophic exhausted medium (E). The medium appeared pale-yellow and was free from algae, bacteria and protozoa; its optical density at a

th wavelength of 750 nm (OD750) was 0.02. For EG+ medium, at the 8 day of cultivation in the PBR, the same aliquots of algal culture were harvested by centrifugation (2,000 g for 10 min). In this case, the mixotrophic exhausted medium (EG) was straw-yellow in colour and presented a weak bacterial contamination due to the glucose addition (OD750 =0.009). Glucose was proved to be totally consumed during the previous cultivation inside the PBR by 3,5-dinitrosalicylic (DNS;

Sigma-Aldrich, Gallarate, Milan, Italy) acid assay, according to Giovanardi et al. (2014). In both exhausted media, pH was not adjusted. After determining the nitrate and phosphate concentrations of the two exhausted media (see “Nitrate and phosphate analyses”), KNO3 and K2HPO4 were axenically added to reach final concentrations of 0.2 and 0.02 g L-1, respectively, i.e., the typical concentration of those components in BM, thus obtaining the replenished exhausted media (E+ and

6 EG+) used for experiments.

Experimental design

When autotrophic cultures of N. oleoabundans, grown in BM in the PBR, reached a cell density of

10 x 106 cells mL-1 (after about 9 days of cultivation), aliquots of cells were inoculated into 300-mL

Erlenmeyer flasks (150 mL total volume) containing C, E+, EG+ media, to obtain an initial cell density of about 0.5 x 106 cells mL-1. The cultures were placed in a growth chamber (24 ± 1 °C, 80

-2 -1 μmolphotons m s of PAR with a 16:8 h light-darkness photoperiod), and cultivated with continuous shaking at 80 rpm, without external CO2 supply. Experiments were performed in triplicate. Aliquots of cultures (cells and media) were collected at different times of cultivation up to 25 days, depending on the analysis.

Analyses

Growth evaluation

Aliquots of cell samples cultivated in control and exhausted media were counted at 0, 3, 7, 12, 17 and 25 days of cultivation using a Thoma haemocytometer (HBG, Giessen, Germany) under a light microscope (Zeiss, model Axiophot) and growth curves were obtained.

The growth rate (μ, number of divisions per day) during the exponential phase was calculated with the following equation:

-1 μ (div d ) = (log2N1 − log2N0) / (t1− t0), where μ is the growth rate, N1 the cell number at time t1, N0 the cell number at time 0 and t1−t0 the time interval (days) (Giovanardi et al. 2013).

In parallel to growth, pH was also periodically monitored on small aliquots of samples.

Nitrate and phosphate analyses

After 12 and 25 days of cultivation, samples of C, E+ and EG+ media were harvested by centrifugation to analyze nitrate and phosphate concentrations. These nutrients were quantified colorimetrically using a flow-injection autoanalyzer (FlowSys, Systea, Roma, Italy).

Photosynthetic pigment extraction and quantification

For photosynthetic pigment analysis, cell samples were collected by centrifugation after 0, 7, 17, 21 and 25 days of cultivation. Extraction of photosynthetic pigments was performed according to

Baldisserotto et al. (2014). The extracts were measured with a Pharmacia Ultrospec 2000 UV-Vis spectrophotometer (1-nm bandwidth; Amersham Biosciences, Piscataway, NJ, USA) at 666

(chlorophyll a - Chla), 653 (chlorophyll b - Chlb) and 470 nm (carotenoids - Car). Quantification was performed according to equations reported in Wellburn (1994).

PAM fluorimetry

The PSII maximum quantum yield of algae was determined at the same cultivation times considered for growth kinetics measurements. A pulse amplitude modulated fluorometer (ADC

Bioscientific Ltd, Hoddesdon, Hertfordshire, UK) was used to determine the in vivo chlorophyll fluorescence of PSII. The PSII maximum quantum yield is reported as FV/FM ratio, i.e. (FM – F0)/

FM, where variable fluorescence is FV = (FM−F0), FM is the maximum fluorescence and F0 is the initial fluorescence of samples (Lichtenthaler et al. 2005). This measurement is considered a valid method to probe the maximum quantum yield of photochemistry in PSII (Kalaji et al. 2014).

Moreover, it is useful to estimate the physiological state of plants and microorganisms also under nutrient stress (White et al. 2011). Samples were prepared as reported in Ferroni et al. (2011) after

15 min of dark incubation.

Light and fluorescence microscopy

For microscopic observations, cell samples were routinely collected throughout the cultivation period inside the PBR and during the experiment with exhausted media. Aliquots of samples were observed using a microscope (Zeiss, model Axiophot) with conventional and fluorescent attachments. The light source for chlorophyll fluorescence observation was a HBO 100 W pressure mercury vapour lamp (filter set, BP436/10, LP470). Pictures of cells were taken with a Canon

IXUS 110 IS digital camera (12.1 megapixels), mounted on the ocular lens through a Leica DC150 system (Leica Camera AG, Solms, Germany).

Lipid staining

During experiments, the intracellular presence of lipids was evaluated by staining cells with the fluorochrome Nile Red (9-diethylamina-5Hbenzo[α]phenoxazine-5-one, 0.5 mg dissolved in 100 mL acetone; Sigma-Aldrich, Gallarate, Milan, Italy), as described in Giovanardi et al. (2014). After incubation at 37°C in darkness for 15 min, cells were observed with the microscope described above at an excitation wavelength of 485 nm (filter set, BP485, LP520). Photographs were taken with the camera described above.

Transmission electron microscopy (TEM)

TEM observations were made on cells harvested by centrifugation (600 g, 10 min) at the 12th day of cultivation. Cells were fixed, post-fixed and dehydrated as reported in Baldisserotto et al. (2012).

Embedding in resin and staining procedures were performed as previously described (Pancaldi et al.

2002). Sections were observed with a Hitachi H800 electron microscope (Hitachi, Tokyo, Japan) at the Electron Microscopy Centre, University of Ferrara.

Extracellular free fatty acids analysis

To analyze the FFA composition of the exhausted E and EG media used to prepare E+ and EG+ replenished media, aliquots of N. oleoabundans cultures grown autotrophically and mixotrophically in the PBR were collected at the 15th and at the 8th day of cultivation, respectively. Samples were centrifuged at 2,000 g for 10 min in order to separate the medium from algae. Fifty mL of media obtained from the autotrophic (E) and mixotrophic (EG) cultures were freeze-dried for analysis.

The extracellular FFA composition was determined in duplicate by gas chromatography-mass spectrometry (GC-MS). For extraction, samples were dissolved in 3 mL of hexane, sonicated for 15 min and extracted overnight. Fatty acid methyl esters were prepared by transesterification with 1.5 mL of 5% of sodium hydroxide in methanol solution. Sample volumes of 1 μl were injected into the

GC-MS apparatus, which consisted of a Varian Saturn 2100 MS/MS ion trap mass spectrometer.

Separations were performed using a Zebron ZB-WAX Phenomenex capillary column (60 cm in length, 0.25 mm i.d) supplied with helium carrier gas at 1 mL min-1 constant flow. The injector temperature was 250 °C and the oven temperature programme was the following: start 100°C for 2 min, ramp to 200°C at 10 °C/min, and hold for 108 min. The MS acquisitions were performed by full scan mode.

Determination of polyamines

To analyze the free PA composition of N. oleoabundans cells cultivated autotrophically and mixotrophically in PBR and in the corresponding E and EG media, samples were collected as described for FFA analyses. Freeze-dried E or EG media and algae (50 and 20 mg, respectively) were extracted in cold 4% perchloric acid, kept for 1 h on ice, and then centrifuged at 15,000 g for

15 min. Aliquots (200 L) of the supernatants and standard solutions of putrescine (Put), spermidine (Spd) and spermine (Spm) were derivatised with dansyl chloride (Scaramagli et al.

10 1999). Dansylated derivatives were extracted with toluene, taken to dryness and resuspended in acetonitrile. PAs were separated and quantified by HPLC using a reverse phase C18 column

(Spherisorb ODS2, 5-μm particle diameter, 4.6 x 250 mm, Waters, Wexford, Ireland) and a programmed acetonitrile:water step gradient (flow rate 1 mL min-1) on a Jasco system (Jasco Corp.,

Tokyo, Japan) consisting of a PU-1580 pump, an LG-1580-02 ternary gradient unit, a DG-1580-53 three-line degasser and a FP-1529 fluorescence detector, linked to an autosampler (AS 2055 Plus).

Data treatment analysis

Data were processed with Graphpad Prism 6 (Graph Pad Software, San Diego CA, USA). In each case, means ± standard deviations for n number of samples are given. Statistical analyses for comparison of means were carried out using ANOVA, followed by Student’s t-test (significance level, 0.05).

Results

Growth and morphology of N. oleoabundans cultivated autotrophically and mixotrophically in a

20-L PBR

Growth kinetics and cell morphology of N. oleoabundans cultivated autotrophically and mixotrophically in a 20-L PBR are shown in Fig 1. Autotrophic cells rapidly entered the exponential phase starting on the 2nd-3rd day up to the 7th-9th day of cultivation (Fig 1a). After this period, cultures had a short late exponential phase and then entered the stationary phase of growth, reaching a final cell density of about 22 × 106 cells mL-1 at the end of the of cultivation time (25 days). During the entire period, cells showed the typical morphology of N. oleoabundans grown in

11 BM medium, i.e., almost spherical, with a cell diameter of 3-5 µm (Fig 1b, d). One cup-shaped chloroplast, containing a large pyrenoid, was present and emitted an intense red fluorescence due to the presence of chlorophyll (Fig 1b,c). On the 25th day of cultivation, cells maintained their normal features and dimensions, but some lipid globules accumulated inside the cytoplasm, as revealed by

Nile Red staining (Fig 1d,e).

During the first four days of mixotrophic growth in the PBR in the presence of glucose, cells showed an evident increase in cell density, reaching values of ca. 33 x 106 cells mL-1 (Fig 1f).

Subsequently, cells entered the stationary phase of growth, and then cell density leveled off to values of about 36 x 106 cells mL-1 up to the end of the cultivation period, i.e., eight days. At the beginning of cultivation, both cell shape and size, and chloroplast features in mixotrophic cultures

(Fig 1g, h) were similar to those of autotrophic cultures (Fig 1b, c). Starting on the 3rd day and up to the end of cultivation, however, mixotrophic cells showed peculiar features. In fact, the chloroplast lost its characteristic cup shape and translucent globules, which tended to occupy almost the entire cell volume, accumulated inside the cytoplasm (Fig 1i). Nile Red staining confirmed the lipidic nature of these droplets (Fig 1j).

Extracellular FFA composition in E and EG media

The extracellular FFA composition in the exhausted autotrophic (E) and mixotrophic (EG) media of

N. oleoabundans is shown in Table 1. While EG medium comprised both saturated and mono- unsaturated FFAs, E medium contained only saturated FFAs. In fact, both samples contained similar percentages of saturated myristic acid (C14:0) and palmitic acid (C16:0), while the percentage of stearic acid (C18:0) was different between the two samples. In E medium, stearic acid was present at about 64%, while in EG medium at about 56%; however, C18:0 represented the main saturated FFA for both samples. It is noteworthy that only EG medium contained about 6% of the monoenoic oleic acid (C18:1ω9).

12 Polyamine content in E and EG media and in the corresponding N. oleoabundans cells cultivated in a 20-L PBR

As shown in Fig 9a, the PA composition of N. oleoabundans cells cultivated mixotrophically in the

PBR were characterized by higher putrescine, spermine, and, especially, spermidine levels compared to cells cultivated autotrophically. In fact, Spm, Put and Spd concentrations in mixotrophic cells were ca. 2, 30 and 57 times higher, respectively, than those found in autotrophic cells (p<0.001). Conversely, the PA composition in E and EG medium was not significantly different. In fact, both media contained similar amounts of Put and Spm. Interestingly, spermidine was present in high concentration in both media (119 ± 16.91 and 101 ± 5.00 pmol mL-1 in E and

EG, respectively; Fig 9b).

Growth kinetics of N. oleoabundans in exhausted and control growth media

As shown in Fig 2a, the growth of N. oleoabundans in E+ medium was promoted for the entire experiment relative to controls grown on BM medium; on the contrary, growth in EG+ medium was similar to that of the C. All samples entered the exponential phase very soon (during the first 3 days of cultivation), despite some differences in growth rates (Fig 2b). The highest μ values were observed for algae cultivated in the two exhausted media (E+ and EG+, 1.29 and 1.18 div d-1, respectively; control, 0.98 div d-1; p <0.01 in both cases relative to C) (Fig 2b). Subsequently, from the 3rd day onwards, no relevant differences were observed between C and EG+ samples, as both of them showed similar growth kinetics and cell densities (Fig 2a). Conversely, E+ samples reached and maintained the highest cell density; the major difference as compared to the other samples was observed starting on the 7th day, with a cell density of about 16 × 106 cells mL-1 (45 and 50% more than EG+ and C samples, respectively; p <0.01 in both cases relative to E+ samples) at the same time point. After the 17th day, all samples entered the stationary phase, reaching, at the end of the

13 experiment, a final cell density of 25-30 × 106 cells mL-1. During the experiments, aliquots of samples were periodically harvested to measure the pH of the culture media. As observed in Figure

S1, initial pH was very different between control medium (about 7.0) and exhausted media (about

+ + 9.5 in both cases). pH gradually increased up to 9.2 in the control medium, whilst in E and EG media the pH varied between 9.5 and 8.0 without any obvious trend.

Consumption of nitrate and phosphate by N. oleoabundans cultivated in exhausted and control growth media

The consumption of nitrate and phosphate by N. oleoabundans cells in the course of the experiment is depicted in Fig 3. At time 0 (inoculation), the exhausted replenished growth media contained

- 2- approximately 2 mM NO3 and 0.115 mM PO4 , i.e., the typical concentration of those components in BM medium. By the end of the experiment, cells had consumed practically all nitrate and phosphate present in the growth media. However, nutrient consumption by C, E+ and EG+ algal cultures followed different trends, since C samples maintained higher nitrate and phosphate concentrations than samples in exhausted media. In fact, after 12 days of cultivation, cells grown in

E+ and EG+ had consumed about 34% and 68% of nitrate, respectively, while C only 15%. Nitrate concentration, decreased from 1.43 mM at time 0 for all culture media to 0.94 mM for E+, 0.45 mM for EG+ and 1.21 mM for C media (p <0.001 in both treated samples relative to C; Fig 3a).

Differently, during the first 12 days of growth, phosphate exhibited a dramatic decrease, with a reduction by cells cultivated in EG+ and E+ of 94% and 98%, respectively, while in C medium it was only about 60%. In fact, phosphate concentration decreased from about 0.086 mM at time 0 for all culture media to 0.002 for E+, 0.005 for EG+ and 0.035 mM for C media (p <0.001 in both treated samples relative to C; Fig 3b).

Photosynthetic pigment content in N. oleoabundans cultivated in exhausted and control media

14 During the experiment a gradual increase in photosynthetic pigment content was observed in all samples (Fig 4). Chla in algae grown in C medium showed an evident increase up to the end of the experiment, reaching an average concentration of 0.5 nmol 10-6 cells (Fig 4a). A similar increasing trend was also observed in E+ samples, though with values 10-15% lower than those of controls.

From the 17th day of cultivation, Chla content of E+ samples was significantly lower than in C

(32%; p<0.01). Conversely, in EG+ samples Chla content showed a slightly increasing trend as compared with C and E+ samples; this led to a final value of 0.35 nmol 10-6 cells at the end of the experiment (32 and 21% lower than C and E+ samples, respectively; Fig 4a). An increasing trend in

Chlb content was also observed for all samples, without significant differences between C and E+ samples (Fig 4b). Similar to Chla content, EG+ cells contained lower quantities of Chlb than C and

E+ samples (15-25% and 20-35% lower than C and E+ samples, respectively). More evident differences were observed for Car content (Fig 4c). In fact, C samples showed an evident increase up to the end of the experiment, always containing higher quantities compared to the other samples

(20-45% and 15-30% more than E+ and EG+ samples, respectively; p<0.05). It is noteworthy that

EG+ and E+ cells shared a similar trend of Car concentration during the experiment (Fig 4c).

Maximum quantum yield of PSII of N. oleoabundans cultivated in exhausted and control media

The variations in PSII maximum quantum yield measured during the experiments are shown in Fig

5. C, E+ and EG+ samples showed a slight increase of the FV/FM ratio during the first three days of growth, reaching values of ca. 0.70. Subsequently, C cells maintained stable values around 0.70-

0.75, while the FV/FM ratio of E+ and EG+ samples decreased drastically down to values below

0.50 at the end of experiment. In EG+ cells the decrease was dramatic already from the 3rd day of cultivation, reaching the lowest value after 17 days (ca. 0.30 for EG+ vs 0.75 for C and 0.50 for

E+; p <0.01 in both cases); thereafter, samples maintained stable values of ca. 0.35 until the end of the experiment. On the contrary, in E+ cells, FV/FM started to decrease very strongly only from the

15 7th day of cultivation until the 17th day (from about 0.70 at day 7 down to 0.50 at day 17). During

th th the subsequent experimental times (21 and 25 day), the FV/FM ratio of E+ cells remained stable around 0.45.

Morphological observations of N. oleoabundans cultivated in exhausted and control media

Light microscopy and Nile Red staining

Light microscopy of both control and treated samples showed that N. oleoabundans maintained similar cell morphology and dimensions throughout the experiment (Fig 6, 7). In fact, cells were almost spherical with a cell diameter of 3-5 μm. One cup-shaped chloroplast, containing a large pyrenoid, was present inside the cells (Fig 6a, c, e). Moreover, the chloroplast emitted an intense red fluorescence due to Chl (Fig 6b, d, f). Interestingly, after 25 days of cultivation, while cell size and shape remained substantially unchanged, all algal samples showed some translucent granulation at the cytoplasmic level (Fig 7a, c, e). In order to understand the nature of those granulations and to investigate if the recycled growth media could promote the production of lipid globules, all samples were periodically stained with the lipid-specific fluorochrome Nile Red. The reaction was positive only at the end of the experiment and the translucent globules were then unequivocally identified as lipid droplets (Fig 7b, d, f).

TEM observations

To investigate the morphological and cytological changes induced by cultivation in recycled growth media, the ultrastructure of C, E+ and EG+ samples was observed by TEM. At 12 days of cultivation, most of the C cell volume was occupied by a characteristic cup-shaped chloroplast.

Inside the plastid one large pyrenoid, which was crossed by two elongated and appressed thylakoids, was present (Fig 8a). In particular, the organelle contained starch in the shape of a shell around the pyrenoid, and showed the typical thylakoid organization (Fig 8b). Typically featured

16 chloroplasts, as described for cells grown in C, were observed in cells grown in E+ medium (Fig

8c); however, thylakoids were more appressed than in C (Fig 8d). EG+ samples showed a more strongly altered chloroplast structure (Fig 8e), as compared with C and E+ samples. Photosynthetic membranes showed different degrees of thylakoid appression: some portions of thylakoid membranes were appressed while others were loose and sometimes swollen (Fig 8f,g). In addition, large portions of the stroma were free of thylakoids and some plastoglobules in proximity of the thylakoid membranes were also visible (Fig 8g, h). Finally, in EG+ samples the pyrenoid lost its round shape and appeared malformed (Fig 8e).

Extracellular FFA composition in E and EG media

The extracellular FFA composition in the exhausted autotrophic (E) and mixotrophic (EG) media of

Polyamine content in E and EG media and in the corresponding N. oleoabundans cells cultivated in a 20-L PBR

As shown in Fig 9a, the PA composition of N. oleoabundans cells cultivated mixotrophically in the

PBR were characterized by higher putrescine, spermine, and, especially, spermidine levels compared to cells cultivated autotrophically. In fact, Spm, Put and Spd concentrations in

17 mixotrophic cells were ca. 2, 30 and 57 times higher, respectively, than those found in autotrophic cells (p<0.001). Conversely, the PA composition in E and EG medium was not significantly different. In fact, both media contained similar amounts of Put and Spm. Interestingly, spermidine was present in high concentration in both media (119 ± 16.91 and 101 ± 5.00 pmol mL-1 in E and

EG, respectively; Fig 9b).

Discussion

The recycling of culture medium has been proposed as a possible solution in order to reduce water consumption for algal cultivation, thereby making the process more economically feasible and environmentally sustainable (Yang et al. 2011; Hadj-Romdhane et al. 2013; Farooq et al. 2014).

