Improving the reliability of fluorescence-based neutral lipid content measurements in microalgal cultures

Algal Research 1 (2012) 176–184

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Improving the reliability of fluorescence-based neutral lipid content measurements in microalgal cultures H. De la Hoz Siegler a, W. Ayidzoe a, A. Ben-Zvi a, R.E. Burrell a, b, W.C. McCaffrey a,⁎ a b

Chemical and Materials Engineering Department, University of Alberta, Edmonton, T6G 2V4, Canada Biomedical Engineering Department, University of Alberta, Edmonton, T6G 2V2, Canada

a r t i c l e

i n f o

Article history: Received 21 January 2012 Received in revised form 15 July 2012 Accepted 16 July 2012 Available online 10 August 2012 Keywords: Nile Red BODIPY Fluorescence Microalgae High-throughput

a b s t r a c t Fast and accurate determination of lipid content in microalgal cultures is a required step for algal bioprocess development and optimization. An improved high-throughput Nile Red staining protocol using a microplate fluorescence reader is proposed, resulting in an increased correlation coefficient and a reduction in the relative standard deviation of 84% when compared to previously reported staining protocols. Differences in the staining efficacy of Nile Red among different algal strains were observed, even though the performance of the proposed method was found to be satisfactory for the different Trebouxiophyceae and Chlorophyceae microalgal cultures evaluated. Therefore, the proposed Nile Red method should only be used for evaluating lipid content variations in previously standardized strains. BODIPY 505/515 was evaluated as a potential substitute for Nile Red, but was found to be unsuitable as a quantitative stain for algal lipids in the microplate-based staining technique. © 2012 Elsevier B.V. All rights reserved.

1. Introduction Several microalgal species have the capacity to accumulate large amounts of neutral lipids when cultured under different conditions [1,2]. Overproduction and accumulation of neutral lipids in microalgae is of interest as these lipids, mainly triglycerides, can then be converted into biodiesel [2,3]. The ability to rapidly quantify the lipid content in algal cultures is critical for the determination of the optimal time for harvesting. Fast lipid quantification is a key for the economic success of algal biofuels, as the accumulation of oil in algae is the result of the unbalance between growth and oil production rate and changes in the dynamics of the culture may result in a rapid depletion of intracellular oil reserves, affecting process productivity [4,5]. Quantitation of neutral lipids is usually performed by solvent extraction and gravimetry following well-known standard methods [6,7]. These methods, however, are time and labor intensive, and require the use of large sample volumes [8]. For screening purposes and small-scale batch experiments, the use of gravimetric quantitation of lipids is, in general, unsuitable. For instance, when frequent monitoring of lipid content is required over extended periods of time for bench-scale cultures, the total sample volume required for appropriate lipid analysis exceeds the initial culture volume. Several alternative techniques have been proposed in order to both increase the reliability of lipid measurements and reduce the amount of samples required for analysis. Quantitation of total lipids in microalgae has been achieved using fluorometry [9–11], colorimetry ⁎ Corresponding author. Fax: +1 780 492 2881. E-mail address: [email protected] (W.C. McCaffrey). 2211-9264/$ – see front matter © 2012 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.algal.2012.07.004

[12,13], time-domain nuclear magnetic resonance (TD-NMR) [14], in-situ gas chromatography followed by mass spectrometry (GC-MS) [5], flow-cytometry [15], and Raman spectroscopy [16,17]. There exist, however, wide variations among these methods in terms of cost per sample, easiness of preparation, reproducibility, and reliability [18]. Moreover, these alternative methods have not yet been tested for a wider range of strains and conditions, and neither a critical comparison of the advantages and disadvantages for each method has been reported. Consequently, gravimetry continues to be the standard method for monitoring lipid content in microalgae [18]. Fluorescence spectrometry offers a cost-effective alternative to gravimetry for quantifying the lipid content, provided that an appropriate fluorescent marker is available. Nile Red (9-diethylamino-5H-benzo [α]-phenoxazine-5-one) has been widely used as an in-situ lipid marker with the advantage of having strong fluorescence in non-polar environments, while its fluorescence is quenched in water [19,20]. It has been used, with relative success, for measuring the lipid content in microalgae [8,10,11,21]. The use of the Nile Red lipid probe, however, has been hindered by the high variability of the fluorescence measurements, limiting its application to qualitative, and semiquantitative uses [8]. Gocze and Freeman [22] studied the cell to cell variability of fluorescence measurements of lipid droplets in tumor cells, using Nile Red as staining agent. A large variation in lipid vacuole size and distribution among the cell population, as determined by Nile Red staining followed by flow cytometry, led Gocze and Freeman [22] to conclude that such variation was due to the staining agent and not due to an intrinsic difference in lipid content among the cells. When a different dye (4,4-Difluoro-1,3,5,7,8-pentamethyl-4-bora-3A-4A-diaza-S-indacene,