Yang et al. (2011) estimated that the large-scale cultivation of the microalga Chlorella vulgaris in recycled culture medium could reduce water use by about 84%. Present results clearly suggest that

N. oleoabundans can also efficiently grow in exhausted growth media, especially in the autotrophic medium replenished with nitrate and phosphate (E+). Observing the consumption trend of these main nutrients (Fig. 3), it is clear that in the exhausted media their concentration decreased faster with respect to controls in parallel to the enhanced cell growth. Moreover, phosphate was consumed faster than nitrate by all samples, probably because of its 100 times lower concentration with respect to the latter nutrient (0.2 gL-1 of nitrate vs 0.002 gL-1 of phosphate; Baldisserotto et al.,

2012). Thus, in order to make the use of recycled culture medium feasible, exhausted media should be replenished with the correct ratio of nitrate and phosphate, as they are the main nutrients that guarantee cell growth (Stephens et al. 2010). This aspect was preliminarily verified by employing unmodified exhausted growth media (E and EG) to re-cultivate N. oleoabundans. In fact, cells showed a slight growth, reaching, at the end of the cultivation period, low cell densities (ca. 11 vs

32 × 106 cells mL-1 of C cultures) (data not shown). These cells accumulated intracellular lipids throughout the experiment (data not shown) as a consequence of the limitation of nitrate and

18 phosphate (Mata et al. 2010; Popovich et al. 2012). These results are consistent with those of Zhu et al. (2013), who found that C. zofingensis cultivated in recycled medium, without nitrogen and phosphorus, displayed enhanced lipid production compared to cultures with full nutrients.

In order to understand if the growth promotion of N. oleoabundans cultivated in E+ and EG+, relative to C, was influenced by some molecules released from the algae, a characterization of autotrophic (E) and mixotrophic (EG) media was performed. First of all, pH of the control and exhausted media was monitored. In particular, an increasing trend was observed in control, which reached the pH of the exhausted media at the end of the experiment. This was not surprisingly, considering that during the microalgal growth pH usually increases because of the consumption of

+ CO2 during the photosynthetic process (Zhang et al., 2014). However, considering that in E and

EG+ pH fluctuated around their initial values, or at least gradually decreased, it is not possible to attribute to pH any evident role in the cell growth promotion. In fact, Moheimani and Borowitzka

(2006) and Stephen et al. (2010) reported that molecules released from cells can positively/negatively alter cell metabolism and biomass production.

Here we show that both exhausted media contained the main PAs (putrescine, spermidine, and spermine). It is known that these plant growth regulators are involved in a great variety of developmental processes in plant cells, e.g., cell division and protein synthesis (Kaur-Sawhney et al. 2003; Kuznetsova et al. 2006). PA biosynthetic pathways have also been studied in green algae

(Cohen et al. 1984; Theiss et al. 2002; Fuell et al. 2010). Although algae produce “unusual” PAs, such as homospermidine and thermospermine (Hamana et al. 2013), the more common PAs (Put,

Spd and Spm), when added to the media, promoted growth and metabolism in C. vulgaris (Czerpak et al. 2003). PAs are also known to alleviate the effect of biotic and abiotic stress in plants as well as algae (Tate et al. 2013).

In the present work, a very strong increase in Put and, especially, Spd concentrations was observed in N. oleoabundans mixotrophic cells as compared to the autotrophic ones, both cultivated in a 20-L

PBR. The growth-promoting role of PAs could explain the faster (0-4 days) and stronger cell

19 density increase (ca. 33 x 106 cell mL-1) in mixotrophic cultures relative to the autotrophic ones (0-9 days; ca. 22 x 106 cell mL-1). The higher PA concentrations in mixotrophically grown cells also suggest that, under these culture conditions, algal cells may be better protected from stress-inducing factors, e.g., bacterial contamination. To our knowledge, the presence of PAs released into algal growth medium in PBRs has not been documented before. The release of PAs (predominantly Spd) from autotrophic and mixotrophic cells into their respective (E and EG) exhausted media was not significantly different. However, the presence of these plant growth regulators seems to contribute to the promotion of N. oleoabundans growth in both media replenished with nitrate and phosphate.

In fact, during the entire experiment, a higher cell density was observed in E+ relative to C medium.

Moreover, the EG+ samples showed a promotion of cell density, albeit similar to that of C.

Both autotrophic and mixotrophic exhausted media also contained FFAs, which could have induced a change in the normal metabolism of N. oleoabundans. This assumption is demonstrated by the more rapid consumption of nitrate and phosphate, the decreased photosynthetic pigments content and the strong decline in the FV/FM ratio observed in cells grown in E+ and EG+ media. These results corroborate the assumption that microalgae release metabolites in high cell density cultures

(Richmond 2004) or as a consequence of stressful conditions (Ikawa 2004; Wu et al. 2006). Even if the role of FFAs is currently under debate (Ikawa 2004; Stephen et al. 2010), recent studies showed that they can strongly inhibit growth or exert cytotoxic effects on microalgae (Kogteva and

Bezuglov 1998; Wu et al. 2006; Bosma et al. 2008; Stephen et al. 2010). In fact, the presence of these metabolites, and their oxidative products, negatively affect biomass productivity, especially when the microalgae are cultivated in recycled medium (Lívanský et al. 1996; Rodolfi et al. 2003).

Granum et al. (2002) reported that the accumulation of intracellular lipids in microalgae enhanced the release of FFAs into the culture medium, and Harun et al. (2010) observed that this release was caused by cell lysis.

In this work, FFA composition in autotrophic (E) and mixotrophic (EG) media after N. oleoabundans cultivation was similar. The only main difference was represented by the presence of

20 the monoenoic oleic acid (C18:1ω9) in EG medium. Oleic acid is one of the major lipid components in lipid-enriched N. oleoabundans grown under N-stress conditions (Popovich et al.

2012). It is reported that mixotrophic cultivation alters N:C ratio, inducing similar lipid production as N-depleted autotrophic cultures (Giovanardi et al. 2014). Probably for this reason, the release of oleic acid occurred only in the mixotrophic growth medium. Moreover, Wu et al. (2006) observed altered plasma membrane permeability due to the toxic effects of FFAs in two Chlorophyta (C. Formatted: Font: Italic vulgaris and Monoraphidium contortum) and in a cyanobacterium (Anabaena P-9). Then, in addition to the lower concentration with respect to nitrate in BM medium, Tthe rapid consumption of phosphate in both exhausted media could be linked also to an alteration in membrane permeability, due to the activity of FFAs (Ikawa 2004; Wu et al. 2006). In addition, FFAs can cause inhibition of the PSII and PSI electron transport chains (Siegenthaler 1973) and disorganization of thylakoids (Wu et al. 2006). Indeed, N. oleoabundans cells in E+ and EG+ were characterized by alterations of the photosynthetic apparatus. These cells, in fact, contained less photosynthetic pigments than C cells, and exhibited a drastic decrease of the PSII maximum quantum yield

(FV/FM). These variations are linked to an alteration of photosynthetic efficiency, especially as regards PSII (White et al. 2011). The decreased photosynthetic pigment contents, observed in cells grown in E+ and EG+, also reflected the changes in thylakoid membrane arrangement

(Baldisserotto et al. 2012). Therefore, correct thylakoid organization is necessary to maintain optimal photosynthetic activity and this is often influenced by culture conditions (Nevo et al. 2012).

In addition, the presence of some plastoglobules in chloroplasts of algal cells, grown on exhausted media, could be another indicator of an alteration of photosynthetic membranes (Besagni and

Kessler 2013). On the contrary, N. oleoabundans cells grown in C medium showed stable and normal FV/FM values throughout the experiment (Giovanardi et al. 2014), and the typical assembly of thylakoid membranes (Nevo et al. 2012; Baldisserotto et al. 2012; Giovanardi et al. 2014).

The presence of lipid droplets, observed only at the end of the experiment in C, E+ and EG+ cells, is probably related to aging of the microalgal cultures (Hu et al. 2008; Baldisserotto et al. 2012),

21 rather than to the use of recycled culture media.

In conclusion, results presented here demonstrate that recycling autotrophic and mixotrophic growth media is a suitable solution to obtain high cell density cultures of the microalga N. oleoabundans. For this reason, they represent a contribution for improving the scale-up of microalgal cultivation while providing a more sustainable ecological impact on water resources.

However, further studies are needed to deepen knowledge on the nature and the specific role of some molecules that are released into the growth media, in order to obtain useful information for the advancement of the biotechnological use of this strain.

Acknowledgements

This work was financially supported by grants from the University of Ferrara. We are grateful to

Dr. Immacolata Maresca of the University of Ferrara for technical assistance in freeze-drying of samples.

Conflict of interest

The authors declare they have no conflict of interest.

22 References:

Baldisserotto C, Ferroni L, Giovanardi M, Pantaleoni L, Boccaletti L, Pancaldi S (2012) Salinity

promotes growth of freshwater Neochloris oleoabundans UTEX 1185 (Sphaeropleales, Formatted: Font: Italic

Neochloridaceae): morpho-physiological aspects. Phycologia 51:700-710

Baldisserotto C, Giovanardi M, Ferroni L, Pancaldi S (2014) Growth, morphology and

photosynthetic responses of Neochloris oleoabundans during cultivation in a mixotrophic

brackish medium and subsequent starvation. Acta Physiol Plant 36:461-472

Batan L, Quinn CJ, Bradley TH (2013) Analysis of water footprint of a photobioreactor microalgae

biofuel production system from blue, green and lifecycle perspectives. Algal Research 2:196-

203

Besagni C and Kessler F (2013) A mechanism implicating plastoglobules in thylakoid disassembly

during senescence and nitrogen starvation. Planta 237:463-470

Borowitzka MA (2005) Culturing microalgae in outdoor ponds. In: Anderson RA (ed) Algal

culturing techniques. Elsevier Academic, London, pp 205-218

Borowitzka MA and Moheimani NR (2013) Sustainable biofuels from algae. Mitig Adapt Strateg

Glob Change 18:13-25

Bosma R, Miazek K, Willemsen SM, Vermuë MH, Wijffels RH (2008) Growth inhibition of

Monodus subterraneus by free fatty acids. Biotechnol Bioeng 101:1108–1114

Chisti Y (2007) Biodiesel from microalgae. Biotech Adv 25:294-306

Czerpak R, Bajguz A, Pietrowska A, Dobrogowska R, Matejczyk M, Weislawski W (2003)

Biochemical activity of di- and polyamines in the green alga Chlorella vulgaris Beijerinck

(Chlorophyceae). Acta Soc Bot Pol 72:19–24 Formatted: Font: Italic

Cohen E, Arad SM, Heimer YH, Mizrahi Y (1984) Polyamine biosynthetic enzymes in the cell cycle

of Chlorella. Plant Physiol 74:385-388

Farooq W, Suh WI, Park MS, Yang J (2014) Water use and its recycling in microalgae cultivation for

biofuel application. Bioresource Technol doi: 10.1016/j.biortech.2014.10.140

23 Ferroni L, Baldisserotto C, Pantaleoni L, Billi P, Fasulo MP, Pancaldi S (2007) High salinity alters

chloroplast morpho-physiology in a freshwater Kichneriella species (Selenastraceae) from Formatted: Font: Not Italic Formatted: Font: Italic Ethipian Lake Awasa. Am J Bot 94:1973-1983

Ferroni L, Baldisserotto C, Giovanardi M, Pantaleoni L, Morosinotto T, Pancaldi S (2011) Revised

assignment of room-temperature chlorophyll fluorescence emission bands in single living cells

of Chlamydomonas reinhardtii. J Bioenerg Biomembr 43:163-173

Fuell C, Elliott KA, Hanfrey CC, Franceschetti M, Michael AJ (2010) Polyamine biosynthetic

diversity in plants and algae. Plant Physiol Biochem 48:513–520

Giovanardi M, Ferroni L, Baldisserotto C, Tedeschi P, Maietti A, Pantaleoni L, Pancaldi S (2013)

Morpho-physiological analyses of Neochloris oleoabundans (Chlorophyta) grown Formatted: Font: Italic

mixotrophically in a carbon-rich waste product. Protoplasma 250:161-174

Giovanardi M, Baldisserotto C, Ferroni L, Longoni P, Cella R, Pancaldi S (2014) Growth and lipid

synthesis promotion in mixotrophic Neochloris oleoabundans (Chlorophyta) cultivated with Formatted: Font: Italic

glucose. Protoplasma 251:115-125

Gouveia L, Marques AE, Lopes da Silva T, Reis A (2009) Neochloris oleoabundans UTEX #1185: a

suitable renewable lipid source for biofuel production. J Ind Microbiol Biotechnol 36: 821-826

Granum E, Kirkvold S, Myklestad SM (2002) Cellular and extracellular production of carbohydrates

and amino acids by the marine diatom Skeletonema costatum: diel variations and effects of N

depletion. Mar Ecol Prog Ser 242:83–94.

Grobbelaar JU (2009) Factors governing algal growth in photobioreactors: the “open” versus

“closed” debate. J Appl Phycol 21:489-492

Hadj-Romdhane F, Jaouen P, Pruvost J, Grizeau D, Van Vooren G, Bourseau P (2012) Development

and validation of a minimal growth medium for recycling Chlorella vulgaris culture.

Bioresource Technol 123:366-374

Hadj-Romdhane F, Zheng X, Jaouen P, Pruvost J, Grizeau D, Croué JP, Bourseau P (2013) The

culture of Chlorella vulgaris in a recycled supernatant: Effects on biomass production and

24 medium quality. Bioresource Technol 132: 285–292

Hamana K, Niitsu M, Hayashi H (2013) Occurrence of homospermidine and thermospermine as a

cellular polyamine in unicellular chlorophyte and multicellular charophyte green algae. J Gen

Appl Microbiol 59: 313-319

Harun R, Singh M, Forde GM, Danquah MK (2010) Bioprocess engineering of microalgae to

produce a variety of consumer products. Renew Sust Energ Rev 14:1037-1047

Hu Q, Sommerfeld M, Jarvis E, Ghirardi M, Posewitz M, Seibert M, Darzins A (2008) Microalgal

triacylglycerols as a feedstocks for biofuel production: perspective and advances. Plant J

54:621-639

Ikawa M (2004) Algal polyunsaturated fatty acids and effects on plankton ecology and other

organism. UNH Center for Freshwater Biology Research 6:17-44

Kalaji HM, Schansker G, Ladle RJ, Vasilij Goltsev V, Bosa K, Allakhverdiev SI, Brestic M, Bussotti

F, Calatayud A, Dąbrowski P, Elsheery NI, Ferroni L, Guidi L, Hogewoning SW, Jajoo A,

Misra AN, Nebauer SG, Pancaldi S, Penella C, Poli D, Pollastrini M, Romanowska-Duda ZB,

Rutkowska B, Serôdio J, Suresh K, Szulc W, Tambussi E, Yanniccari M, Zivcak M (2014)

Frequently asked questions about in vivo chlorophyll fluorescence: practical issues. Photosynth

Res 122:121-158

Kaur-Sawhney R, Tiburcio AF, Altabella T, Galston AW (2003) Polyamines in plants: an overview.

J Cell Mol Biol 2:1-12

Kogteva GS and Bezuglov VV (1998) Unsaturated fatty acids as endogenous bioregulators.

Biochemistry (Moscow) 63:4-12

Lam MK and Lee KT (2012) Potential of using organic fertilizer to cultivate Chlorella vulgaris for

biodiesel production. Appl Energy 94:303-308

Li Y, Horsman M, Wang B, Wu N, Lan CQ (2008) Effect on nitrogen sources on cell growth and

lipid accumulation of green alga Neochloris oleoabundans. Appl Microbiol Biotechnol 81:629-

636

25 Lichtenthaler HK, Buschmann C, Knapp M (2005) How to correctly determine the different

chlorophyll fluorescence parameters and the chlorophyll fluorescence decrease ratio RFd of

leaves with the PAM fluorometer. Photosynthetica 43:379-393

Lívanský K, Dědič K, Bínová J, Tichý V, Novotný P, Doucha J (1996) Influence of the nutrient

solution recycling on the productivity of Scenedesmus obliquus, utilization of nutrients and

water in outdoor cultures. Algological Studies/ Archiv für Hydrobiologie 81:105–113

Mata TM, Martins AA, Caetano NS (2010) Microalgae for biodiesel production and other

applications: a review. Renew Sust Energ Rev 14:217- 232

Moheimani NR and Borowitzka MA (2006) The long-term culture of the coccolithophore

Pleurochrysis carterae (Haptophyta) in outdoor raceway ponds. J Appl Phycol (2006) 18:703- Formatted: Font: Italic

712

Molina Grima E, Belarbi H, Acién Fernández FG, Robles Medina A, Chisti Y (2003) Recovery of

microalgal biomass and metabolites: process options and economics. Biotech Adv 20: 491-515

Ndimba BK, Ndimba RJ, Johnson TS, Waditee-Sirisattha R, Baba M, Sirisattha S, Shiraiwa Y,

Rakwal R, Agrawal GK, Rakwal R (2013) Biofuels as a sustainable energy source: An update

of the applications of proteomics in bioenergy crops and algae. J Prot 93:234-244

Nevo R, Charuni D, Tsabari O, Reich Z (2012) Composition, architecture and dynamics of the

photosynthetic apparatus in higher plants. Plant J 70:157-176

Pancaldi S, Baldisserotto C, Ferroni L, Bonora A, Fasulo MP (2002) Room temperature

microspectrofluorimetry as a useful tool for studying the assembly of the PSII chlorophyll–

protein complexes in single living cells of etiolated Euglena gracilis Klebs during the greening

process. J Exp Bot 53:1753–1763

Popovich CA, Damiani MC, Constenla D, Martínez AM, Giovanardi M, Pancaldi S, Leonardi PI

(2012) Neochloris oleoabundans grown in natural enriched seawater for biodiesel feedstock:

Evaluation of its growth and biochemical composition. Bioresource Technol 114:287-293

Pruvost J, Van Vooren G, Cogne G, Legrand J (2009) Investigation of biomass and lipids production

26 with Neochloris oleoabundans in photobioreactor. Bioresource Technol 100:5988-5995

Pulz O (2001) Photobioreactors: production systems for phototrophic microorganisms. Appl

Microbiol Biotechnol 57:287-293

Richmond A (2004) Handbook of Microalgal Culture: Biotechnology and Applied Phycology.

Blackwell Sciences Ltd., Oxford

Rodolfi L, Zittelli GC, Barsanti L, Rosati G, Tredici MR (2003) Growth medium recycling in

Nannochloropsis sp. mass cultivation. Biomol Eng 20:243–248 Formatted: Font: Italic

Scaramagli S, Biondi S, Capitani F, Gerola P, Altamura MM, Torrigiani P (1999) Polyamine

conjugate levels and ethylene biosynthesis: inverse relationship with vegetative bud formation

in tobacco thin layers. Physiol Plant 105:366-375

Sheehan JJ (2009) Biofuels and the conundrum of sustainability. Curr Opin Biotechnol 20:318–324

Siegenthaler PA (1973) Change in pH dependence and sequential inhibition of photosynthetic

activity in chloroplasts by unsaturated fatty acids. Biochim Biophys Acta 305:153-162

Smith VH, Sturm BSM, deNoyelles FJ, Billings SA (2010) The ecology of algal biodiesel

production. Trends Ecol Evo 25:301-309

Solomon BD (2010) Biofuels and sustainability. Ann NY Acad Sci 1185:119-134

Stephens E, Ross IL, Mussgnug JH,Wagner LD, BorowitzkaMA, Posten C, Kruse O, Hankamer B

(2010) Future prospects of microalgal biofuel production systems. Trends Plant Sci 15:554-564

Tate JJ, Gutierrez-Wing MT, Rush KA, Benton MG (2013) The Effects of Plant Growth Substances

and Mixed Cultures on Growth and Metabolite Production of Green Algae Chlorella sp.: A

Review. J Plant Growth Regul 32:417–428

Tornabene TG, Holzer G, Lien S, Burris N (1983) Lipid composition of the nitrogen starved green

alga Neochloris oleoabundans. Enzyme Microbiol Tech 5:435-440

Theiss C, Bohley P, Voigt J (2002) Regulation by polyamines of ornithine decarboxylase activity and

cell division in the unicellular green alga Chlamydomonas reinhardtii Plant Physiol 128:1470-

1479

27 Wellburn AR (1994) The spectral determination of chlorophylls a and b, as well as total carotenoids,

using various solvents with spectrophotometer of different resolution. J Plant Physiol 144:307–

313

White S, Anandraj A, Bux F (2011) PAM fluorometry as a tool to assess microalgal nutrient stress

and monitor cellular neutral lipids. Bioresource Technol 102:1675-1682

Wu JT, Chiang YR, Huang WY, Jane WN (2006) Cytotoxic effects of free fatty acids on

phytoplankton algae and cyanobacteria. Aquat Toxicol 80:338–345

Yang, J, Xu M, Zhang X, Hu Q, Sommerfeld M, Chen Y (2011) Lifecycle analysis on biodiesel

production from microalgae: Water footprint and nutrients balance. Bioresource Technol

102:159-165

Zhang Q, Wang T, Hong Y (2014) Investigation of initial pH effects on growth of an oleaginous

microalgae Chlorella sp. HQ for lipid and nutrient uptake. Water Sci Technol 70:712-719

Zhu LD, Takala J, Hiltunen E, Wang ZM (2013) Recycling harvest water to cultivate Chlorella Formatted: Font: Italic

zofingiensis under nutrient limitation for biodiesel production. Bioresource Technol 144:14-20

28 Figure captions:

Fig 1 (a-e) Growth kinetics (a) and morphology (b-e) of N. oleoabundans cells autotrophically cultivated in a 20-L PBR in BM medium. Light (b, d) and fluorescence microscope observations (c, e) of N. oleoabundans cells after 3 (b, c) and 25 (d, e) days of autotrophic cultivation. (f-j) Growth kinetics (f) and cell morphology (g-j) of N. oleoabundans cultivated in a 20-L PBR in BM medium under mixotrophic conditions (supplemented with 2.5 gL-1 of glucose). Light (g, i) and fluorescence microscope observations (h, j) of N. oleoabundans cells after 3 (g, h) and 8 (d, e) days of mixotrophic cultivation. In both graphs, curves are constructed on a log2 scale and data are means ± s.d. (n=3). In all micrographs, bars: 2 μm

Fig 2 (a) Growth kinetics of N. oleoabundans in BM medium (filled circles), E+ medium (empty squares) and EG+ medium (empty diamonds). (b) Growth rates, calculated during the exponential phase

(0–3 days time interval of cultivation), of cells grown in BM (white), E+ (light grey) and EG+ (dark grey) media. The growth curve is constructed on a log2 scale and data are means ± s.d. (n=3)

Fig 3 Time-course variations in the concentrations of nitrate (a) and of phosphate (b) in culture media of

N. oleoabundans during 25 days of cultivation on BM medium (white), E+ medium (light grey) and

EG+ medium (dark grey). Values are means ± s.d. (n=3)

Fig 4 Time-course variations of Chla (a), Chlb (b) and Car content (c) in N. oleoabundans cells grown in BM (filled circles), E+ (empty squares) and EG+ (empty diamonds) media during the 25 days of cultivation. Values are means ± s.d. (n=3)

Fig 5 Time-course variations of PSII maximum quantum yield (FV/FM ratio) in N. oleoabundans cells grown in BM (filled circles), E+ (empty squares) and EG+ (empty diamonds) media during the 25 days of cultivation. Values are means ± s.d. (n=3)

29 Fig 6 Light and fluorescence microscopy observations of N. oleoabundans cells at 12 days of cultivation. (a) Control cells and (b) the relative fluorescence of the chloroplast; (c) cell grown in

E+ medium and (d) the relative fluorescence of the chloroplast; (e) cell grown in EG+ medium and

(f) the relative fluorescence of the chloroplast. In all micrographs, bars: 2 μm

Fig 7 Light and epifluorescence pictures of N. oleoabundans cells after 25 days of growth. (a)

Control cells and (b) relative Nile Red-staining observation, (c) cells grown in E+ medium and (d) relative Nile Red-staining observation, and (e) cells grown in E+ medium and (f) relative Nile Red- staining observation in EG+ medium. Translucent globules are indicated with arrows. In all micrographs, bars: 2 μm

Fig 8 Transmission electron micrographs of N. oleoabundans cells at 12 days of cultivation. (a) Cell grown in BM medium and (b) detail of its chloroplast showing the typical thylakoid membranes organization (white arrow). (c) Cell grown in E+ medium and (d) detail of its chloroplast with quite compact and appressed (white arrow) thylakoids. (e) Cell grown in EG+ medium and (f-h) details of its chloroplast. The presence of intermediate stages of thylakoid membranes are evident in EG+ cells, showing a general disorganisation of thylakoid membranes (f-h), which appeared wavy, loose and sometimes swollen (f, g). (h) Some plastoglobules in proximity of thylakoid membranes are also observed (arrowheads). P, pyrenoid; N, nucleus, M, mithochondrion. Bars: 0.5 μm (a, c, e);

0.05 μm (b, d, f, g, h)

Fig 9 Polyamine concentrations (a) in N. oleoabundans cells cultivated in a 20-L PBR autotrophically and mixotrophically at 15 and 8 days of cultivation, respectively, and (b) in the corresponding exhausted E and EG growth media. Autotrophic cells (white), mixotrophic cells (dark),

E+ (light grey) and EG+ medium (dark grey). Values are means ± s.d. (n=3)

30 Manuscript Click here to download Manuscript: Sabia et al. Revised MS.doc Click here to view linked References

1 2 Introduction 3 4 5 6 Over the last decades, microalgae have received much attention as promising high-potential 7 8 feedstock for biodiesel (Chisti 2007; Li et al. 2008; Borowitzka and Moheimani 2013) and as an 9 10 11 attractive raw material for the production of a wide range of high value bioproducts (Molina Grima 12 13 et al. 2003; Mata et al. 2010). They are photosynthetic microorganisms with numerous key 14 15 16 features, which potentially make them a more appreciable renewable source than terrestrial plants 17 18 (Richmond 2004; Hu et al 2008; Ndimba et al. 2013). For instance, these microorganisms have 19 20 faster growth rates, higher photosynthetic efficiencies, higher rates of carbon dioxide fixation, and 21 22 23 higher biomass productivities compared to plants (Richmond 2004; Gouveia et al. 2009; Harun et 24 25 al. 2010; Mata et al. 2010). Moreover, microalgae can thrive in non-arable lands being able to grow 26 27 28 in waters having different salinity levels and chemical composition (Smith et al. 2010; Borowitzka 29 30 and Moheimani 2013). 31 32 33 The environmental and economical impact of large-scale production of microalgae has been widely 34 35 discussed, with the aim of making the process increasingly sustainable (Sheehan 2009; Solomon 36 37 2010). In this perspective, microalgal cultivation systems are crucial factors to be taken into account 38 39 40 (Smith et al. 2010; Stephens et al. 2010; Borowitzka and Moheimani 2013). A wide variety of 41 42 systems has been described by Mata and co-workers (2010) and recently by Borowitzka and 43 44 45 Moheimani (2013). Microalgae can be cultivated in open pond reactors or closed photobioreactors 46 47 (PBRs), depending on algal strain, issue of research, type of desired products and environmental 48 49 50 conditions (Chisti 2007; Mata et al. 2010; Borowitzka and Moheimani 2013). Although open ponds 51 52 are the most commonly used culture systems, due to their low cost of production and operation, 53 54 55 recently closed PBRs are receiving much attention for the possibility of producing valuable 56 57 compounds (Pulz 2001; Grobbelaar 2009; Smith et al. 2010). However, algal cultures require large 58 59 amounts of water in both culture systems (Lam and Lee 2012; Borowitzka and Moheimani 2013). 60 61 62 63 64 3 65 Considering that water is becoming a scarce natural resource, water demand represents an important 1 2 factor when performing large-scale cultivation (Batan et al. 2013; Farooq et al. 2014). In order to 3 4 5 reduce the high production costs and, at the same time, to make microalgal production more 6 7 environmentally sustainable, recycling culture medium has been proposed as a possible solution 8 9 10 (Yang et al. 2011; Hadj-Romdhane et al. 2012; Farooq et al. 2014), especially in large-scale culture 11 12 systems (Borowitzka 2005; Lam and Lee 2012; Zhu et al. 2013). However, the feasibility of 13 14 recycling growth medium to re-cultivate microalgae has been tested only in a few cases (Lívanský 15 16 17 et al. 1996; Rodolfi et al. 2003; Hadj-Romdhane et al. 2013; Zhu et al. 2013). It is reported that the 18 19 recycling medium could have negative effects on biomass productivity, due to the release of 20 21 22 inhibitory secondary metabolites (Richmond 2004). Harmful metabolites are, in fact, released 23 24 during microalgal growth under physiological stress (Ikawa 2004; Moheimani and Borowitzka 25 26 27 2006). Free fatty acids (FFAs), and substances derived from the photooxidation of unsaturated fatty 28 29 acids, are the most common metabolites with inhibitory effects on microalgae (Ikawa 2004; Wu et 30 31 32 al. 2006; Bosma et al. 2008; Stephen et al. 2010). 33 34 In the present work, the suitability of recycling growth media for the re-cultivation of Neochloris 35 36 oleoabundans (Chlorophyta) was investigated. This species was chosen because it is considered one 37 38 39 of the most promising oil-rich microalgae, due to its capability to accumulate lipids when grown 40 41 under nitrogen starvation (Tornabene et al. 1983; Chisti 2007; Li et al. 2008; Pruvost et al. 2009) or 42 43 44 mixotrophically, in the presence of glucose, or carbon-rich wastes, as organic carbon sources 45 46 (Giovanardi et al. 2013; Baldisserotto et al. 2014). Present results clearly indicate that N. 47 48 49 oleoabundans can efficiently grow in its exhausted growth medium, if suitably replenished with the 50 51 main nutrients. Based on this encouraging result, and with the aim of gaining further knowledge on 52 53 the morpho-physiological aspects and biotechnological applications of this algal strain, the 54 55 56 following features were analyzed: (1) growth kinetics in parallel to nitrate and phosphate 57 58 consumption; (2) photosynthetic pigment content; (3) PSII maximum quantum yield; (4) cell 59 60 61 morphology, with special attention to intracellular lipid accumulation, (5) FFA accumulation in 62 63 64 4 65 recycled growth media, and (6) polyamine (PA) concentration both inside cells and in the 1 2 corresponding culture media. PAs were determined in order to understand if these plant growth 3 4 5 regulators could be responsible for the growth promotion of N. oleoabundans in recycled growth 6 7 medium. It is known, in fact, that PAs, together with other plant growth regulators, have stimulatory 8 9 10 effects on algal growth and metabolism, and are involved in mitigating various types of biotic and 11 12 abiotic stress (Tate et al. 2013). 13 14 15 16 17 18 19 Materials and Methods 20 21 22 Algal strains and culture condition 23 24 25 The strain used in this study was the Chlorophyta N. oleoabundans UTEX 1185 (syn. Ettlia 26 27 oleoabundans) (Sphaeropleales, Neochloridaceae), obtained from the Culture Collection of Algae 28 29 30 of the University of Texas (UTEX, Austin, Texas, USA; www.utex.org). 31 32 N. oleoabundans was cultivated in axenic liquid brackish medium (BM) (Baldisserotto et al. 2012) 33 34 35 in a coaxial 20-L capacity PBR (M2M Engineering, Grazzanise, Caserta, Italy). Algae were 36 37 cultivated autotrophically in BM for 25 days or mixotrophically, by addition of 2.5 g L-1 glucose, 38 39 for 8 days, according to previously described protocols (Baldisserotto et al. 2014; Giovanardi et al. 40 41 42 2014). For autotrophic cultivation, cells were inoculated into the PBR to obtain an initial cell 43 44 density of about 0.6 x 106 cells mL-1, while for mixotrophic cultivation initial cell density was 45 46 6 -1 47 higher (10 x 10 cells mL ). Culture conditions in the PBR were: 24 ± 1°C; sterile air injection at 48 -2 -1 49 the bottom of the PBR, with 0.5/3.5 h bubbling/static cycles; irradiance 65 µmolphotons m s of 50 51 52 PAR (16:8 h light:dark photoperiod). Light was supplied with inner cool–white fluorescent Philips 53 54 tubes. 55 56 Algal growth and morphology in the PBR were monitored as described below. 57 58 59 60 61 62 63 64 5 65

1 2 Preparation of growth media 3 4 5 Experiments were performed by testing the following media: 6 7 8 - C: freshly prepared BM medium (control); 9 10 - E: autotrophic exhausted medium; 11 12 13 - EG: mixotrophic exhausted medium after cultivation with glucose; 14 15 - E+: autotrophic exhausted medium replenished with nitrate and phosphate concentrations as for 16 17 BM medium; 18 19 20 - EG+: mixotrophic exhausted medium after cultivation with glucose and replenished with nutrients 21 22 as in E+. 23 24 25 In order to obtain the exhausted media for the recycling experiment, N. oleoabundans was grown 26 27 inside the PBR under the culture conditions described in the previous section. For E+ medium 28 29 th 30 preparation, about 500 mL of autotrophic algal culture in the stationary phase of growth (at the 15 31 32 day of cultivation in the PBR) were centrifuged at 2,000 g for 10 min in order to separate the 33 34 35 medium from the algae, thus obtaining an autotrophic exhausted medium (E). The medium 36 37 appeared pale-yellow and was free from algae, bacteria and protozoa; its optical density at a 38 39 th wavelength of 750 nm (OD750) was 0.02. For EG+ medium, at the 8 day of cultivation in the PBR, 40 41 42 the same aliquots of algal culture were harvested by centrifugation (2,000 g for 10 min). In this 43 44 case, the mixotrophic exhausted medium (EG) was straw-yellow in colour and presented a weak 45 46 47 bacterial contamination due to the glucose addition (OD750 =0.009). Glucose was proved to be 48 49 totally consumed during the previous cultivation inside the PBR by 3,5-dinitrosalicylic (DNS; 50 51 52 Sigma-Aldrich, Gallarate, Milan, Italy) acid assay, according to Giovanardi et al. (2014). In both 53 54 exhausted media, pH was not adjusted. After determining the nitrate and phosphate concentrations 55 56 of the two exhausted media (see “Nitrate and phosphate analyses”), KNO and K HPO were 57 3 2 4 58 -1 59 axenically added to reach final concentrations of 0.2 and 0.02 g L , respectively, i.e., the typical 60 61 concentration of those components in BM, thus obtaining the replenished exhausted media (E+ and 62 63 64 6 65 EG+) used for experiments. 1 2 3 4 5 Experimental design 6 7 8 When autotrophic cultures of N. oleoabundans, grown in BM in the PBR, reached a cell density of 9 10 10 x 106 cells mL-1 (after about 9 days of cultivation), aliquots of cells were inoculated into 300-mL 11 12 13 Erlenmeyer flasks (150 mL total volume) containing C, E+, EG+ media, to obtain an initial cell 14 15 density of about 0.5 x 106 cells mL-1. The cultures were placed in a growth chamber (24 ± 1 °C, 80 16 17 μmol m-2 s-1 of PAR with a 16:8 h light-darkness photoperiod), and cultivated with continuous 18 photons 19 20 shaking at 80 rpm, without external CO2 supply. Experiments were performed in triplicate. Aliquots 21 22 of cultures (cells and media) were collected at different times of cultivation up to 25 days, 23 24 25 depending on the analysis. 26 27 28 29 30 Analyses 31 32 33 Growth evaluation 34 35 Aliquots of cell samples cultivated in control and exhausted media were counted at 0, 3, 7, 12, 17 36 37 and 25 days of cultivation using a Thoma haemocytometer (HBG, Giessen, Germany) under a light 38 39 40 microscope (Zeiss, model Axiophot) and growth curves were obtained. 41 42 The growth rate (μ, number of divisions per day) during the exponential phase was calculated with 43 44 45 the following equation: 46 47 -1 μ (div d ) = (log2N1 − log2N0) / (t1− t0), 48 49 50 where μ is the growth rate, N1 the cell number at time t1, N0 the cell number at time 0 and t1−t0 the 51 52 time interval (days) (Giovanardi et al. 2013). 53 54 55 In parallel to growth, pH was also periodically monitored on small aliquots of samples. 56 57 58 59 60 61 62 63 64 7 65

1 2 Nitrate and phosphate analyses 3 4 5 After 12 and 25 days of cultivation, samples of C, E+ and EG+ media were harvested by 6 7 centrifugation to analyze nitrate and phosphate concentrations. These nutrients were quantified 8 9 10 colorimetrically using a flow-injection autoanalyzer (FlowSys, Systea, Roma, Italy). 11 12 13 14 Photosynthetic pigment extraction and quantification 15 16 17 For photosynthetic pigment analysis, cell samples were collected by centrifugation after 0, 7, 17, 21 18 19 and 25 days of cultivation. Extraction of photosynthetic pigments was performed according to 20 21 22 Baldisserotto et al. (2014). The extracts were measured with a Pharmacia Ultrospec 2000 UV-Vis 23 24 spectrophotometer (1-nm bandwidth; Amersham Biosciences, Piscataway, NJ, USA) at 666 25 26 27 (chlorophyll a - Chla), 653 (chlorophyll b - Chlb) and 470 nm (carotenoids - Car). Quantification 28 29 was performed according to equations reported in Wellburn (1994). 30 31 32 33 34 PAM fluorimetry 35 36 The PSII maximum quantum yield of algae was determined at the same cultivation times 37 38 39 considered for growth kinetics measurements. A pulse amplitude modulated fluorometer (ADC 40 41 Bioscientific Ltd, Hoddesdon, Hertfordshire, UK) was used to determine the in vivo chlorophyll 42 43 44 fluorescence of PSII. The PSII maximum quantum yield is reported as FV/FM ratio, i.e. (FM – F0)/ 45 46 FM, where variable fluorescence is FV = (FM−F0), FM is the maximum fluorescence and F0 is the 47 48 49 initial fluorescence of samples (Lichtenthaler et al. 2005). This measurement is considered a valid 50 51 method to probe the maximum quantum yield of photochemistry in PSII (Kalaji et al. 2014). 52 53 Moreover, it is useful to estimate the physiological state of plants and microorganisms also under 54 55 56 nutrient stress (White et al. 2011). Samples were prepared as reported in Ferroni et al. (2011) after 57 58 15 min of dark incubation. 59 60 61 62 63 64 8 65

1 2 Light and fluorescence microscopy 3 4 5 For microscopic observations, cell samples were routinely collected throughout the cultivation 6 7 period inside the PBR and during the experiment with exhausted media. Aliquots of samples were 8 9 10 observed using a microscope (Zeiss, model Axiophot) with conventional and fluorescent 11 12 attachments. The light source for chlorophyll fluorescence observation was a HBO 100 W pressure 13 14 mercury vapour lamp (filter set, BP436/10, LP470). Pictures of cells were taken with a Canon 15 16 17 IXUS 110 IS digital camera (12.1 megapixels), mounted on the ocular lens through a Leica DC150 18 19 system (Leica Camera AG, Solms, Germany). 20 21 22 23 24 Lipid staining 25 26 27 During experiments, the intracellular presence of lipids was evaluated by staining cells with the 28 29 fluorochrome Nile Red (9-diethylamina-5Hbenzo[α]phenoxazine-5-one, 0.5 mg dissolved in 100 30 31 32 mL acetone; Sigma-Aldrich, Gallarate, Milan, Italy), as described in Giovanardi et al. (2014). After 33 34 incubation at 37°C in darkness for 15 min, cells were observed with the microscope described 35 36 above at an excitation wavelength of 485 nm (filter set, BP485, LP520). Photographs were taken 37 38 39 with the camera described above. 40 41 42 43 44 Transmission electron microscopy (TEM) 45 46 TEM observations were made on cells harvested by centrifugation (600 g, 10 min) at the 12th day of 47 48 49 cultivation. Cells were fixed, post-fixed and dehydrated as reported in Baldisserotto et al. (2012). 50 51 Embedding in resin and staining procedures were performed as previously described (Pancaldi et al. 52 53 2002). Sections were observed with a Hitachi H800 electron microscope (Hitachi, Tokyo, Japan) at 54 55 56 the Electron Microscopy Centre, University of Ferrara. 57 58 59 60 61 62 63 64 9 65