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aka BODIPY 493/503) was used, variability was reduced by 44%. This result suggests that BODIPY has the potential to be a better neutral lipid fluorescent marker than Nile Red. The use of BODIPY 505/515 (4,4-Difluoro-1,3,5,7-tetramethyl-4-bora-3A-4A-diaza-S-indacene) for the staining of lipid bodies in microalgae has more recently been reported by Cooper et al. [23], who used BODIPY 505/515 in combination with fluorescence activated cell sorting (FACS) to detect algal cells with high amounts of oil. In this study, the factors affecting the variability of fluorescence measurements in microalgal cells stained with Nile Red are investigated. The staining and measurement protocol reported by Chen et al. [8] is used as a reference, and modifications to this protocol are suggested in order to reduce variability. The proposed optimized protocol is applied to four different algal strains, representing the classes Chlorophyceae and Trebouxiophyceae: Chlorella vulgaris, Auxenochlorella protothecoides, Scenedesmus dimorphus, and Scenedesmus obliquus. C. vulgaris has been widely studied as a model microalgae, and lipid quantification in C. vulgaris using Nile Red has been previously reported [8,21]. Chen et al. [8] used C. vulgaris as a model strain for evaluating the validity of the microplate based fluorometric method used in this report as a reference. Lipid content in A. protothecoides using Nile Red as an indicator has been reported by Gao et al. [14]. In [14], a correlation coefficient between fluorescence intensity and intracellular lipid concentration equal to r2 =0.9067 was reported, which indicates that the standard Nile Red staining protocol used for Gao et al. [14] can only be used as a semiquantitative method for lipid quantitation due to the large relative error associated with this correlation coefficient. In a modification of their original protocol, Chen et al. [24] pre-treated S. dimorphus cells with microwaves with the aim of facilitating the penetration of the Nile Red dye into the lipid bodies. While it was reported that the use of microwaves in combination with DMSO resulted in increased fluorescence intensity in S. dimorphus [24], it was not reported whether there was a linear relationship between fluorescence intensity count and intracellular neutral lipid content. Quantitation of lipids in S. obliquus using Nile Red as the fluorophore has only been reported in combination with multiparameter flow-cytometry [25], with a relatively low correlation coefficient (r2 =0.80). The modifications to the microplate-based Nile Red staining protocol proposed in this report resulted in a 80–90 % reduction in the relative standard deviation among replicates, allowing the use of Nile Red fluorescence as a quantitative probe. The potential use of BODIPY 505/515 as a staining agent for the quantitative determination of lipid content in microalgae was also investigated. It was found that, even though BODIPY 505/515 provides superior and more specific staining of lipid bodies in microalgal cells, the fluorescence intensity of BODIPY stained cells, measured in a multiwell microplate after in-well staining, is not proportional to the lipid content of the cells. 2. Materials and methods 2.1. Organisms and culture conditions Four species of fresh-water microalgae, representing the classes Chlorophyceae and Trebouxiophyceae, were obtained from the University of Texas Culture Collection of Algae. The cultured strains were Auxenochlorella protothecoides (UTEX 25), Chlorella vulgaris (UTEX 395), Scenedesmus dimorphus (UTEX 746), and Scenedesmus obliquus (UTEX 2630). Algal cells were cultured axenically under heterotrophic conditions in synthetic media containing glucose as the carbon source and glycine as the nitrogen source. Details of media composition and preparation were as previously described by De la Hoz Siegler et al. [4]. To induce lipid accumulation in the cells, the carbon to nitrogen ratio was set at a very high value which is equivalent to a nitrogen depleted condition. The Carbon:Nitrogen (C:N) molar ratio in the cultures varied from 100:1 to 500:1, with the initial glucose concentration ranging from 15