1 2 Extracellular free fatty acids analysis 3 4 5 To analyze the FFA composition of the exhausted E and EG media used to prepare E+ and EG+ 6 7 replenished media, aliquots of N. oleoabundans cultures grown autotrophically and mixotrophically 8 9 th th 10 in the PBR were collected at the 15 and at the 8 day of cultivation, respectively. Samples were 11 12 centrifuged at 2,000 g for 10 min in order to separate the medium from algae. Fifty mL of media 13 14 obtained from the autotrophic (E) and mixotrophic (EG) cultures were freeze-dried for analysis. 15 16 17 The extracellular FFA composition was determined in duplicate by gas chromatography-mass 18 19 spectrometry (GC-MS). For extraction, samples were dissolved in 3 mL of hexane, sonicated for 15 20 21 22 min and extracted overnight. Fatty acid methyl esters were prepared by transesterification with 1.5 23 24 mL of 5% of sodium hydroxide in methanol solution. Sample volumes of 1 μl were injected into the 25 26 27 GC-MS apparatus, which consisted of a Varian Saturn 2100 MS/MS ion trap mass spectrometer. 28 29 Separations were performed using a Zebron ZB-WAX Phenomenex capillary column (60 cm in 30 31 -1 32 length, 0.25 mm i.d) supplied with helium carrier gas at 1 mL min constant flow. The injector 33 34 temperature was 250 °C and the oven temperature program was the following: start 100°C for 2 35 36 min, ramp to 200°C at 10 °C/min, and hold for 108 min. The MS acquisitions were performed by 37 38 39 full scan mode. 40 41 42 43 44 Determination of polyamines 45 46 To analyze the free PA composition of N. oleoabundans cells cultivated autotrophically and 47 48 49 mixotrophically in PBR and in the corresponding E and EG media, samples were collected as 50 51 described for FFA analyses. Freeze-dried E or EG media and algae (50 and 20 mg, respectively) 52 53 were extracted in cold 4% perchloric acid, kept for 1 h on ice, and then centrifuged at 15,000 g for 54 55 56 15 min. Aliquots (200 L) of the supernatants and standard solutions of putrescine (Put), 57 58 spermidine (Spd) and spermine (Spm) were derivatised with dansyl chloride (Scaramagli et al. 59 60 61 1999). Dansylated derivatives were extracted with toluene, taken to dryness and resuspended in 62 63 64 10 65 acetonitrile. PAs were separated and quantified by HPLC using a reverse phase C18 column 1 2 (Spherisorb ODS2, 5-μm particle diameter, 4.6 x 250 mm, Waters, Wexford, Ireland) and a 3 4 -1 5 programmed acetonitrile:water step gradient (flow rate 1 mL min ) on a Jasco system (Jasco Corp., 6 7 Tokyo, Japan) consisting of a PU-1580 pump, an LG-1580-02 ternary gradient unit, a DG-1580-53 8 9 10 three-line degasser and a FP-1529 fluorescence detector, linked to an autosampler (AS 2055 Plus). 11 12 13 14 Data analysis 15 16 17 Data were processed with Graphpad Prism 6 (Graph Pad Software, San Diego CA, USA). In each 18 19 20 case, means ± standard deviations for n number of samples are given. Statistical analyses for 21 22 comparison of means were carried out using ANOVA, followed by Student’s t-test (significance 23 24 25 level, 0.05). 26 27 28 29 30 31 32 Results 33 34 35 36 Growth and morphology of N. oleoabundans cultivated autotrophically and mixotrophically in a 37 38 20-L PBR 39 40 Growth kinetics and cell morphology of N. oleoabundans cultivated autotrophically and 41 42 43 mixotrophically in a 20-L PBR are shown in Fig 1. Autotrophic cells rapidly entered the 44 45 exponential phase starting on the 2nd-3rd day up to the 7th-9th day of cultivation (Fig 1a). After this 46 47 48 period, cultures had a short late exponential phase and then entered the stationary phase of growth, 49 50 reaching a final cell density of about 22 × 106 cells mL-1 at the end of the of cultivation time (25 51 52 53 days). During the entire period, cells showed the typical morphology of N. oleoabundans grown in 54 55 BM medium, i.e., almost spherical, with a cell diameter of 3-5 µm (Fig 1b, d). One cup-shaped 56 57 chloroplast, containing a large pyrenoid, was present and emitted an intense red fluorescence due to 58 59 60 the presence of chlorophyll (Fig 1b,c). On the 25th day of cultivation, cells maintained their normal 61 62 63 64 11 65 features and dimensions, but some lipid globules accumulated inside the cytoplasm, as revealed by 1 2 Nile Red staining (Fig 1d,e). 3 4 5 During the first four days of mixotrophic growth in the PBR in the presence of glucose, cells 6 7 showed an evident increase in cell density, reaching values of ca. 33 x 106 cells mL-1 (Fig 1f). 8 9 10 Subsequently, cells entered the stationary phase of growth, and then cell density leveled off to 11 12 values of about 36 x 106 cells mL-1 up to the end of the cultivation period, i.e., eight days. At the 13 14 beginning of cultivation, both cell shape and size, and chloroplast features in mixotrophic cultures 15 16 17 (Fig 1g, h) were similar to those of autotrophic cultures (Fig 1b, c). Starting on the 3rd day and up to 18 19 the end of cultivation, however, mixotrophic cells showed peculiar features. In fact, the chloroplast 20 21 22 lost its characteristic cup shape and translucent globules, which tended to occupy almost the entire 23 24 cell volume, accumulated inside the cytoplasm (Fig 1i). Nile Red staining confirmed the lipidic 25 26 27 nature of these droplets (Fig 1j). 28 29 30 31

32 Extracellular FFA composition in E and EG media 33 34 The extracellular FFA composition in the exhausted autotrophic (E) and mixotrophic (EG) media of 35 36 N. oleoabundans is shown in Table 1. While EG medium comprised both saturated and mono- 37 38 39 unsaturated FFAs, E medium contained only saturated FFAs. In fact, both samples contained 40 41 similar percentages of saturated myristic acid (C14:0) and palmitic acid (C16:0), while the 42 43 44 percentage of stearic acid (C18:0) was different between the two samples. In E medium, stearic acid 45 46 was present at about 64%, while in EG medium at about 56%; however, C18:0 represented the main 47 48 49 saturated FFA for both samples. It is noteworthy that only EG medium contained about 6% of the 50 51 monoenoic oleic acid (C18:1ω9). 52 53 54 55 56 Polyamine content in E and EG media and in the corresponding N. oleoabundans cells cultivated in 57 58 a 20-L PBR 59 60 61 As shown in Fig 9a, the PA composition of N. oleoabundans cells cultivated mixotrophically in the 62 63 64 12 65 PBR were characterized by higher putrescine, spermine, and, especially, spermidine levels 1 2 compared to cells cultivated autotrophically. In fact, Spm, Put and Spd concentrations in 3 4 5 mixotrophic cells were ca. 2, 30 and 57 times higher, respectively, than those found in autotrophic 6 7 cells (p<0.001). Conversely, the PA composition in E and EG medium was not significantly 8 9 10 different. In fact, both media contained similar amounts of Put and Spm. Interestingly, spermidine 11 12 was present in high concentration in both media (119 ± 16.91 and 101 ± 5.00 pmol mL-1 in E and 13 14 EG, respectively; Fig 9b). 15 16 17 18 19 Growth kinetics of N. oleoabundans in exhausted and control growth media 20 21 22 As shown in Fig 2a, the growth of N. oleoabundans in E+ medium was promoted for the entire 23 24 experiment relative to controls grown on BM medium; on the contrary, growth in EG+ medium was 25 26 27 similar to that of the C. All samples entered the exponential phase very soon (during the first 3 days 28 29 of cultivation), despite some differences in growth rates (Fig 2b). The highest μ values were 30 31 -1 32 observed for algae cultivated in the two exhausted media (E+ and EG+, 1.29 and 1.18 div d , 33 34 respectively; control, 0.98 div d-1; p <0.01 in both cases relative to C) (Fig 2b). Subsequently, from 35 36 the 3rd day onwards, no relevant differences were observed between C and EG+ samples, as both of 37 38 39 them showed similar growth kinetics and cell densities (Fig 2a). Conversely, E+ samples reached 40 41 and maintained the highest cell density; the major difference as compared to the other samples was 42 43 th 6 -1 44 observed starting on the 7 day, with a cell density of about 16 × 10 cells mL (45 and 50% more 45 46 than EG+ and C samples, respectively; p <0.01 in both cases relative to E+ samples) at the same 47 48 th 49 time point. After the 17 day, all samples entered the stationary phase, reaching, at the end of the 50 51 experiment, a final cell density of 25-30 × 106 cells mL-1. During the experiments, aliquots of 52 53 samples were periodically harvested to measure the pH of the culture media. As observed in Figure 54 55 56 S1, initial pH was very different between control medium (about 7.0) and exhausted media (about 57 58 + + 9.5 in both cases). pH gradually increased up to 9.2 in the control medium, whilst in E and EG 59 60 61 media the pH varied between 9.5 and 8.0 without any obvious trend. 62 63 64 13 65

1 2 Consumption of nitrate and phosphate by N. oleoabundans cultivated in exhausted and control 3 4 5 growth media 6 7 The consumption of nitrate and phosphate by N. oleoabundans cells in the course of the experiment 8 9 10 is depicted in Fig 3. At time 0 (inoculation), the exhausted replenished growth media contained 11 - 2- 12 approximately 2 mM NO3 and 0.115 mM PO4 , i.e., the typical concentration of those components 13 14 in BM medium. By the end of the experiment, cells had consumed practically all nitrate and 15 16 17 phosphate present in the growth media. However, nutrient consumption by C, E+ and EG+ algal 18 19 cultures followed different trends, since C samples maintained higher nitrate and phosphate 20 21 22 concentrations than samples in exhausted media. In fact, after 12 days of cultivation, cells grown in 23 24 E+ and EG+ had consumed about 34% and 68% of nitrate, respectively, while C only 15%. Nitrate 25 26 27 concentration decreased from 1.43 mM at time 0 for all culture media to 0.94 mM for E+, 0.45 mM 28 29 for EG+ and 1.21 mM for C media (p <0.001 in both treated samples relative to C; Fig 3a). 30 31 32 Differently, during the first 12 days of growth, phosphate exhibited a dramatic decrease, with a 33 34 reduction by cells cultivated in EG+ and E+ of 94% and 98%, respectively, while in C medium it 35 36 was only about 60%. In fact, phosphate concentration decreased from about 0.086 mM at time 0 for 37 38 39 all culture media to 0.002 for E+, 0.005 for EG+ and 0.035 mM for C media (p <0.001 in both 40 41 treated samples relative to C; Fig 3b). 42 43 44 45 46 Photosynthetic pigment content in N. oleoabundans cultivated in exhausted and control media 47 48 49 During the experiment a gradual increase in photosynthetic pigment content was observed in all 50 51 samples (Fig 4). Chla in algae grown in C medium showed an evident increase up to the end of the 52 53 experiment, reaching an average concentration of 0.5 nmol 10-6 cells (Fig 4a). A similar increasing 54 55 56 trend was also observed in E+ samples, though with values 10-15% lower than those of controls. 57 58 From the 17th day of cultivation, Chla content of E+ samples was significantly lower than in C 59 60 61 (32%; p<0.01). Conversely, in EG+ samples Chla content showed a slightly increasing trend as 62 63 64 14 65 compared with C and E+ samples; this led to a final value of 0.35 nmol 10-6 cells at the end of the 1 2 experiment (32 and 21% lower than C and E+ samples, respectively; Fig 4a). An increasing trend in 3 4 5 Chlb content was also observed for all samples, without significant differences between C and E+ 6 7 samples (Fig 4b). Similar to Chla content, EG+ cells contained lower quantities of Chlb than C and 8 9 10 E+ samples (15-25% and 20-35% lower than C and E+ samples, respectively). More evident 11 12 differences were observed for Car content (Fig 4c). In fact, C samples showed an evident increase 13 14 up to the end of the experiment, always containing higher quantities compared to the other samples 15 16 17 (20-45% and 15-30% more than E+ and EG+ samples, respectively; p<0.05). It is noteworthy that 18 19 EG+ and E+ cells shared a similar trend of Car concentration during the experiment (Fig 4c). 20 21 22 23 24 Maximum quantum yield of PSII of N. oleoabundans cultivated in exhausted and control media 25 26 27 The variations in PSII maximum quantum yield measured during the experiments are shown in Fig 28 29 5. C, E+ and EG+ samples showed a slight increase of the FV/FM ratio during the first three days of 30 31 32 growth, reaching values of ca. 0.70. Subsequently, C cells maintained stable values around 0.70- 33 34 0.75, while the FV/FM ratio of E+ and EG+ samples decreased drastically down to values below 35 36 0.50 at the end of experiment. In EG+ cells the decrease was dramatic already from the 3rd day of 37 38 39 cultivation, reaching the lowest value after 17 days (ca. 0.30 for EG+ vs 0.75 for C and 0.50 for 40 41 E+; p <0.01 in both cases); thereafter, samples maintained stable values of ca. 0.35 until the end of 42 43 44 the experiment. On the contrary, in E+ cells, FV/FM started to decrease very strongly only from the 45 46 7th day of cultivation until the 17th day (from about 0.70 at day 7 down to 0.50 at day 17). During 47 48 th th 49 the subsequent experimental times (21 and 25 day), the FV/FM ratio of E+ cells remained stable 50 51 around 0.45. 52 53 54 55 56 Morphological observations of N. oleoabundans cultivated in exhausted and control media 57 58 59 Light microscopy and Nile Red staining 60 61 Light microscopy of both control and treated samples showed that N. oleoabundans maintained 62 63 64 15 65 similar cell morphology and dimensions throughout the experiment (Fig 6, 7). In fact, cells were 1 2 almost spherical with a cell diameter of 3-5 μm. One cup-shaped chloroplast, containing a large 3 4 5 pyrenoid, was present inside the cells (Fig 6a, c, e). Moreover, the chloroplast emitted an intense 6 7 red fluorescence due to Chl (Fig 6b, d, f). Interestingly, after 25 days of cultivation, while cell size 8 9 10 and shape remained substantially unchanged, all algal samples showed some translucent granulation 11 12 at the cytoplasmic level (Fig 7a, c, e). In order to understand the nature of those granulations and to 13 14 investigate if the recycled growth media could promote the production of lipid globules, all samples 15 16 17 were periodically stained with the lipid-specific fluorochrome Nile Red. The reaction was positive 18 19 only at the end of the experiment and the translucent globules were then unequivocally identified as 20 21 22 lipid droplets (Fig 7b, d, f). 23 24 25 26 27 TEM observations 28 29 To investigate the morphological and cytological changes induced by cultivation in recycled growth 30 31 32 media, the ultrastructure of C, E+ and EG+ samples was observed by TEM. At 12 days of 33 34 cultivation, most of the C cell volume was occupied by a characteristic cup-shaped chloroplast. 35 36 Inside the plastid one large pyrenoid, which was crossed by two elongated and appressed 37 38 39 thylakoids, was present (Fig 8a). In particular, the organelle contained starch in the shape of a shell 40 41 around the pyrenoid, and showed the typical thylakoid organization (Fig 8b). Typically featured 42 43 44 chloroplasts, as described for cells grown in C, were observed in cells grown in E+ medium (Fig 45 46 8c); however, thylakoids were more appressed than in C (Fig 8d). EG+ samples showed a more 47 48 49 strongly altered chloroplast structure (Fig 8e), as compared with C and E+ samples. Photosynthetic 50 51 membranes showed different degrees of thylakoid appression: some portions of thylakoid 52 53 membranes were appressed while others were loose and sometimes swollen (Fig 8f,g). In addition, 54 55 56 large portions of the stroma were free of thylakoids and some plastoglobules in proximity of the 57 58 thylakoid membranes were also visible (Fig 8g, h). Finally, in EG+ samples the pyrenoid lost its 59 60 61 round shape and appeared malformed (Fig 8e). 62 63 64 16 65

1 2 3 Discussion 4 5 6 7 The recycling of culture medium has been proposed as a possible solution in order to reduce water 8 9 consumption for algal cultivation, thereby making the process more economically feasible and 10 11 12 environmentally sustainable (Yang et al. 2011; Hadj-Romdhane et al. 2013; Farooq et al. 2014). 13 14 Yang et al. (2011) estimated that the large-scale cultivation of the microalga Chlorella vulgaris in 15 16 17 recycled culture medium could reduce water use by about 84%. Present results clearly suggest that 18 19 N. oleoabundans can also efficiently grow in exhausted growth media, especially in the autotrophic 20 21 medium replenished with nitrate and phosphate (E+). Observing the consumption trend of these 22 23 24 main nutrients (Fig. 3), it is clear that in the exhausted media their concentration decreased faster 25 26 with respect to controls in parallel to the enhanced cell growth. Moreover, phosphate was consumed 27 28 29 faster than nitrate by all samples, probably because of its 100 times lower concentration with 30 31 respect to the latter nutrient (0.2 gL-1 of nitrate vs 0.002 gL-1 of phosphate; Baldisserotto et al., 32 33 34 2012).Thus, in order to make the use of recycled culture medium feasible, exhausted media should 35 36 be replenished with the correct ratio of nitrate and phosphate, as they are the main nutrients that 37 38 39 guarantee cell growth (Stephens et al. 2010). This aspect was preliminarily verified by employing 40 41 unmodified exhausted growth media (E and EG) to re-cultivate N. oleoabundans. In fact, cells 42 43 showed a slight growth, reaching, at the end of the cultivation period, low cell densities (ca. 11 vs 44 45 6 -1 46 32 × 10 cells mL of C cultures) (data not shown). These cells accumulated intracellular lipids 47 48 throughout the experiment (data not shown) as a consequence of the limitation of nitrate and 49 50 51 phosphate (Mata et al. 2010; Popovich et al. 2012). These results are consistent with those of Zhu et 52 53 al. (2013), who found that C. zofingensis cultivated in recycled medium, without nitrogen and 54 55 56 phosphorus, displayed enhanced lipid production compared to cultures with full nutrients. 57 58 In order to understand if the growth promotion of N. oleoabundans cultivated in E+ and EG+, 59 60 relative to C, was influenced by some molecules released from the algae, a characterization of 61 62 63 64 17 65 autotrophic (E) and mixotrophic (EG) media was performed. First of all, pH of the control and 1 2 exhausted media was monitored. In particular, an increasing trend was observed in control, which 3 4 5 reached the pH of the exhausted media at the end of the experiment. This was not surprisingly, 6 7 considering that during the microalgal growth pH usually increases because of the consumption of 8 9 + 10 CO2 during the photosynthetic process (Zhang et al., 2014). However, considering that in E and 11 12 EG+ pH fluctuated around their initial values, or at least gradually decreased, it is not possible to 13 14 attribute to pH any evident role in the cell growth promotion. Moheimani and Borowitzka (2006) 15 16 17 and Stephen et al. (2010) reported that molecules released from cells can positively/negatively alter 18 19 cell metabolism and biomass production. 20 21 22 Here we show that both exhausted media contained the main PAs (putrescine, spermidine, and 23 24 spermine). It is known that these plant growth regulators are involved in a great variety of 25 26 27 developmental processes in plant cells, e.g., cell division and protein synthesis (Kaur-Sawhney et 28 29 al. 2003; Kuznetsova et al. 2006). PA biosynthetic pathways have also been studied in green algae 30 31 32 (Cohen et al. 1984; Theiss et al. 2002; Fuell et al. 2010). Although algae produce “unusual” PAs, 33 34 such as homospermidine and thermospermine (Hamana et al. 2013), the more common PAs (Put, 35 36 Spd and Spm), when added to the media, promoted growth and metabolism in C. vulgaris (Czerpak 37 38 39 et al. 2003). PAs are also known to alleviate the effect of biotic and abiotic stress in plants as well 40 41 as algae (Tate et al. 2013). 42 43 44 In the present work, a very strong increase in Put and, especially, Spd concentrations was observed 45 46 in N. oleoabundans mixotrophic cells as compared to the autotrophic ones, both cultivated in a 20-L 47 48 49 PBR. The growth-promoting role of PAs could explain the faster (0-4 days) and stronger cell 50 51 density increase (ca. 33 x 106 cell mL-1) in mixotrophic cultures relative to the autotrophic ones (0-9 52 53 days; ca. 22 x 106 cell mL-1). The higher PA concentrations in mixotrophically grown cells also 54 55 56 suggest that, under these culture conditions, algal cells may be better protected from stress-inducing 57 58 factors, e.g., bacterial contamination. To our knowledge, the presence of PAs released into algal 59 60 61 growth medium in PBRs has not been documented before. The release of PAs (predominantly Spd) 62 63 64 18 65 from autotrophic and mixotrophic cells into their respective (E and EG) exhausted media was not 1 2 significantly different. However, the presence of these plant growth regulators seems to contribute 3 4 5 to the promotion of N. oleoabundans growth in both media replenished with nitrate and phosphate. 6 7 In fact, during the entire experiment, a higher cell density was observed in E+ relative to C medium. 8 9 10 Moreover, the EG+ samples showed a promotion of cell density, albeit similar to that of C. 11 12 Both autotrophic and mixotrophic exhausted media also contained FFAs, which could have induced 13 14 a change in the normal metabolism of N. oleoabundans. This assumption is demonstrated by the 15 16 17 more rapid consumption of nitrate and phosphate, the decreased photosynthetic pigments content 18 19 and the strong decline in the FV/FM ratio observed in cells grown in E+ and EG+ media. These 20 21 22 results corroborate the assumption that microalgae release metabolites in high cell density cultures 23 24 (Richmond 2004) or as a consequence of stressful conditions (Ikawa 2004; Wu et al. 2006). Even if 25 26 27 the role of FFAs is currently under debate (Ikawa 2004; Stephen et al. 2010), recent studies showed 28 29 that they can strongly inhibit growth or exert cytotoxic effects on microalgae (Kogteva and 30 31 32 Bezuglov 1998; Wu et al. 2006; Bosma et al. 2008; Stephen et al. 2010). In fact, the presence of 33 34 these metabolites, and their oxidative products, negatively affect biomass productivity, especially 35 36 when the microalgae are cultivated in recycled medium (Lívanský et al. 1996; Rodolfi et al. 2003). 37 38 39 Granum et al. (2002) reported that the accumulation of intracellular lipids in microalgae enhanced 40 41 the release of FFAs into the culture medium, and Harun et al. (2010) observed that this release was 42 43 44 caused by cell lysis. 45 46 In this work, FFA composition in autotrophic (E) and mixotrophic (EG) media after N. 47 48 49 oleoabundans cultivation was similar. The only main difference was represented by the presence of 50 51 the monoenoic oleic acid (C18:1ω9) in EG medium. Oleic acid is one of the major lipid 52 53 components in lipid-enriched N. oleoabundans grown under N-stress conditions (Popovich et al. 54 55 56 2012). It is reported that mixotrophic cultivation alters N:C ratio, inducing similar lipid production 57 58 as N-depleted autotrophic cultures (Giovanardi et al. 2014). Probably for this reason, the release of 59 60 61 oleic acid occurred only in the mixotrophic growth medium. Moreover, Wu et al. (2006) observed 62 63 64 19 65 altered plasma membrane permeability due to the toxic effects of FFAs in two Chlorophyta (C. 1 2 vulgaris and Monoraphidium contortum) and in a cyanobacterium (Anabaena P-9). Then, in 3 4 5 addition to the lower concentration with respect to nitrate in BM medium, the rapid consumption of 6 7 phosphate in both exhausted media could be linked also to an alteration in membrane permeability, 8 9 10 due to the activity of FFAs (Ikawa 2004; Wu et al. 2006). In addition, FFAs can cause inhibition of 11 12 the PSII and PSI electron transport chains (Siegenthaler 1973) and disorganization of thylakoids 13 14 (Wu et al. 2006). Indeed, N. oleoabundans cells in E+ and EG+ were characterized by alterations of 15 16 17 the photosynthetic apparatus. These cells, in fact, contained less photosynthetic pigments than C 18 19 cells, and exhibited a drastic decrease of the PSII maximum quantum yield (FV/FM). These 20 21 22 variations are linked to an alteration of photosynthetic efficiency, especially as regards PSII (White 23 24 et al. 2011). The decreased photosynthetic pigment contents, observed in cells grown in E+ and 25 26 27 EG+, also reflected the changes in thylakoid membrane arrangement (Baldisserotto et al. 2012). 28 29 Therefore, correct thylakoid organization is necessary to maintain optimal photosynthetic activity 30 31 32 and this is often influenced by culture conditions (Nevo et al. 2012). In addition, the presence of 33 34 some plastoglobules in chloroplasts of algal cells, grown on exhausted media, could be another 35 36 indicator of an alteration of photosynthetic membranes (Besagni and Kessler 2013). On the 37 38 39 contrary, N. oleoabundans cells grown in C medium showed stable and normal FV/FM values 40 41 throughout the experiment (Giovanardi et al. 2014), and the typical assembly of thylakoid 42 43 44 membranes (Nevo et al. 2012; Baldisserotto et al. 2012; Giovanardi et al. 2014). 45 46 The presence of lipid droplets, observed only at the end of the experiment in C, E+ and EG+ cells, 47 48 49 is probably related to aging of the microalgal cultures (Hu et al. 2008; Baldisserotto et al. 2012), 50 51 rather than to the use of recycled culture media. 52 53 54 55 56 In conclusion, results presented here demonstrate that recycling autotrophic and mixotrophic 57 58 growth media is a suitable solution to obtain high cell density cultures of the microalga N. 59 60 61 oleoabundans. For this reason, they represent a contribution for improving the scale-up of 62 63 64 20 65 microalgal cultivation while providing a more sustainable ecological impact on water resources. 1 2 However, further studies are needed to deepen knowledge on the nature and the specific role of 3 4 5 some molecules that are released into the growth media, in order to obtain useful information for 6 7 the advancement of the biotechnological use of this strain. 8 9 10 11 12 Acknowledgements 13 14 This work was financially supported by grants from the University of Ferrara. We are grateful to 15 16 17 Dr. Immacolata Maresca of the University of Ferrara for technical assistance in freeze-drying of 18 19 samples. 20 21 22 23 24 Conflict of interest 25 26 27 The authors declare they have no conflict of interest. 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 21 65 References: 1 2 Baldisserotto C, Ferroni L, Giovanardi M, Pantaleoni L, Boccaletti L, Pancaldi S (2012) Salinity 3 4 5 promotes growth of freshwater Neochloris oleoabundans UTEX 1185 (Sphaeropleales, 6 7 Neochloridaceae): morpho-physiological aspects. Phycologia 51:700-710 8 9 10 Baldisserotto C, Giovanardi M, Ferroni L, Pancaldi S (2014) Growth, morphology and 11 12 photosynthetic responses of Neochloris oleoabundans during cultivation in a mixotrophic 13 14 brackish medium and subsequent starvation. Acta Physiol Plant 36:461-472 15 16 17 Batan L, Quinn CJ, Bradley TH (2013) Analysis of water footprint of a photobioreactor microalgae 18 19 biofuel production system from blue, green and lifecycle perspectives. Algal Research 2:196- 20 21 22 203 23 24 Besagni C and Kessler F (2013) A mechanism implicating plastoglobules in thylakoid disassembly 25 26 27 during senescence and nitrogen starvation. Planta 237:463-470 28 29 Borowitzka MA (2005) Culturing microalgae in outdoor ponds. In: Anderson RA (ed) Algal 30 31 32 culturing techniques. Elsevier Academic, London, pp 205-218 33 34 Borowitzka MA and Moheimani NR (2013) Sustainable biofuels from algae. Mitig Adapt Strateg 35 36 Glob Change 18:13-25 37 38 39 Bosma R, Miazek K, Willemsen SM, Vermuë MH, Wijffels RH (2008) Growth inhibition of 40 41 Monodus subterraneus by free fatty acids. Biotechnol Bioeng 101:1108–1114 42 43 44 Chisti Y (2007) Biodiesel from microalgae. Biotech Adv 25:294-306 45 46 Czerpak R, Bajguz A, Pietrowska A, Dobrogowska R, Matejczyk M, Weislawski W (2003) 47 48 49 Biochemical activity of di- and polyamines in the green alga Chlorella vulgaris Beijerinck 50 51 (Chlorophyceae). Acta Soc Bot Pol 72:19–24 52 53 Cohen E, Arad SM, Heimer YH, Mizrahi Y (1984) Polyamine biosynthetic enzymes in the cell cycle 54 55 56 of Chlorella. Plant Physiol 74:385-388 57 58 59 Farooq W, Suh WI, Park MS, Yang J (2014) Water use and its recycling in microalgae cultivation for 60 61 biofuel application. Bioresource Technol doi: 10.1016/j.biortech.2014.10.140 62 63 64 22 65 Ferroni L, Baldisserotto C, Pantaleoni L, Billi P, Fasulo MP, Pancaldi S (2007) High salinity alters 1 2 chloroplast morpho-physiology in a freshwater Kichneriella species (Selenastraceae) from 3 4 5 Ethipian Lake Awasa. Am J Bot 94:1973-1983 6 7 Ferroni L, Baldisserotto C, Giovanardi M, Pantaleoni L, Morosinotto T, Pancaldi S (2011) Revised 8 9 10 assignment of room-temperature chlorophyll fluorescence emission bands in single living cells 11 12 of Chlamydomonas reinhardtii. J Bioenerg Biomembr 43:163-173 13 14 Fuell C, Elliott KA, Hanfrey CC, Franceschetti M, Michael AJ (2010) Polyamine biosynthetic 15 16 17 diversity in plants and algae. Plant Physiol Biochem 48:513–520 18 19 Giovanardi M, Ferroni L, Baldisserotto C, Tedeschi P, Maietti A, Pantaleoni L, Pancaldi S (2013) 20 21 22 Morpho-physiological analyses of Neochloris oleoabundans (Chlorophyta) grown 23 24 mixotrophically in a carbon-rich waste product. Protoplasma 250:161-174 25 26 27 Giovanardi M, Baldisserotto C, Ferroni L, Longoni P, Cella R, Pancaldi S (2014) Growth and lipid 28 29 synthesis promotion in mixotrophic Neochloris oleoabundans (Chlorophyta) cultivated with 30 31 32 glucose. Protoplasma 251:115-125 33 34 Gouveia L, Marques AE, Lopes da Silva T, Reis A (2009) Neochloris oleoabundans UTEX #1185: a 35 36 suitable renewable lipid source for biofuel production. J Ind Microbiol Biotechnol 36: 821-826 37 38 39 Granum E, Kirkvold S, Myklestad SM (2002) Cellular and extracellular production of carbohydrates 40 41 and amino acids by the marine diatom Skeletonema costatum: diel variations and effects of N 42 43 44 depletion. Mar Ecol Prog Ser 242:83–94. 45 46 Grobbelaar JU (2009) Factors governing algal growth in photobioreactors: the “open” versus 47 48 49 “closed” debate. J Appl Phycol 21:489-492 50 51 Hadj-Romdhane F, Jaouen P, Pruvost J, Grizeau D, Van Vooren G, Bourseau P (2012) Development 52 53 and validation of a minimal growth medium for recycling Chlorella vulgaris culture. 54 55 56 Bioresource Technol 123:366-374 57 58 Hadj-Romdhane F, Zheng X, Jaouen P, Pruvost J, Grizeau D, Croué JP, Bourseau P (2013) The 59 60 61 culture of Chlorella vulgaris in a recycled supernatant: Effects on biomass production and 62 63 64 23 65 medium quality. Bioresource Technol 132: 285–292 1 2 Hamana K, Niitsu M, Hayashi H (2013) Occurrence of homospermidine and thermospermine as a 3 4 5 cellular polyamine in unicellular chlorophyte and multicellular charophyte green algae. J Gen 6 7 Appl Microbiol 59: 313-319 8 9 10 Harun R, Singh M, Forde GM, Danquah MK (2010) Bioprocess engineering of microalgae to 11 12 produce a variety of consumer products. Renew Sust Energ Rev 14:1037-1047 13 14 Hu Q, Sommerfeld M, Jarvis E, Ghirardi M, Posewitz M, Seibert M, Darzins A (2008) Microalgal 15 16 17 triacylglycerols as a feedstocks for biofuel production: perspective and advances. Plant J 18 19 54:621-639 20 21 22 Ikawa M (2004) Algal polyunsaturated fatty acids and effects on plankton ecology and other 23 24 organism. UNH Center for Freshwater Biology Research 6:17-44 25 26 27 Kalaji HM, Schansker G, Ladle RJ, Vasilij Goltsev V, Bosa K, Allakhverdiev SI, Brestic M, Bussotti 28 29 F, Calatayud A, Dąbrowski P, Elsheery NI, Ferroni L, Guidi L, Hogewoning SW, Jajoo A, 30 31 32 Misra AN, Nebauer SG, Pancaldi S, Penella C, Poli D, Pollastrini M, Romanowska-Duda ZB, 33 34 Rutkowska B, Serôdio J, Suresh K, Szulc W, Tambussi E, Yanniccari M, Zivcak M (2014) 35 36 Frequently asked questions about in vivo chlorophyll fluorescence: practical issues. Photosynth 37 38 39 Res 122:121-158 40 41 Kaur-Sawhney R, Tiburcio AF, Altabella T, Galston AW (2003) Polyamines in plants: an overview. 42 43 44 J Cell Mol Biol 2:1-12 45 46 Kogteva GS and Bezuglov VV (1998) Unsaturated fatty acids as endogenous bioregulators. 47 48 49 Biochemistry (Moscow) 63:4-12 50 51 Lam MK and Lee KT (2012) Potential of using organic fertilizer to cultivate Chlorella vulgaris for 52 53 54 biodiesel production. Appl Energy 94:303-308 55 56 Li Y, Horsman M, Wang B, Wu N, Lan CQ (2008) Effect on nitrogen sources on cell growth and 57 58 lipid accumulation of green alga Neochloris oleoabundans. Appl Microbiol Biotechnol 81:629- 59 60 61 636 62 63 64 24 65 Lichtenthaler HK, Buschmann C, Knapp M (2005) How to correctly determine the different 1 2 chlorophyll fluorescence parameters and the chlorophyll fluorescence decrease ratio RFd of 3 4 5 leaves with the PAM fluorometer. Photosynthetica 43:379-393 6 7 Lívanský K, Dědič K, Bínová J, Tichý V, Novotný P, Doucha J (1996) Influence of the nutrient 8 9 10 solution recycling on the productivity of Scenedesmus obliquus, utilization of nutrients and 11 12 water in outdoor cultures. Algological Studies/ Archiv für Hydrobiologie 81:105–113 13 14 Mata TM, Martins AA, Caetano NS (2010) Microalgae for biodiesel production and other 15 16 17 applications: a review. Renew Sust Energ Rev 14:217- 232 18 19 Moheimani NR and Borowitzka MA (2006) The long-term culture of the coccolithophore 20 21 22 Pleurochrysis carterae (Haptophyta) in outdoor raceway ponds. J Appl Phycol (2006) 18:703- 23 24 712 25 26 27 Molina Grima E, Belarbi H, Acién Fernández FG, Robles Medina A, Chisti Y (2003) Recovery of 28 29 microalgal biomass and metabolites: process options and economics. Biotech Adv 20: 491-515 30 31 32 Ndimba BK, Ndimba RJ, Johnson TS, Waditee-Sirisattha R, Baba M, Sirisattha S, Shiraiwa Y, 33 34 Rakwal R, Agrawal GK, Rakwal R (2013) Biofuels as a sustainable energy source: An update 35 36 of the applications of proteomics in bioenergy crops and algae. J Prot 93:234-244 37 38 39 Nevo R, Charuni D, Tsabari O, Reich Z (2012) Composition, architecture and dynamics of the 40 41 photosynthetic apparatus in higher plants. Plant J 70:157-176 42 43 44 Pancaldi S, Baldisserotto C, Ferroni L, Bonora A, Fasulo MP (2002) Room temperature 45 46 microspectrofluorimetry as a useful tool for studying the assembly of the PSII chlorophyll– 47 48 49 protein complexes in single living cells of etiolated Euglena gracilis Klebs during the greening 50 51 process. J Exp Bot 53:1753–1763 52 53 Popovich CA, Damiani MC, Constenla D, Martínez AM, Giovanardi M, Pancaldi S, Leonardi PI 54 55 56 (2012) Neochloris oleoabundans grown in natural enriched seawater for biodiesel feedstock: 57 58 Evaluation of its growth and biochemical composition. Bioresource Technol 114:287-293 59 60 61 Pruvost J, Van Vooren G, Cogne G, Legrand J (2009) Investigation of biomass and lipids production 62 63 64 25 65 with Neochloris oleoabundans in photobioreactor. Bioresource Technol 100:5988-5995 1 2 Pulz O (2001) Photobioreactors: production systems for phototrophic microorganisms. Appl 3 4 5 Microbiol Biotechnol 57:287-293 6 7 Richmond A (2004) Handbook of Microalgal Culture: Biotechnology and Applied Phycology. 8 9 10 Blackwell Sciences Ltd., Oxford 11 12 Rodolfi L, Zittelli GC, Barsanti L, Rosati G, Tredici MR (2003) Growth medium recycling in 13 14 Nannochloropsis sp. mass cultivation. Biomol Eng 20:243–248 15 16 17 Scaramagli S, Biondi S, Capitani F, Gerola P, Altamura MM, Torrigiani P (1999) Polyamine 18 19 conjugate levels and ethylene biosynthesis: inverse relationship with vegetative bud formation 20 21 22 in tobacco thin layers. Physiol Plant 105:366-375 23 24 Sheehan JJ (2009) Biofuels and the conundrum of sustainability. Curr Opin Biotechnol 20:318–324 25 26 27 Siegenthaler PA (1973) Change in pH dependence and sequential inhibition of photosynthetic 28 29 activity in chloroplasts by unsaturated fatty acids. Biochim Biophys Acta 305:153-162 30 31 32 Smith VH, Sturm BSM, deNoyelles FJ, Billings SA (2010) The ecology of algal biodiesel 33 34 production. Trends Ecol Evo 25:301-309 35 36 Solomon BD (2010) Biofuels and sustainability. Ann NY Acad Sci 1185:119-134 37 38 39 Stephens E, Ross IL, Mussgnug JH,Wagner LD, BorowitzkaMA, Posten C, Kruse O, Hankamer B 40 41 (2010) Future prospects of microalgal biofuel production systems. Trends Plant Sci 15:554-564 42 43 44 Tate JJ, Gutierrez-Wing MT, Rush KA, Benton MG (2013) The Effects of Plant Growth Substances 45 46 and Mixed Cultures on Growth and Metabolite Production of Green Algae Chlorella sp.: A 47 48 49 Review. J Plant Growth Regul 32:417–428 50 51 Tornabene TG, Holzer G, Lien S, Burris N (1983) Lipid composition of the nitrogen starved green 52 53 alga Neochloris oleoabundans. Enzyme Microbiol Tech 5:435-440 54 55 56 Theiss C, Bohley P, Voigt J (2002) Regulation by polyamines of ornithine decarboxylase activity and 57 58 cell division in the unicellular green alga Chlamydomonas reinhardtii Plant Physiol 128:1470- 59 60 61 1479 62 63 64 26 65 Wellburn AR (1994) The spectral determination of chlorophylls a and b, as well as total carotenoids, 1 2 using various solvents with spectrophotometer of different resolution. J Plant Physiol 144:307– 3 4 5 313 6 7 White S, Anandraj A, Bux F (2011) PAM fluorometry as a tool to assess microalgal nutrient stress 8 9 10 and monitor cellular neutral lipids. Bioresource Technol 102:1675-1682 11 12 Wu JT, Chiang YR, Huang WY, Jane WN (2006) Cytotoxic effects of free fatty acids on 13 14 phytoplankton algae and cyanobacteria. Aquat Toxicol 80:338–345 15 16 17 Yang, J, Xu M, Zhang X, Hu Q, Sommerfeld M, Chen Y (2011) Lifecycle analysis on biodiesel 18 19 production from microalgae: Water footprint and nutrients balance. Bioresource Technol 20 21 22 102:159-165 23 24 Zhang Q, Wang T, Hong Y (2014) Investigation of initial pH effects on growth of an oleaginous 25 26 27 microalgae Chlorella sp. HQ for lipid and nutrient uptake. Water Sci Technol 70:712-719 28 29 Zhu LD, Takala J, Hiltunen E, Wang ZM (2013) Recycling harvest water to cultivate Chlorella 30 31 32 zofingiensis under nutrient limitation for biodiesel production. Bioresource Technol 144:14-20 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 27 65 Figure captions: 1 2 Fig 1 (a-e) Growth kinetics (a) and morphology (b-e) of N. oleoabundans cells autotrophically 3 4 5 cultivated in a 20-L PBR in BM medium. Light (b, d) and fluorescence microscope observations (c, 6 7 e) of N. oleoabundans cells after 3 (b, c) and 25 (d, e) days of autotrophic cultivation. (f-j) Growth 8 9 10 kinetics (f) and cell morphology (g-j) of N. oleoabundans cultivated in a 20-L PBR in BM medium 11 12 under mixotrophic conditions (supplemented with 2.5 gL-1 of glucose). Light (g, i) and fluorescence 13 14 microscope observations (h, j) of N. oleoabundans cells after 3 (g, h) and 8 (d, e) days of 15 16 17 mixotrophic cultivation. In both graphs, curves are constructed on a log2 scale and data are means ± 18 19 s.d. (n=3). In all micrographs, bars: 2 μm 20 21 22 23 24 Fig 2 (a) Growth kinetics of N. oleoabundans in BM medium (filled circles), E+ medium (empty 25 26 27 squares) and EG+ medium (empty diamonds). (b) Growth rates, calculated during the exponential phase 28 29 (0–3 days time interval of cultivation), of cells grown in BM (white), E+ (light grey) and EG+ (dark 30 31 32 grey) media. The growth curve is constructed on a log2 scale and data are means ± s.d. (n=3) 33 34 35 36 Fig 3 Time-course variations in the concentrations of nitrate (a) and of phosphate (b) in culture media of 37 38 39 N. oleoabundans during 25 days of cultivation on BM medium (white), E+ medium (light grey) and 40 41 EG+ medium (dark grey). Values are means ± s.d. (n=3) 42 43 44 45 46 Fig 4 Time-course variations of Chla (a), Chlb (b) and Car content (c) in N. oleoabundans cells grown 47 48 49 in BM (filled circles), E+ (empty squares) and EG+ (empty diamonds) media during the 25 days of 50 51 cultivation. Values are means ± s.d. (n=3) 52 53 54 55 56 Fig 5 Time-course variations of PSII maximum quantum yield (FV/FM ratio) in N. oleoabundans cells 57 58 grown in BM (filled circles), E+ (empty squares) and EG+ (empty diamonds) media during the 25 days 59 60 61 of cultivation. Values are means ± s.d. (n=3) 62 63 64 28 65 Fig 6 Light and fluorescence microscopy observations of N. oleoabundans cells at 12 days of 1 2 cultivation. (a) Control cells and (b) the relative fluorescence of the chloroplast; (c) cell grown in 3 4 5 E+ medium and (d) the relative fluorescence of the chloroplast; (e) cell grown in EG+ medium and 6 7 (f) the relative fluorescence of the chloroplast. In all micrographs, bars: 2 μm 8 9 10 11 12 Fig 7 Light and epifluorescence pictures of N. oleoabundans cells after 25 days of growth. (a) 13 14 Control cells and (b) relative Nile Red-staining observation, (c) cells grown in E+ medium and (d) 15 16 17 relative Nile Red-staining observation, and (e) cells grown in E+ medium and (f) relative Nile Red- 18 19 staining observation in EG+ medium. Translucent globules are indicated with arrows. In all 20 21 22 micrographs, bars: 2 μm 23 24 25 26 27 Fig 8 Transmission electron micrographs of N. oleoabundans cells at 12 days of cultivation. (a) Cell 28 29 grown in BM medium and (b) detail of its chloroplast showing the typical thylakoid membranes 30 31 32 organization (white arrow). (c) Cell grown in E+ medium and (d) detail of its chloroplast with quite 33 34 compact and appressed (white arrow) thylakoids. (e) Cell grown in EG+ medium and (f-h) details 35 36 of its chloroplast. The presence of intermediate stages of thylakoid membranes are evident in EG+ 37 38 39 cells, showing a general disorganisation of thylakoid membranes (f-h), which appeared wavy, loose 40 41 and sometimes swollen (f, g). (h) Some plastoglobules in proximity of thylakoid membranes are 42 43 44 also observed (arrowheads). P, pyrenoid; N, nucleus, M, mithochondrion. Bars: 0.5 μm (a, c, e); 45 46 0.05 μm (b, d, f, g, h) 47 48 49 50 51 Fig 9 Polyamine concentrations (a) in N. oleoabundans cells cultivated in a 20-L PBR 52 53 autotrophically and mixotrophically at 15 and 8 days of cultivation, respectively, and (b) in the 54 55 56 corresponding exhausted E and EG growth media. Autotrophic cells (white), mixotrophic cells (dark), 57 58 E+ (light grey) and EG+ medium (dark grey). Values are means ± s.d. (n=3) 59 60 61 62 63 64 29 65 Manuscript Click here to download Manuscript: Sabia et al. MS.doc Click here to view linked References