177

to 45 g/L and the initial glycine concentration ranging from 0.2 to 0.6 g/L. The iron content of the media was also varied as iron also has a significant effect on lipid accumulation [26]. The Nitrogen:Iron (N:Fe) molar ratio varied from 15:1 to 185:1, corresponding to an initial iron sulfate concentration of 24–48 mg/L. All cultures were conducted in wide-mouth culture flasks and incubated in the dark, at 25±0.2 °C, in a shaker incubator at 100 rpm. All reagents for growth media preparation were obtained from Fisher Scientific Co. 2.2. Fluorometric measurements with Nile Red The high-throughput technique reported by Chen et al. [8] was followed with the exception that ethanol replaced DMSO as the carrier solvent. The base procedure was as follows: a 10 μL aliquot of a Nile Red ethanolic solution, containing 10 μg/mL of Nile Red, was added to each well of a 96-well microplate containing a fixed volume of algal suspension at a concentration of 5 g/L. The volume in each well was brought to 200 μL by adding an aqueous ethanol solution (20% v/v), and then the plate was incubated at 40 °C for 10 min. Nile Red was obtained from Sigma (N3013). Fluorescence emissions were recorded in a multiplate reader spectrophotometer (Fluoroskan Ascent, Thermo Labsystems), with a dynamic range greater than 6 decades, and a sensitivity of 2 fmol fluorescein per well. A 530 nm excitation filter, with half bandwidth =10 nm, and a 604 nm emission filter, with half bandwidth= 10 nm, were used for the Nile Red fluorescence intensity measurements. 2.2.1. Effect of biomass and solvent concentration In order to determine the conditions that produce a better (more linear) correlation between intracellular oil content and fluorescence intensity count, algal biomass concentration was varied from 0.0 g/L (no biomass) to 20 g/L (cell dry weight), following the staining procedure described above. Similarly ethanol concentration was varied from 0 % (no ethanol, only deionized water) to 30 % v/v. 2.2.2. Effect of staining time and stain concentration Nile Red concentration was evaluated at 1.0 μg/mL and 10μg/mL, and at staining times (length of incubation at 40 °C) of 0, 10, 20, and 30 min. 2.2.3. Effect of mixing To determine if proper mixing took place in the microwells, and to evaluate the effect of mixing on the variability of the fluorometric measurements, all reagents were mixed in a 16 × 150 glass test tube, and vortexed at 3000 rpm. Afterwards, 200 μL aliquots of the pre-mixed suspension were placed in each well of the 96-well microplate, and the microplate was incubated for 10 min at 40 °C. The variability of the fluorescence measurements in this case was compared to the variability when reagents were added individually to each well of the microplate following the procedure indicated above. Further evaluation of the effect of mixing parameters involved the variation of the mixing rate in the microplate reader from 600 rpm to 1200 rpm, and the reduction of the total volume in the wells from 200 μL, to 120 μL. This reduction in volume was achieved by decreasing the amount of aqueous ethanol solution from 180 μL to 100 μL per well. 2.3. BODIPY staining The high-throughput fluorometric technique was further modified by replacing Nile Red as the staining agent with BODIPY 505/515. BODIPY 505/515 was obtained from Invitrogen Molecular Probes (D3921). Measurements for BODIPY stained cells were carried out using a 485 nm excitation filter, half bandwidth = 14 nm; and a 510 nm emission filter, half bandwidth = 10 nm. All fluorescence emission measurements were recorded with a multiplate reader

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spectrophotometer (Fluoroskan Ascent, Thermo Labsystems). BODIPY 505/515 stock solution was as described in Cooper et al. [23].