1 2 Re-cultivation of Neochloris oleoabundans in exhausted autotrophic and mixotrophic media: 3 4 5 the potential role of polyamines and free fatty acids 6 7 8 9 1 1 2 1 10 Alessandra Sabia , Costanza Baldisserotto , Stefania Biondi , Roberta Marchesini , Paola 11 12 Tedeschi3, Annalisa Maietti3, Martina Giovanardi1, Lorenzo Ferroni1, Simonetta Pancaldi1 13 14 1Department of Life Sciences and Biotechnology, University of Ferrara, Corso Ercole I d'Este 32, 15 16 17 44121 Ferrara, Italy. 18 19 2Department of Biological, Geological and Environmental Sciences, University of Bologna, via 20 21 22 Irnerio 42, 40126 Bologna, Italy. 23 24 3Department of Chemical and Pharmaceutical Science, University of Ferrara, via Fossato di 25 26 27 Mortara 17, 44121 Ferrara, Italy. 28 29 30 31 32 *Corresponding Author. Tel: +39 0532 293786; Fax: +39 0532 208561; E-mail address: 33 34 [email protected]. 35 36 37 38 39 40 41 ABSTRACT 42 43 44 45 Neochloris oleoabundans (Chlorophyta) is widely considered one of the most promising microalgae 46 47 for biotechnological applications. However, the large-scale production of microalgae requires large 48 49 50 amounts of water. In this perspective, the possibility of using exhausted growth media for the re- 51 52 cultivation of N. oleoabundans was investigated in order to simultaneously make the cultivation 53 54 55 more economically feasible and environmentally sustainable. Experiments were performed by 56 57 testing the following media: autotrophic exhausted medium (E+) and mixotrophic exhausted 58 59 medium after cultivation with glucose (EG+) of N. oleoabundans cells grown in a 20-L 60 61 62 63 64 1 65 photobioreactor (PBR). Both exhausted media were replenished with the same amounts of nitrate 1 2 and phosphate as the control brackish medium (C). Growth kinetics, nitrate and phosphate 3 4 5 consumption, photosynthetic pigments content, photosynthetic efficiency, cell morphology, and 6 7 lipid production were evaluated. Moreover, the free fatty acid (FFA) composition of exhausted 8 9 10 media and the polyamine (PA) concentrations of both algae and media were analyzed in order to 11 12 test if some molecules, released into the medium, could influence algal growth and metabolism. 13 14 Results showed that N. oleoabundans can efficiently grow in both exhausted media (E+ and EG+), 15 16 17 especially in E+, and to the same extent as in C medium. Growth promotion of N. oleoabundans 18 19 was attributed to PAs and alteration of the photosynthetic apparatus to FFAs. Taken together, 20 21 22 results show that recycling growth medium is a suitable solution to obtain good N. oleoabundans 23 24 biomass concentrations, while providing a more sustainable ecological impact on water resources. 25 26 27 28 29 30 31 32 Keywords: 33 34 Neochloris oleoabundans 35 36 Recycling culture media 37 38 Photosynthetic apparatus 39 40 Biomass production 41 42 Polyamines 43 44 Free fatty acids 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 2 65 Introduction 1 2 3 Over the last decades, microalgae have received much attention as promising high-potential 4 5 6 feedstock for biodiesel (Chisti 2007; Li et al. 2008; Borowitzka and Moheimani 2013) and as an 7 8 attractive raw material for the production of a wide range of high value bioproducts (Molina Grima 9 10 11 et al. 2003; Mata et al. 2010). They are photosynthetic microorganisms with numerous key 12 13 features, which potentially make them a more appreciable renewable source than terrestrial plants 14 15 16 (Richmond 2004; Hu et al 2008; Ndimba et al. 2013). For instance, these microorganisms have 17 18 faster growth rates, higher photosynthetic efficiencies, higher rates of carbon dioxide fixation, and 19 20 higher biomass productivities compared to plants (Richmond 2004; Gouveia et al. 2009; Harun et 21 22 23 al. 2010; Mata et al. 2010). Moreover, microalgae can thrive in non-arable lands being able to grow 24 25 in waters having different salinity levels and chemical composition (Smith et al. 2010; Borowitzka 26 27 28 and Moheimani 2013). 29 30 The environmental and economical impact of large-scale production of microalgae has been widely 31 32 33 discussed, with the aim of making the process increasingly sustainable (Sheehan 2009; Solomon 34 35 2010). In this perspective, microalgal cultivation systems are crucial factors to be taken into account 36 37 (Smith et al. 2010; Stephens et al. 2010; Borowitzka and Moheimani 2013). A wide variety of 38 39 40 systems has been described by Mata and co-workers (2010) and recently by Borowitzka and 41 42 Moheimani (2013). Microalgae can be cultivated in open pond reactors or closed photobioreactors 43 44 45 (PBRs), depending on algal strain, issue of research, type of desired products and environmental 46 47 conditions (Chisti 2007; Mata et al. 2010; Borowitzka and Moheimani 2013). Although open ponds 48 49 50 are the most commonly used culture systems, due to their low cost of production and operation, 51 52 recently closed PBRs are receiving much attention for the possibility of producing valuable 53 54 55 compounds (Pulz 2001; Grobbelaar 2009; Smith et al. 2010). However, algal cultures require large 56 57 amounts of water in both culture systems (Lam and Lee 2012; Borowitzka and Moheimani 2013). 58 59 Considering that water is becoming a scarce natural resource, water demand represents an important 60 61 62 63 64 3 65 factor when performing large-scale cultivation (Batan et al. 2013; Farooq et al. 2014). In order to 1 2 reduce the high production costs and, at the same time, to make microalgal production more 3 4 5 environmentally sustainable, recycling culture medium has been proposed as a possible solution 6 7 (Yang et al. 2011; Hadj-Romdhane et al. 2012; Farooq et al. 2014), especially in large-scale culture 8 9 10 systems (Borowitzka 2005; Lam and Lee 2012; Zhu et al. 2013). However, the feasibility of 11 12 recycling growth medium to re-cultivate microalgae has been tested only in a few cases (Lívanský 13 14 et al. 1996; Rodolfi et al. 2003; Hadj-Romdhane et al. 2013; Zhu et al. 2013). It is reported that the 15 16 17 recycling medium could have negative effects on biomass productivity, due to the release of 18 19 inhibitory secondary metabolites (Richmond 2004). Harmful metabolites are, in fact, released 20 21 22 during microalgal growth under physiological stress (Ikawa 2004; Moheimani and Borowitzka 23 24 2006). Free fatty acids (FFAs), and substances derived from the photooxidation of unsaturated fatty 25 26 27 acids, are the most common metabolites with inhibitory effects on microalgae (Ikawa 2004; Wu et 28 29 al. 2006; Bosma et al. 2008; Stephen et al. 2010). 30 31 32 In the present work, the suitability of recycling growth media for the re-cultivation of Neochloris 33 34 oleoabundans (Chlorophyta) was investigated. This species was chosen because it is considered one 35 36 of the most promising oil-rich microalgae, due to its capability to accumulate lipids when grown 37 38 39 under nitrogen starvation (Tornabene et al. 1983; Chisti 2007; Li et al. 2008; Pruvost et al. 2009) or 40 41 mixotrophically, in the presence of glucose, or carbon-rich wastes, as organic carbon sources 42 43 44 (Giovanardi et al. 2013; Baldisserotto et al. 2014). Present results clearly indicate that N. 45 46 oleoabundans can efficiently grow in its exhausted growth medium, if suitably replenished with the 47 48 49 main nutrients. Based on this encouraging result, and with the aim of gaining further knowledge on 50 51 the morpho-physiological aspects and biotechnological applications of this algal strain, the 52 53 following features were analyzed: (1) growth kinetics in parallel to nitrate and phosphate 54 55 56 consumption; (2) photosynthetic pigment content; (3) PSII maximum quantum yield; (4) cell 57 58 morphology, with special attention to intracellular lipid accumulation, (5) FFA accumulation in 59 60 61 recycled growth media, and (6) polyamine (PA) concentration both inside cells and in the 62 63 64 4 65 corresponding culture media. PAs were determined in order to understand if these plant growth 1 2 regulators could be responsible for the growth promotion of N. oleoabundans in recycled growth 3 4 5 medium. It is known, in fact, that PAs, together with other plant growth regulators, have stimulatory 6 7 effects on algal growth and metabolism, and are involved in mitigating various types of biotic and 8 9 10 abiotic stress (Tate et al. 2013). 11 12 13 14 15 16 17 Materials and Methods 18 19 Algal strains and culture condition 20 21 22 The strain used in this study was the Chlorophyta N. oleoabundans UTEX 1185 (syn. Ettlia 23 24 25 oleoabundans) (Sphaeropleales, Neochloridaceae), obtained from the Culture Collection of Algae 26 27 of the University of Texas (UTEX, Austin, Texas, USA; www.utex.org). 28 29 30 N. oleoabundans was cultivated in axenic liquid brackish medium (BM) (Baldisserotto et al. 2012) 31 32 in a coaxial 20-L capacity PBR (M2M Engineering, Grazzanise, Caserta, Italy). Algae were 33 34 -1 35 cultivated autotrophically in BM for 25 days or mixotrophically, by addition of 2.5 g L glucose, 36 37 for 8 days, according to previously described protocols (Baldisserotto et al. 2014; Giovanardi et al. 38 39 2014). For autotrophic cultivation, cells were inoculated into the PBR to obtain an initial cell 40 41 6 -1 42 density of about 0.6 x 10 cells mL , while for mixotrophic cultivation initial cell density was 43 44 higher (10 x 106 cells mL-1). Culture conditions in the PBR were: 24 ± 1°C; sterile air injection at 45 46 -2 -1 47 the bottom of the PBR, with 0.5/3.5 h bubbling/static cycles; irradiance 65 µmolphotons m s of 48 49 PAR (16:8 h light:dark photoperiod). Light was supplied with inner cool–white fluorescent Philips 50 51 52 tubes. 53 54 Algal growth and morphology in the PBR were monitored as described below. 55 56 57 58 59 60 61 62 63 64 5 65 Preparation of growth media 1 2 3 Experiments were performed by testing the following media: 4 5 - C: freshly prepared BM medium (control); 6 7 8 - E: autotrophic exhausted medium; 9 10 - EG: mixotrophic exhausted medium after cultivation with glucose; 11 12 13 - E+: autotrophic exhausted medium replenished with nitrate and phosphate concentrations as for 14 15 BM medium; 16 17 - EG+: mixotrophic exhausted medium after cultivation with glucose and replenished with nutrients 18 19 20 as in E+. 21 22 In order to obtain the exhausted media for the recycling experiment, N. oleoabundans was grown 23 24 25 inside the PBR under the culture conditions described in the previous section. For E+ medium 26 27 preparation, about 500 mL of autotrophic algal culture in the stationary phase of growth (at the 15th 28 29 30 day of cultivation in the PBR) were centrifuged at 2,000 g for 10 min in order to separate the 31 32 medium from the algae, thus obtaining an autotrophic exhausted medium (E). The medium 33 34 35 appeared pale-yellow and was free from algae, bacteria and protozoa; its optical density at a 36 th 37 wavelength of 750 nm (OD750) was 0.02. For EG+ medium, at the 8 day of cultivation in the PBR, 38 39 the same aliquots of algal culture were harvested by centrifugation (2,000 g for 10 min). In this 40 41 42 case, the mixotrophic exhausted medium (EG) was straw-yellow in colour and presented a weak 43 44 bacterial contamination due to the glucose addition (OD750 =0.009). After determining the nitrate 45 46 47 and phosphate concentrations of the two exhausted media (see “Nitrate and phosphate analyses”), 48 -1 49 KNO3 and K2HPO4 were axenically added to reach final concentrations of 0.2 and 0.02 g L , 50 51 52 respectively, i.e., the typical concentration of those components in BM, thus obtaining the 53 54 replenished exhausted media (E+ and EG+) used for experiments. 55 56 57 58 59 60 61 62 63 64 6 65 Experimental design 1 2 3 When autotrophic cultures of N. oleoabundans, grown in BM in the PBR, reached a cell density of 4 5 10 x 106 cells mL-1 (after about 9 days of cultivation), aliquots of cells were inoculated into 300-mL 6 7 8 Erlenmeyer flasks (150 mL total volume) containing C, E+, EG+ media, to obtain an initial cell 9 10 density of about 0.5 x 106 cells mL-1. The cultures were placed in a growth chamber (24 ± 1 °C, 80 11 12 -2 -1 13 μmolphotons m s of PAR with a 16:8 h light-darkness photoperiod), and cultivated with continuous 14 15 shaking at 80 rpm, without external CO2 supply. Experiments were performed in triplicate. Aliquots 16 17 of cultures (cells and media) were collected at different times of cultivation up to 25 days, 18 19 20 depending on the analysis. 21 22 23 24 25 Analyses 26 27 28 Growth evaluation 29 30 Aliquots of cell samples cultivated in control and exhausted media were counted at 0, 3, 7, 12, 17 31 32 33 and 25 days of cultivation using a Thoma haemocytometer (HBG, Giessen, Germany) under a light 34 35 microscope (Zeiss, model Axiophot) and growth curves were obtained. 36 37 The growth rate (μ, number of divisions per day) during the exponential phase was calculated with 38 39 40 the following equation: 41 42 μ (div d-1) = (log N − log N ) / (t − t ), 43 2 1 2 0 1 0 44 45 where μ is the growth rate, N1 the cell number at time t1, N0 the cell number at time 0 and t1−t0 the 46 47 48 time interval (days) (Giovanardi et al. 2013). 49 50 51 52 Nitrate and phosphate analyses 53 54 55 After 12 and 25 days of cultivation, samples of C, E+ and EG+ media were harvested by 56 57 centrifugation to analyze nitrate and phosphate concentrations. These nutrients were quantified 58 59 60 colorimetrically using a flow-injection autoanalyzer (FlowSys, Systea, Roma, Italy). 61 62 63 64 7 65

1 2 Photosynthetic pigment extraction and quantification 3 4 5 For photosynthetic pigment analysis, cell samples were collected by centrifugation after 0, 7, 17, 21 6 7 and 25 days of cultivation. Extraction of photosynthetic pigments was performed according to 8 9 10 Baldisserotto et al. (2014). The extracts were measured with a Pharmacia Ultrospec 2000 UV-Vis 11 12 spectrophotometer (1-nm bandwidth; Amersham Biosciences, Piscataway, NJ, USA) at 666 13 14 (chlorophyll a - Chla), 653 (chlorophyll b - Chlb) and 470 nm (carotenoids - Car). Quantification 15 16 17 was performed according to equations reported in Wellburn (1994). 18 19 20 21 22 PAM fluorimetry 23 24 The PSII maximum quantum yield of algae was determined at the same cultivation times 25 26 27 considered for growth kinetics measurements. A pulse amplitude modulated fluorometer (ADC 28 29 Bioscientific Ltd, Hoddesdon, Hertfordshire, UK) was used to determine the in vivo chlorophyll 30 31 32 fluorescence of PSII. The PSII maximum quantum yield is reported as FV/FM ratio, i.e. (FM – F0)/ 33 34 FM, where variable fluorescence is FV = (FM−F0), FM is the maximum fluorescence and F0 is the 35 36 initial fluorescence of samples (Lichtenthaler et al. 2005). This measurement is considered a valid 37 38 39 method to probe the maximum quantum yield of photochemistry in PSII (Kalaji et al. 2014). 40 41 Moreover, it is useful to estimate the physiological state of plants and microorganisms also under 42 43 44 nutrient stress (White et al. 2011). Samples were prepared as reported in Ferroni et al. (2011) after 45 46 15 min of dark incubation. 47 48 49 50 51 Light and fluorescence microscopy 52 53 For microscopic observations, cell samples were routinely collected throughout the cultivation 54 55 56 period inside the PBR and during the experiment with exhausted media. Aliquots of samples were 57 58 observed using a microscope (Zeiss, model Axiophot) with conventional and fluorescent 59 60 61 attachments. The light source for chlorophyll fluorescence observation was a HBO 100 W pressure 62 63 64 8 65 mercury vapour lamp (filter set, BP436/10, LP470). Pictures of cells were taken with a Canon 1 2 IXUS 110 IS digital camera (12.1 megapixels), mounted on the ocular lens through a Leica DC150 3 4 5 system (Leica Camera AG, Solms, Germany). 6 7 8 9 10 Lipid staining 11 12 During experiments, the intracellular presence of lipids was evaluated by staining cells with the 13 14 fluorochrome Nile Red (9-diethylamina-5Hbenzo[α]phenoxazine-5-one, 0.5 mg dissolved in 100 15 16 17 mL acetone; Sigma-Aldrich, Gallarate, Milan, Italy), as described in Giovanardi et al. (2014). After 18 19 incubation at 37°C in darkness for 15 min, cells were observed with the microscope described 20 21 22 above at an excitation wavelength of 485 nm (filter set, BP485, LP520). Photographs were taken 23 24 with the camera described above. 25 26 27 28 29 Transmission electron microscopy (TEM) 30 31 th 32 TEM observations were made on cells harvested by centrifugation (600 g, 10 min) at the 12 day of 33 34 cultivation. Cells were fixed, post-fixed and dehydrated as reported in Baldisserotto et al. (2012). 35 36 Embedding in resin and staining procedures were performed as previously described (Pancaldi et al. 37 38 39 2002). Sections were observed with a Hitachi H800 electron microscope (Hitachi, Tokyo, Japan) at 40 41 the Electron Microscopy Centre, University of Ferrara. 42 43 44 45 46 Extracellular free fatty acids analysis 47 48 49 To analyze the FFA composition of the exhausted E and EG media used to prepare E+ and EG+ 50 51 replenished media, aliquots of N. oleoabundans cultures grown autotrophically and mixotrophically 52 53 in the PBR were collected at the 15th and at the 8th day of cultivation, respectively. Samples were 54 55 56 centrifuged at 2,000 g for 10 min in order to separate the medium from algae. Fifty mL of media 57 58 obtained from the autotrophic (E) and mixotrophic (EG) cultures were freeze-dried for analysis. 59 60 61 The extracellular FFA composition was determined in duplicate by gas chromatography-mass 62 63 64 9 65 spectrometry (GC-MS). For extraction, samples were dissolved in 3 mL of hexane, sonicated for 15 1 2 min and extracted overnight. Fatty acid methyl esters were prepared by transesterification with 1.5 3 4 5 mL of 5% of sodium hydroxide in methanol solution. Sample volumes of 1 μl were injected into the 6 7 GC-MS apparatus, which consisted of a Varian Saturn 2100 MS/MS ion trap mass spectrometer. 8 9 10 Separations were performed using a Zebron ZB-WAX Phenomenex capillary column (60 cm in 11 12 length, 0.25 mm i.d) supplied with helium carrier gas at 1 mL min-1 constant flow. The injector 13 14 temperature was 250 °C and the oven temperature programme was the following: start 100°C for 2 15 16 17 min, ramp to 200°C at 10 °C/min, and hold for 108 min. The MS acquisitions were performed by 18 19 full scan mode. 20 21 22 23 24 Determination of polyamines 25 26 27 To analyze the free PA composition of N. oleoabundans cells cultivated autotrophically and 28 29 mixotrophically in PBR and in the corresponding E and EG media, samples were collected as 30 31 32 described for FFA analyses. Freeze-dried E or EG media and algae (50 and 20 mg, respectively) 33 34 were extracted in cold 4% perchloric acid, kept for 1 h on ice, and then centrifuged at 15,000 g for 35 36 15 min. Aliquots (200 L) of the supernatants and standard solutions of putrescine (Put), 37 38 39 spermidine (Spd) and spermine (Spm) were derivatised with dansyl chloride (Scaramagli et al. 40 41 1999). Dansylated derivatives were extracted with toluene, taken to dryness and resuspended in 42 43 44 acetonitrile. PAs were separated and quantified by HPLC using a reverse phase C18 column 45 46 (Spherisorb ODS2, 5-μm particle diameter, 4.6 x 250 mm, Waters, Wexford, Ireland) and a 47 48 -1 49 programmed acetonitrile:water step gradient (flow rate 1 mL min ) on a Jasco system (Jasco Corp., 50 51 Tokyo, Japan) consisting of a PU-1580 pump, an LG-1580-02 ternary gradient unit, a DG-1580-53 52 53 54 three-line degasser and a FP-1529 fluorescence detector, linked to an autosampler (AS 2055 Plus). 55 56 57 58 59 60 61 62 63 64 10 65 Data treatment 1 2 3 Data were processed with Graphpad Prism 6 (Graph Pad Software, San Diego CA, USA). In each 4 5 case, means ± standard deviations for n number of samples are given. Statistical analyses for 6 7 8 comparison of means were carried out using ANOVA, followed by Student’s t-test (significance 9 10 level, 0.05). 11 12 13 14 15 16 17 Results 18 19 20 21 Growth and morphology of N. oleoabundans cultivated autotrophically and mixotrophically in a 22 23 20-L PBR 24 25 26 Growth kinetics and cell morphology of N. oleoabundans cultivated autotrophically and 27 28 mixotrophically in a 20-L PBR are shown in Fig 1. Autotrophic cells rapidly entered the 29 30 nd rd th th 31 exponential phase starting on the 2 -3 day up to the 7 -9 day of cultivation (Fig 1a). After this 32 33 period, cultures had a short late exponential phase and then entered the stationary phase of growth, 34 35 6 -1 36 reaching a final cell density of about 22 × 10 cells mL at the end of the of cultivation time (25 37 38 days). During the entire period, cells showed the typical morphology of N. oleoabundans grown in 39 40 BM medium, i.e., almost spherical, with a cell diameter of 3-5 µm (Fig 1b, d). One cup-shaped 41 42 43 chloroplast, containing a large pyrenoid, was present and emitted an intense red fluorescence due to 44 45 the presence of chlorophyll (Fig 1b,c). On the 25th day of cultivation, cells maintained their normal 46 47 48 features and dimensions, but some lipid globules accumulated inside the cytoplasm, as revealed by 49 50 Nile Red staining (Fig 1d,e). 51 52 53 During the first four days of mixotrophic growth in the PBR in the presence of glucose, cells 54 55 showed an evident increase in cell density, reaching values of ca. 33 x 106 cells mL-1 (Fig 1f). 56 57 Subsequently, cells entered the stationary phase of growth, and then cell density leveled off to 58 59 6 -1 60 values of about 36 x 10 cells mL up to the end of the cultivation period, i.e., eight days. At the 61 62 63 64 11 65 beginning of cultivation, both cell shape and size, and chloroplast features in mixotrophic cultures 1 2 (Fig 1g, h) were similar to those of autotrophic cultures (Fig 1b, c). Starting on the 3rd day and up to 3 4 5 the end of cultivation, however, mixotrophic cells showed peculiar features. In fact, the chloroplast 6 7 lost its characteristic cup shape and translucent globules, which tended to occupy almost the entire 8 9 10 cell volume, accumulated inside the cytoplasm (Fig 1i). Nile Red staining confirmed the lipidic 11 12 nature of these droplets (Fig 1j). 13 14 15 16 17 Growth kinetics of N. oleoabundans in exhausted and control growth media 18 19 As shown in Fig 2a, the growth of N. oleoabundans in E+ medium was promoted for the entire 20 21 22 experiment relative to controls grown on BM medium; on the contrary, growth in EG+ medium was 23 24 similar to that of the C. All samples entered the exponential phase very soon (during the first 3 days 25 26 27 of cultivation), despite some differences in growth rates (Fig 2b). The highest μ values were 28 29 observed for algae cultivated in the two exhausted media (E+ and EG+, 1.29 and 1.18 div d-1, 30 31 -1 32 respectively; control, 0.98 div d ; p <0.01 in both cases relative to C) (Fig 2b). Subsequently, from 33 34 the 3rd day onwards, no relevant differences were observed between C and EG+ samples, as both of 35 36 them showed similar growth kinetics and cell densities (Fig 2a). Conversely, E+ samples reached 37 38 39 and maintained the highest cell density; the major difference as compared to the other samples was 40 41 observed starting on the 7th day, with a cell density of about 16 × 106 cells mL-1 (45 and 50% more 42 43 44 than EG+ and C samples, respectively; p <0.01 in both cases relative to E+ samples) at the same 45 46 time point. After the 17th day, all samples entered the stationary phase, reaching, at the end of the 47 48 6 -1 49 experiment, a final cell density of 25-30 × 10 cells mL . 50 51 52 53 Consumption of nitrate and phosphate by N. oleoabundans cultivated in exhausted and control 54 55 56 growth media 57 58 The consumption of nitrate and phosphate by N. oleoabundans cells in the course of the experiment 59 60 61 is depicted in Fig 3. At time 0 (inoculation), the exhausted replenished growth media contained 62 63 64 12 65 - 2- approximately 2 mM NO3 and 0.115 mM PO4 , i.e., the typical concentration of those components 1 2 in BM medium. By the end of the experiment, cells had consumed practically all nitrate and 3 4 5 phosphate present in the growth media. However, nutrient consumption by C, E+ and EG+ algal 6 7 cultures followed different trends, since C samples maintained higher nitrate and phosphate 8 9 10 concentrations than samples in exhausted media. In fact, after 12 days of cultivation, cells grown in 11 12 E+ and EG+ had consumed about 34% and 68% of nitrate, respectively, while C only 15%. Nitrate 13 14 concentration, decreased from 1.43 mM at time 0 for all culture media to 0.94 mM for E+, 0.45 mM 15 16 17 for EG+ and 1.21 mM for C media (p <0.001 in both treated samples relative to C; Fig 3a). 18 19 Differently, during the first 12 days of growth, phosphate exhibited a dramatic decrease, with a 20 21 22 reduction by cells cultivated in EG+ and E+ of 94% and 98%, respectively, while in C medium it 23 24 was only about 60%. In fact, phosphate concentration decreased from about 0.086 mM at time 0 for 25 26 27 all culture media to 0.002 for E+, 0.005 for EG+ and 0.035 mM for C media (p <0.001 in both 28 29 treated samples relative to C; Fig 3b). 30 31 32 33 34 Photosynthetic pigment content in N. oleoabundans cultivated in exhausted and control media 35 36 During the experiment a gradual increase in photosynthetic pigment content was observed in all 37 38 39 samples (Fig 4). Chla in algae grown in C medium showed an evident increase up to the end of the 40 41 experiment, reaching an average concentration of 0.5 nmol 10-6 cells (Fig 4a). A similar increasing 42 43 44 trend was also observed in E+ samples, though with values 10-15% lower than those of controls. 45 46 From the 17th day of cultivation, Chla content of E+ samples was significantly lower than in C 47 48 49 (32%; p<0.01). Conversely, in EG+ samples Chla content showed a slightly increasing trend as 50 51 compared with C and E+ samples; this led to a final value of 0.35 nmol 10-6 cells at the end of the 52 53 experiment (32 and 21% lower than C and E+ samples, respectively; Fig 4a). An increasing trend in 54 55 56 Chlb content was also observed for all samples, without significant differences between C and E+ 57 58 samples (Fig 4b). Similar to Chla content, EG+ cells contained lower quantities of Chlb than C and 59 60 61 E+ samples (15-25% and 20-35% lower than C and E+ samples, respectively). More evident 62 63 64 13 65 differences were observed for Car content (Fig 4c). In fact, C samples showed an evident increase 1 2 up to the end of the experiment, always containing higher quantities compared to the other samples 3 4 5 (20-45% and 15-30% more than E+ and EG+ samples, respectively; p<0.05). It is noteworthy that 6 7 EG+ and E+ cells shared a similar trend of Car concentration during the experiment (Fig 4c). 8 9 10 11 12 13 14 Maximum quantum yield of PSII of N. oleoabundans cultivated in exhausted and control media 15 16 17 The variations in PSII maximum quantum yield measured during the experiments are shown in Fig 18 19 5. C, E+ and EG+ samples showed a slight increase of the F /F ratio during the first three days of 20 V M 21 22 growth, reaching values of ca. 0.70. Subsequently, C cells maintained stable values around 0.70- 23 24 0.75, while the FV/FM ratio of E+ and EG+ samples decreased drastically down to values below 25 26 rd 27 0.50 at the end of experiment. In EG+ cells the decrease was dramatic already from the 3 day of 28 29 cultivation, reaching the lowest value after 17 days (ca. 0.30 for EG+ vs 0.75 for C and 0.50 for 30 31 32 E+; p <0.01 in both cases); thereafter, samples maintained stable values of ca. 0.35 until the end of 33 34 the experiment. On the contrary, in E+ cells, FV/FM started to decrease very strongly only from the 35 36 7th day of cultivation until the 17th day (from about 0.70 at day 7 down to 0.50 at day 17). During 37 38 th th 39 the subsequent experimental times (21 and 25 day), the FV/FM ratio of E+ cells remained stable 40 41 around 0.45. 42 43 44 45 46 47 48 49 Morphological observations of N. oleoabundans cultivated in exhausted and control media 50 51 52 Light microscopy and Nile Red staining 53 54 Light microscopy of both control and treated samples showed that N. oleoabundans maintained 55 56 similar cell morphology and dimensions throughout the experiment (Fig 6, 7). In fact, cells were 57 58 59 almost spherical with a cell diameter of 3-5 μm. One cup-shaped chloroplast, containing a large 60 61 pyrenoid, was present inside the cells (Fig 6a, c, e). Moreover, the chloroplast emitted an intense 62 63 64 14 65 red fluorescence due to Chl (Fig 6b, d, f). Interestingly, after 25 days of cultivation, while cell size 1 2 and shape remained substantially unchanged, all algal samples showed some translucent granulation 3 4 5 at the cytoplasmic level (Fig 7a, c, e). In order to understand the nature of those granulations and to 6 7 investigate if the recycled growth media could promote the production of lipid globules, all samples 8 9 10 were periodically stained with the lipid-specific fluorochrome Nile Red. The reaction was positive 11 12 only at the end of the experiment and the translucent globules were then unequivocally identified as 13 14 lipid droplets (Fig 7b, d, f). 15 16 17 18 19 TEM observations 20 21 22 To investigate the morphological and cytological changes induced by cultivation in recycled growth 23 24 media, the ultrastructure of C, E+ and EG+ samples was observed by TEM. At 12 days of 25 26 27 cultivation, most of the C cell volume was occupied by a characteristic cup-shaped chloroplast. 28 29 Inside the plastid one large pyrenoid, which was crossed by two elongated and appressed 30 31 32 thylakoids, was present (Fig 8a). In particular, the organelle contained starch in the shape of a shell 33 34 around the pyrenoid, and showed the typical thylakoid organization (Fig 8b). Typically featured 35 36 chloroplasts, as described for cells grown in C, were observed in cells grown in E+ medium (Fig 37 38 39 8c); however, thylakoids were more appressed than in C (Fig 8d). EG+ samples showed a more 40 41 strongly altered chloroplast structure (Fig 8e), as compared with C and E+ samples. Photosynthetic 42 43 44 membranes showed different degrees of thylakoid appression: some portions of thylakoid 45 46 membranes were appressed while others were loose and sometimes swollen (Fig 8f,g). In addition, 47 48 49 large portions of the stroma were free of thylakoids and some plastoglobules in proximity of the 50 51 thylakoid membranes were also visible (Fig 8g, h). Finally, in EG+ samples the pyrenoid lost its 52 53 round shape and appeared malformed (Fig 8e). 54 55 56 57 58 59 60 61 62 63 64 15 65 Extracellular FFA composition in E and EG media 1 2 The extracellular FFA composition in the exhausted autotrophic (E) and mixotrophic (EG) media of 3 4 5 N. oleoabundans is shown in Table 1. While EG medium comprised both saturated and mono- 6 7 unsaturated FFAs, E medium contained only saturated FFAs. In fact, both samples contained 8 9 10 similar percentages of saturated myristic acid (C14:0) and palmitic acid (C16:0), while the 11 12 percentage of stearic acid (C18:0) was different between the two samples. In E medium, stearic acid 13 14 was present at about 64%, while in EG medium at about 56%; however, C18:0 represented the main 15 16 17 saturated FFA for both samples. It is noteworthy that only EG medium contained about 6% of the 18 19 monoenoic oleic acid (C18:1ω9). 20 21 22 23 24 Polyamine content in E and EG media and in the corresponding N. oleoabundans cells cultivated in 25 26 27 a 20-L PBR 28 29 As shown in Fig 9a, the PA composition of N. oleoabundans cells cultivated mixotrophically in the 30 31 32 PBR were characterized by higher putrescine, spermine, and, especially, spermidine levels 33 34 compared to cells cultivated autotrophically. In fact, Spm, Put and Spd concentrations in 35 36 mixotrophic cells were ca. 2, 30 and 57 times higher, respectively, than those found in autotrophic 37 38 39 cells (p<0.001). Conversely, the PA composition in E and EG medium was not significantly 40 41 different. In fact, both media contained similar amounts of Put and Spm. Interestingly, spermidine 42 43 -1 44 was present in high concentration in both media (119 ± 16.91 and 101 ± 5.00 pmol mL in E and 45 46 EG, respectively; Fig 9b). 47 48 49 50 51 52 Discussion 53 54 55 56 The recycling of culture medium has been proposed as a possible solution in order to reduce water 57 58 consumption for algal cultivation, thereby making the process more economically feasible and 59 60 61 environmentally sustainable (Yang et al. 2011; Hadj-Romdhane et al. 2013; Farooq et al. 2014). 62 63 64 16 65 Yang et al. (2011) estimated that the large-scale cultivation of the microalga Chlorella vulgaris in 1 2 recycled culture medium could reduce water use by about 84%. Present results clearly suggest that 3 4 5 N. oleoabundans can also efficiently grow in exhausted growth media, especially in the autotrophic 6 7 medium replenished with nitrate and phosphate (E+). Thus, in order to make the use of recycled 8 9 10 culture medium feasible, exhausted media should be replenished with the correct ratio of nitrate and 11 12 phosphate, as they are the main nutrients that guarantee cell growth (Stephens et al. 2010). This 13 14 aspect was preliminarily verified by employing unmodified exhausted growth media (E and EG) to 15 16 17 re-cultivate N. oleoabundans. In fact, cells showed a slight growth, reaching, at the end of the 18 19 cultivation period, low cell densities (ca. 11 vs 32 × 106 cells mL-1 of C cultures) (data not shown). 20 21 22 These cells accumulated intracellular lipids throughout the experiment (data not shown) as a 23 24 consequence of the limitation of nitrate and phosphate (Mata et al. 2010; Popovich et al. 2012). 25 26 27 These results are consistent with those of Zhu et al. (2013), who found that C. zofingensis cultivated 28 29 in recycled medium, without nitrogen and phosphorus, displayed enhanced lipid production 30 31 32 compared to cultures with full nutrients. 33 34 In order to understand if the growth promotion of N. oleoabundans cultivated in E+ and EG+, 35 36 relative to C, was influenced by some molecules released from the algae, a characterization of 37 38 39 autotrophic (E) and mixotrophic (EG) media was performed. In fact, Moheimani and Borowitzka 40 41 (2006) and Stephen et al. (2010) reported that molecules released from cells can 42 43 44 positively/negatively alter cell metabolism and biomass production. 45 46 Here we show that both exhausted media contained the main PAs (putrescine, spermidine, and 47 48 49 spermine). It is known that these plant growth regulators are involved in a great variety of 50 51 developmental processes in plant cells, e.g., cell division and protein synthesis (Kaur-Sawhney et 52 53 al. 2003; Kuznetsova et al. 2006). PA biosynthetic pathways have also been studied in green algae 54 55 56 (Cohen et al. 1984; Theiss et al. 2002; Fuell et al. 2010). Although algae produce “unusual” PAs, 57 58 such as homospermidine and thermospermine (Hamana et al. 2013), the more common PAs (Put, 59 60 61 Spd and Spm), when added to the media, promoted growth and metabolism in C. vulgaris (Czerpak 62 63 64 17 65 et al. 2003). PAs are also known to alleviate the effect of biotic and abiotic stress in plants as well 1 2 as algae (Tate et al. 2013). 3 4 5 In the present work, a very strong increase in Put and, especially, Spd concentrations was observed 6 7 in N. oleoabundans mixotrophic cells as compared to the autotrophic ones, both cultivated in a 20-L 8 9 10 PBR. The growth-promoting role of PAs could explain the faster (0-4 days) and stronger cell 11 12 density increase (ca. 33 x 106 cell mL-1) in mixotrophic cultures relative to the autotrophic ones (0-9 13 14 days; ca. 22 x 106 cell mL-1). The higher PA concentrations in mixotrophically grown cells also 15 16 17 suggest that, under these culture conditions, algal cells may be better protected from stress-inducing 18 19 factors, e.g., bacterial contamination. To our knowledge, the presence of PAs released into algal 20 21 22 growth medium in PBRs has not been documented before. The release of PAs (predominantly Spd) 23 24 from autotrophic and mixotrophic cells into their respective (E and EG) exhausted media was not 25 26 27 significantly different. However, the presence of these plant growth regulators seems to contribute 28 29 to the promotion of N. oleoabundans growth in both media replenished with nitrate and phosphate. 30 31 32 In fact, during the entire experiment, a higher cell density was observed in E+ relative to C medium. 33 34 Moreover, the EG+ samples showed a promotion of cell density, albeit similar to that of C. 35 36 Both autotrophic and mixotrophic exhausted media also contained FFAs, which could have induced 37 38 39 a change in the normal metabolism of N. oleoabundans. This assumption is demonstrated by the 40 41 more rapid consumption of nitrate and phosphate, the decreased photosynthetic pigments content 42 43 44 and the strong decline in the FV/FM ratio observed in cells grown in E+ and EG+ media. These 45 46 results corroborate the assumption that microalgae release metabolites in high cell density cultures 47 48 49 (Richmond 2004) or as a consequence of stressful conditions (Ikawa 2004; Wu et al. 2006). Even if 50 51 the role of FFAs is currently under debate (Ikawa 2004; Stephen et al. 2010), recent studies showed 52 53 that they can strongly inhibit growth or exert cytotoxic effects on microalgae (Kogteva and 54 55 56 Bezuglov 1998; Wu et al. 2006; Bosma et al. 2008; Stephen et al. 2010). In fact, the presence of 57 58 these metabolites, and their oxidative products, negatively affect biomass productivity, especially 59 60 61 when the microalgae are cultivated in recycled medium (Lívanský et al. 1996; Rodolfi et al. 2003). 62 63 64 18 65 Granum et al. (2002) reported that the accumulation of intracellular lipids in microalgae enhanced 1 2 the release of FFAs into the culture medium, and Harun et al. (2010) observed that this release was 3 4 5 caused by cell lysis. 6 7 In this work, FFA composition in autotrophic (E) and mixotrophic (EG) media after N. 8 9 10 oleoabundans cultivation was similar. The only main difference was represented by the presence of 11 12 the monoenoic oleic acid (C18:1ω9) in EG medium. Oleic acid is one of the major lipid 13 14 components in lipid-enriched N. oleoabundans grown under N-stress conditions (Popovich et al. 15 16 17 2012). It is reported that mixotrophic cultivation alters N:C ratio, inducing similar lipid production 18 19 as N-depleted autotrophic cultures (Giovanardi et al. 2014). Probably for this reason, the release of 20 21 22 oleic acid occurred only in the mixotrophic growth medium. Moreover, Wu et al. (2006) observed 23 24 altered plasma membrane permeability due to the toxic effects of FFAs in two Chlorophyta (C. 25 26 27 vulgaris and Monoraphidium contortum) and in a cyanobacterium (Anabaena P-9). The rapid 28 29 consumption of phosphate in both exhausted media could be linked to an alteration in membrane 30 31 32 permeability, due to the activity of FFAs (Ikawa 2004; Wu et al. 2006). In addition, FFAs can cause 33 34 inhibition of the PSII and PSI electron transport chains (Siegenthaler 1973) and disorganization of 35 36 thylakoids (Wu et al. 2006). Indeed, N. oleoabundans cells in E+ and EG+ were characterized by 37 38 39 alterations of the photosynthetic apparatus. These cells, in fact, contained less photosynthetic 40 41 pigments than C cells, and exhibited a drastic decrease of the PSII maximum quantum yield 42 43 44 (FV/FM). These variations are linked to an alteration of photosynthetic efficiency, especially as 45 46 regards PSII (White et al. 2011). The decreased photosynthetic pigment contents, observed in cells 47 48 49 grown in E+ and EG+, also reflected the changes in thylakoid membrane arrangement 50 51 (Baldisserotto et al. 2012). Therefore, correct thylakoid organization is necessary to maintain 52 53 optimal photosynthetic activity and this is often influenced by culture conditions (Nevo et al. 2012). 54 55 56 In addition, the presence of some plastoglobules in chloroplasts of algal cells, grown on exhausted 57 58 media, could be another indicator of an alteration of photosynthetic membranes (Besagni and 59 60 61 Kessler 2013). On the contrary, N. oleoabundans cells grown in C medium showed stable and 62 63 64 19 65 normal FV/FM values throughout the experiment (Giovanardi et al. 2014), and the typical assembly 1 2 of thylakoid membranes (Nevo et al. 2012; Baldisserotto et al. 2012; Giovanardi et al. 2014). 3 4 5 The presence of lipid droplets, observed only at the end of the experiment in C, E+ and EG+ cells, 6 7 is probably related to aging of the microalgal cultures (Hu et al. 2008; Baldisserotto et al. 2012), 8 9 10 rather than to the use of recycled culture media. 11 12 13 14 In conclusion, results presented here demonstrate that recycling autotrophic and mixotrophic 15 16 17 growth media is a suitable solution to obtain high cell density cultures of the microalga N. 18 19 oleoabundans. For this reason, they represent a contribution for improving the scale-up of 20 21 22 microalgal cultivation while providing a more sustainable ecological impact on water resources. 23 24 However, further studies are needed to deepen knowledge on the nature and the specific role of 25 26 27 some molecules that are released into the growth media, in order to obtain useful information for 28 29 the advancement of the biotechnological use of this strain. 30 31 32 33 34 Acknowledgements 35 36 This work was financially supported by grants from the University of Ferrara. We are grateful to 37 38 39 Dr. Immacolata Maresca of the University of Ferrara for technical assistance in freeze-drying of 40 41 samples. 42 43 44 45 46 Conflict of interest 47 48 49 The authors declare they have no conflict of interest. 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 20 65 References: 1 2 Baldisserotto C, Ferroni L, Giovanardi M, Pantaleoni L, Boccaletti L, Pancaldi S (2012) Salinity 3 4 5 promotes growth of freshwater Neochloris oleoabundans UTEX 1185 (Sphaeropleales, 6 7 Neochloridaceae): morpho-physiological aspects. Phycologia 51:700-710 8 9 10 Baldisserotto C, Giovanardi M, Ferroni L, Pancaldi S (2014) Growth, morphology and 11 12 photosynthetic responses of Neochloris oleoabundans during cultivation in a mixotrophic 13 14 brackish medium and subsequent starvation. Acta Physiol Plant 36:461-472 15 16 17 Batan L, Quinn CJ, Bradley TH (2013) Analysis of water footprint of a photobioreactor microalgae 18 19 biofuel production system from blue, green and lifecycle perspectives. Algal Research 2:196- 20 21 22 203 23 24 Besagni C and Kessler F (2013) A mechanism implicating plastoglobules in thylakoid disassembly 25 26 27 during senescence and nitrogen starvation. Planta 237:463-470 28 29 Borowitzka MA (2005) Culturing microalgae in outdoor ponds. In: Anderson RA (ed) Algal 30 31 32 culturing techniques. Elsevier Academic, London, pp 205-218 33 34 Borowitzka MA and Moheimani NR (2013) Sustainable biofuels from algae. Mitig Adapt Strateg 35 36 Glob Change 18:13-25 37 38 39 Bosma R, Miazek K, Willemsen SM, Vermuë MH, Wijffels RH (2008) Growth inhibition of 40 41 Monodus subterraneus by free fatty acids. Biotechnol Bioeng 101:1108–1114 42 43 44 Chisti Y (2007) Biodiesel from microalgae. Biotech Adv 25:294-306 45 46 Czerpak R, Bajguz A, Pietrowska A, Dobrogowska R, Matejczyk M, Weislawski W (2003) 47 48 49 Biochemical activity of di- and polyamines in the green alga Chlorella vulgaris Beijerinck 50 51 (Chlorophyceae). Acta Soc Bot Pol 72:19–24 52 53 Cohen E, Arad SM, Heimer YH, Mizrahi Y (1984) Polyamine biosynthetic enzymes in the cell cycle 54 55 56 of Chlorella. Plant Physiol 74:385-388 57 58 59 Farooq W, Suh WI, Park MS, Yang J (2014) Water use and its recycling in microalgae cultivation for 60 61 biofuel application. Bioresource Technol doi: 10.1016/j.biortech.2014.10.140 62 63 64 21 65 Ferroni L, Baldisserotto C, Pantaleoni L, Billi P, Fasulo MP, Pancaldi S (2007) High salinity alters 1 2 chloroplast morpho-physiology in a freshwater Kichneriella species (Selenastraceae) from 3 4 5 Ethipian Lake Awasa. Am J Bot 94:1973-1983 6 7 Ferroni L, Baldisserotto C, Giovanardi M, Pantaleoni L, Morosinotto T, Pancaldi S (2011) Revised 8 9 10 assignment of room-temperature chlorophyll fluorescence emission bands in single living cells 11 12 of Chlamydomonas reinhardtii. J Bioenerg Biomembr 43:163-173 13 14 Fuell C, Elliott KA, Hanfrey CC, Franceschetti M, Michael AJ (2010) Polyamine biosynthetic 15 16 17 diversity in plants and algae. Plant Physiol Biochem 48:513–520 18 19 Giovanardi M, Ferroni L, Baldisserotto C, Tedeschi P, Maietti A, Pantaleoni L, Pancaldi S (2013) 20 21 22 Morpho-physiological analyses of Neochloris oleoabundans (Chlorophyta) grown 23 24 mixotrophically in a carbon-rich waste product. Protoplasma 250:161-174 25 26 27 Giovanardi M, Baldisserotto C, Ferroni L, Longoni P, Cella R, Pancaldi S (2014) Growth and lipid 28 29 synthesis promotion in mixotrophic Neochloris oleoabundans (Chlorophyta) cultivated with 30 31 32 glucose. Protoplasma 251:115-125 33 34 Gouveia L, Marques AE, Lopes da Silva T, Reis A (2009) Neochloris oleoabundans UTEX #1185: a 35 36 suitable renewable lipid source for biofuel production. J Ind Microbiol Biotechnol 36: 821-826 37 38 39 Granum E, Kirkvold S, Myklestad SM (2002) Cellular and extracellular production of carbohydrates 40 41 and amino acids by the marine diatom Skeletonema costatum: diel variations and effects of N 42 43 44 depletion. Mar Ecol Prog Ser 242:83–94. 45 46 Grobbelaar JU (2009) Factors governing algal growth in photobioreactors: the “open” versus 47 48 49 “closed” debate. J Appl Phycol 21:489-492 50 51 Hadj-Romdhane F, Jaouen P, Pruvost J, Grizeau D, Van Vooren G, Bourseau P (2012) Development 52 53 and validation of a minimal growth medium for recycling Chlorella vulgaris culture. 54 55 56 Bioresource Technol 123:366-374 57 58 Hadj-Romdhane F, Zheng X, Jaouen P, Pruvost J, Grizeau D, Croué JP, Bourseau P (2013) The 59 60 61 culture of Chlorella vulgaris in a recycled supernatant: Effects on biomass production and 62 63 64 22 65 medium quality. Bioresource Technol 132: 285–292 1 2 Hamana K, Niitsu M, Hayashi H (2013) Occurrence of homospermidine and thermospermine as a 3 4 5 cellular polyamine in unicellular chlorophyte and multicellular charophyte green algae. J Gen 6 7 Appl Microbiol 59: 313-319 8 9 10 Harun R, Singh M, Forde GM, Danquah MK (2010) Bioprocess engineering of microalgae to 11 12 produce a variety of consumer products. Renew Sust Energ Rev 14:1037-1047 13 14 Hu Q, Sommerfeld M, Jarvis E, Ghirardi M, Posewitz M, Seibert M, Darzins A (2008) Microalgal 15 16 17 triacylglycerols as a feedstocks for biofuel production: perspective and advances. Plant J 18 19 54:621-639 20 21 22 Ikawa M (2004) Algal polyunsaturated fatty acids and effects on plankton ecology and other 23 24 organism. UNH Center for Freshwater Biology Research 6:17-44 25 26 27 Kalaji HM, Schansker G, Ladle RJ, Vasilij Goltsev V, Bosa K, Allakhverdiev SI, Brestic M, Bussotti 28 29 F, Calatayud A, Dąbrowski P, Elsheery NI, Ferroni L, Guidi L, Hogewoning SW, Jajoo A, 30 31 32 Misra AN, Nebauer SG, Pancaldi S, Penella C, Poli D, Pollastrini M, Romanowska-Duda ZB, 33 34 Rutkowska B, Serôdio J, Suresh K, Szulc W, Tambussi E, Yanniccari M, Zivcak M (2014) 35 36 Frequently asked questions about in vivo chlorophyll fluorescence: practical issues. Photosynth 37 38 39 Res 122:121-158 40 41 Kaur-Sawhney R, Tiburcio AF, Altabella T, Galston AW (2003) Polyamines in plants: an overview. 42 43 44 J Cell Mol Biol 2:1-12 45 46 Kogteva GS and Bezuglov VV (1998) Unsaturated fatty acids as endogenous bioregulators. 47 48 49 Biochemistry (Moscow) 63:4-12 50 51 Lam MK and Lee KT (2012) Potential of using organic fertilizer to cultivate Chlorella vulgaris for 52 53 54 biodiesel production. Appl Energy 94:303-308 55 56 Li Y, Horsman M, Wang B, Wu N, Lan CQ (2008) Effect on nitrogen sources on cell growth and 57 58 lipid accumulation of green alga Neochloris oleoabundans. Appl Microbiol Biotechnol 81:629- 59 60 61 636 62 63 64 23 65 Lichtenthaler HK, Buschmann C, Knapp M (2005) How to correctly determine the different 1 2 chlorophyll fluorescence parameters and the chlorophyll fluorescence decrease ratio RFd of 3 4 5 leaves with the PAM fluorometer. Photosynthetica 43:379-393 6 7 Lívanský K, Dědič K, Bínová J, Tichý V, Novotný P, Doucha J (1996) Influence of the nutrient 8 9 10 solution recycling on the productivity of Scenedesmus obliquus, utilization of nutrients and 11 12 water in outdoor cultures. Algological Studies Archiv fu r Hydrobiologie 81:105–113 13 14 Mata TM, Martins AA, Caetano NS (2010) Microalgae for biodiesel production and other 15 16 17 applications: a review. Renew Sust Energ Rev 14:217- 232 18 19 Moheimani NR and Borowitzka MA (2006) The long-term culture of the coccolithophore 20 21 22 Pleurochrysis carterae (Haptophyta) in outdoor raceway ponds. J Appl Phycol (2006) 18:703- 23 24 712 25 26 27 Molina Grima E, Belarbi H, Acién Fernández FG, Robles Medina A, Chisti Y (2003) Recovery of 28 29 microalgal biomass and metabolites: process options and economics. Biotech Adv 20: 491-515 30 31 32 Ndimba BK, Ndimba RJ, Johnson TS, Waditee-Sirisattha R, Baba M, Sirisattha S, Shiraiwa Y, 33 34 Rakwal R, Agrawal GK, Rakwal R (2013) Biofuels as a sustainable energy source: An update 35 36 of the applications of proteomics in bioenergy crops and algae. J Prot 93:234-244 37 38 39 Nevo R, Charuni D, Tsabari O, Reich Z (2012) Composition, architecture and dynamics of the 40 41 photosynthetic apparatus in higher plants. Plant J 70:157-176 42 43 44 Pancaldi S, Baldisserotto C, Ferroni L, Bonora A, Fasulo MP (2002) Room temperature 45 46 microspectrofluorimetry as a useful tool for studying the assembly of the PSII chlorophyll– 47 48 49 protein complexes in single living cells of etiolated Euglena gracilis Klebs during the greening 50 51 process. J Exp Bot 53:1753–1763 52 53 Popovich CA, Damiani MC, Constenla D, Martínez AM, Giovanardi M, Pancaldi S, Leonardi PI 54 55 56 (2012) Neochloris oleoabundans grown in natural enriched seawater for biodiesel feedstock: 57 58 Evaluation of its growth and biochemical composition. Bioresource Technol 114:287-293 59 60 61 Pruvost J, Van Vooren G, Cogne G, Legrand J (2009) Investigation of biomass and lipids production 62 63 64 24 65 with Neochloris oleoabundans in photobioreactor. Bioresource Technol 100:5988-5995 1 2 Pulz O (2001) Photobioreactors: production systems for phototrophic microorganisms. Appl 3 4 5 Microbiol Biotechnol 57:287-293 6 7 Richmond A (2004) Handbook of Microalgal Culture: Biotechnology and Applied Phycology. 8 9 10 Blackwell Sciences Ltd., Oxford 11 12 Rodolfi L, Zittelli GC, Barsanti L, Rosati G, Tredici MR (2003) Growth medium recycling in 13 14 Nannochloropsis sp. mass cultivation. Biomol Eng 20:243–248 15 16 17 Scaramagli S, Biondi S, Capitani F, Gerola P, Altamura MM, Torrigiani P (1999) Polyamine 18 19 conjugate levels and ethylene biosynthesis: inverse relationship with vegetative bud formation 20 21 22 in tobacco thin layers. Physiol Plant 105:366-375 23 24 Sheehan JJ (2009) Biofuels and the conundrum of sustainability. Curr Opin Biotechnol 20:318–324 25 26 27 Siegenthaler PA (1973) Change in pH dependence and sequential inhibition of photosynthetic 28 29 activity in chloroplasts by unsaturated fatty acids. Biochim Biophys Acta 305:153-162 30 31 32 Smith VH, Sturm BSM, deNoyelles FJ, Billings SA (2010) The ecology of algal biodiesel 33 34 production. Trends Ecol Evo 25:301-309 35 36 Solomon BD (2010) Biofuels and sustainability. Ann NY Acad Sci 1185:119-134 37 38 39 Stephens E, Ross IL, Mussgnug JH,Wagner LD, BorowitzkaMA, Posten C, Kruse O, Hankamer B 40 41 (2010) Future prospects of microalgal biofuel production systems. Trends Plant Sci 15:554-564 42 43 44 Tate JJ, Gutierrez-Wing MT, Rush KA, Benton MG (2013) The Effects of Plant Growth Substances 45 46 and Mixed Cultures on Growth and Metabolite Production of Green Algae Chlorella sp.: A 47 48 49 Review. J Plant Growth Regul 32:417–428 50 51 Tornabene TG, Holzer G, Lien S, Burris N (1983) Lipid composition of the nitrogen starved green 52 53 alga Neochloris oleoabundans. Enzyme Microbiol Tech 5:435-440 54 55 56 Theiss C, Bohley P, Voigt J (2002) Regulation by polyamines of ornithine decarboxylase activity and 57 58 cell division in the unicellular green alga Chlamydomonas reinhardtii Plant Physiol 128:1470- 59 60 61 1479 62 63 64 25 65 Wellburn AR (1994) The spectral determination of chlorophylls a and b, as well as total carotenoids, 1 2 using various solvents with spectrophotometer of different resolution. J Plant Physiol 144:307– 3 4 5 313 6 7 White S, Anandraj A, Bux F (2011) PAM fluorometry as a tool to assess microalgal nutrient stress 8 9 10 and monitor cellular neutral lipids. Bioresource Technol 102:1675-1682 11 12 Wu JT, Chiang YR, Huang WY, Jane WN (2006) Cytotoxic effects of free fatty acids on 13 14 phytoplankton algae and cyanobacteria. Aquat Toxicol 80:338–345 15 16 17 Yang, J, Xu M, Zhang X, Hu Q, Sommerfeld M, Chen Y (2011) Lifecycle analysis on biodiesel 18 19 production from microalgae: Water footprint and nutrients balance. Bioresource Technol 20 21 22 102:159-165 23 24 Zhu LD, Takala J, Hiltunen E, Wang ZM (2013) Recycling harvest water to cultivate Chlorella 25 26 27 zofingiensis under nutrient limitation for biodiesel production. Bioresource Technol 144:14-20 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 26 65 Figure captions: 1 2 Fig 1 (a-e) Growth kinetics (a) and morphology (b-e) of N. oleoabundans cells autotrophically 3 4 5 cultivated in a 20-L PBR in BM medium. Light (b, d) and fluorescence microscope observations (c, 6 7 e) of N. oleoabundans cells after 3 (b, c) and 25 (d, e) days of autotrophic cultivation. (f-j) Growth 8 9 10 kinetics (f) and cell morphology (g-j) of N. oleoabundans cultivated in a 20-L PBR in BM medium 11 12 under mixotrophic conditions (supplemented with 2.5 gL-1 of glucose). Light (g, i) and fluorescence 13 14 microscope observations (h, j) of N. oleoabundans cells after 3 (g, h) and 8 (d, e) days of 15 16 17 mixotrophic cultivation. In both graphs, curves are constructed on a log2 scale and data are means ± 18 19 s.d. (n=3). In all micrographs, bars: 2 μm 20 21 22 23 24 Fig 2 (a) Growth kinetics of N. oleoabundans in BM medium (filled circles), E+ medium (empty 25 26 27 squares) and EG+ medium (empty diamonds). (b) Growth rates, calculated during the exponential phase 28 29 (0–3 days time interval of cultivation), of cells grown in BM (white), E+ (light grey) and EG+ (dark 30 31 32 grey) media. The growth curve is constructed on a log2 scale and data are means ± s.d. (n=3) 33 34 35 36 Fig 3 Time-course variations in the concentrations of nitrate (a) and of phosphate (b) in culture media of 37 38 39 N. oleoabundans during 25 days of cultivation on BM medium (white), E+ medium (light grey) and 40 41 EG+ medium (dark grey). Values are means ± s.d. (n=3) 42 43 44 45 46 Fig 4 Time-course variations of Chla (a), Chlb (b) and Car content (c) in N. oleoabundans cells grown 47 48 49 in BM (filled circles), E+ (empty squares) and EG+ (empty diamonds) media during the 25 days of 50 51 cultivation. Values are means ± s.d. (n=3) 52 53 54 55 56 Fig 5 Time-course variations of PSII maximum quantum yield (FV/FM ratio) in N. oleoabundans cells 57 58 grown in BM (filled circles), E+ (empty squares) and EG+ (empty diamonds) media during the 25 days 59 60 61 of cultivation. Values are means ± s.d. (n=3) 62 63 64 27 65 Fig 6 Light and fluorescence microscopy observations of N. oleoabundans cells at 12 days of 1 2 cultivation. (a) Control cells and (b) the relative fluorescence of the chloroplast; (c) cell grown in 3 4 5 E+ medium and (d) the relative fluorescence of the chloroplast; (e) cell grown in EG+ medium and 6 7 (f) the relative fluorescence of the chloroplast. In all micrographs, bars: 2 μm 8 9 10 11 12 Fig 7 Light and epifluorescence pictures of N. oleoabundans cells after 25 days of growth. (a) 13 14 Control cells and (b) relative Nile Red-staining observation, (c) cells grown in E+ medium and (d) 15 16 17 relative Nile Red-staining observation, and (e) cells grown in E+ medium and (f) relative Nile Red- 18 19 staining observation in EG+ medium. Translucent globules are indicated with arrows. In all 20 21 22 micrographs, bars: 2 μm 23 24 25 26 27 Fig 8 Transmission electron micrographs of N. oleoabundans cells at 12 days of cultivation. (a) Cell 28 29 grown in BM medium and (b) detail of its chloroplast showing the typical thylakoid membranes 30 31 32 organization (white arrow). (c) Cell grown in E+ medium and (d) detail of its chloroplast with quite 33 34 compact and appressed (white arrow) thylakoids. (e) Cell grown in EG+ medium and (f-h) details 35 36 of its chloroplast. The presence of intermediate stages of thylakoid membranes are evident in EG+ 37 38 39 cells, showing a general disorganisation of thylakoid membranes (f-h), which appeared wavy, loose 40 41 and sometimes swollen (f, g). (h) Some plastoglobules in proximity of thylakoid membranes are 42 43 44 also observed (arrowheads). P, pyrenoid; N, nucleus, M, mithochondrion. Bars: 0.5 μm (a, c, e); 45 46 0.05 μm (b, d, f, g, h) 47 48 49 50 51 Fig 9 Polyamine concentrations (a) in N. oleoabundans cells cultivated in a 20-L PBR 52 53 autotrophically and mixotrophically at 15 and 8 days of cultivation, respectively, and (b) in the 54 55 56 corresponding exhausted E and EG growth media. Autotrophic cells (white), mixotrophic cells (dark), 57 58 E+ (light grey) and EG+ medium (dark grey). Values are means ± s.d. (n=3) 59 60 61 62 63 64 28 65 Table

Table 1 Extracellular free fatty acids composition in the exhausted E and EG media of N. oleoabundans grown in 20-L PBR, after 15 and 8 days of cultivation, respectively. Fatty Acids Abbreviation E medium EG medium

Myristic Acid C14:0 2.16 ± 0.09 2.09 ± 0.04 Palmitic Acid C16:0 33.54 ± 0.01 34.43 ± 2.80 Stearic Acid C18:0 64.31 ± 0.10 56.56 ± 2.71 Oleic Acid C18:1ω9 nd 5.93 ± 1.47 Values are expressed as percentage of total fatty acids and are means ± s.d. (n=2). Figure

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g hh Figure Click here to download Figure: Fig 9.tif Supplementary Material Click here to download Supplementary Material: Figure S1.pdf