Fluorescence spectra were recorded using a Photon Technology International (PTI) MP1 Fluorescence System. The excitation spectra of Nile Red staining algal cells were generated at an emission wavelength of 610 nm, while the emission spectra were generated using an excitation wavelength of 530 nm. For BODIPY 505/515, the emission spectra were generated using an excitation filter at 485 nm, as recommended by Cooper et al. [23]. 2.5. Gravimetric determination of neutral lipids Neutral lipids in the algal samples were quantified gravimetrically following the methodological guidelines reported by Shahidi and Wanasundara [27], with conditions optimized in-house. The extraction procedure is briefly summarized as follows. Liquid culture samples of microalgae were centrifuged at 4 °C for 10 min at 21,860 g in a Sorvall SLA-1500 fixed angle rotor. Algal pellets were washed and resuspended in a phosphate buffer solution (pH 6.2) and the procedure was repeated twice. The final algal pellets were frozen at −80 °C, for at least 24 h. Cell pellets were later freeze dried at −50 °C and were high vacuumed for 24 h. Known amounts of freeze dried cells were crushed using a ceramic mortar and pestle, and oil was extracted with a commercial blend of hexanes (Fisher Scientific Co.). Cell debris was separated from the hexane–oil mixture by centrifugation. Oil was recovered by evaporating the hexane and was further quantified gravimetrically. Extracted oil was further characterized by HPLC, in order to evaluate the total lipid classes in the extracted lipid. For this, 5 μL of a 1:10 dilution of the oil sample was added to 200 μL of phosphatidyl N,N-dimethylaminoethanol (PDME) in chloroform (1 μg/μL). 1 μL of the resulting solution was injected into the HPLC, and PDME was used as the internal standard for quantification. HPLC analysis revealed that the hexane-extracted oil was in the form of nearly 100% triglycerides, with traces (below the limit of quantitation) of sterol esters and/or phosphatidylcholine. 3. Results and discussion 3.1. Suitability of Nile Red as an staining agent for the fluorometric quantification of lipids The excitation and emission spectra of two samples of A. protothecoides stained with an ethanolic solution of Nile Red (10 μg/mL) are shown in Fig. 1. The two algal samples differed by their relative intracellular oil content, one sample (oil-rich) containing 50.2±0.9% w/w, and the other (oil-lean) containing 22.8± 0.4 % w/w oil content. The spectra shown in Fig. 1 correspond to the average of five individually collected spectra, where each spectrum was collected right after the previous one without removing the sample from the measuring chamber. This averaging is required in order to reduce the noise inherent to the fluorescence measurements of algal cell suspensions. The noise in the fluorescent signal can be explained by the particulate nature of the sample, and the heterogeneity of cell structures. The average standard deviation among spectra replicates was 0.78%, with a maximum variation of 3.8% with respect to the mean count. In Fig. 1, it is observed that the fluorescence intensity of Nile Red stained algal cells increases with increasing oil content. It is also observed that different peaks exist in the excitation and emission spectra of aqueous algal samples. The different peaks observed in the spectra correspond to differences in the absorption properties of different lipid classes, or other structures, in the cells [19]. Elsey et al. [11] compared the emission spectra of Nile Red dissolved in different solvents, and reported that Nile Red fluorescence maximum emission shifted to higher

Fluorescence intensity counts (a.u.)

2.4. Fluorescence spectra

6

x 105

5

Emission spectra Excitation spectra

4

3

2

1

0 400

450

500

550

600

650

700

Wavelength (nm) Fig. 1. Effect of the intracellular oil content on the excitation and emission spectra of A. protothecoides cells stained with a 10 μg/mL ethanolic solution of Nile Red. The cell concentration, as cell dry weight, in each sample was set at 5 g/L. [‐‐‐] Oil-rich sample (50.2±0.9% w/w oil content, cells were batch-cultured for 200 h with an initial glucose concentration of 30 g/L and an initial glycine concentration of 0.2 g/L); [—] oil-lean sample (22.8±0.4% w/w oil content, cells were batch-cultured for 200 h with an initial glucose concentration of 30 g/L and an initial glycine concentration of 0.6 g/L). The maximum absorption/emission associated with neutral lipids occurs at 527/590 nm. A secondary emission peak, corresponding to a more polar lipid class, occurs at 645 nm.

wavelengths as polarity of the solvent increased, being 576 nm for hexane, 600 nm for chloroform, and 632 nm for ethanol. Therefore, it is expected that different absorption bands will be observed in a cellular sample as it will contain different lipid classes of varying polarity. In Fig. 1, it can be seen that for the oil-lean sample there are two peaks on the emission spectra. The peak at around 590 nm corresponds to the expected emission band for Nile Red in a non-polar environment, while the peak at around 645 nm indicates the presence of a more polar lipid environment [19,11]. This observation supports why Nile Red can be used to quantify neutral lipids in algal samples. The correct selection of excitation and emission filters is, therefore, critical to ensure that non-polar lipids are quantified, rather than other lipid classes or other cellular materials. 3.2. Suitability of BODIPY as a staining agent for the fluorometric quantification of lipids BODIPY 505/515 (4,4-Difluoro-1,3,5,7-tetramethyl-4-bora-3A-4Adiaza-S-indacene) has been employed for in-vivo visualization of lipid bodies in algal cells, and has been used as a dye of choice for fluorescence activated cell sorting aimed at selecting cells with high neutral-lipid content [23]. Given the superiority of BODIPY in visualization applications, compared to Nile Red, it is of interest to evaluate the potential use of BODIPY as a quantitative stain as an alternative to Nile Red in microplate fluorometry. Algal suspensions were stained with a solution of the lipid probe BODIPY 505/515 in DMSO, following the procedure outlined by Cooper et al. [23]. Fluorescence microscopy, as shown in Fig. 2, indicates that the BODIPY 505/515 probe is highly selective for neutral lipids, with a strong fluorescence of the liposomes and a low signal from other cellular structures. The emission spectra of BODIPY stained cells of A. protothecoides were recorded for cells with different oil contents, in a similar way as presented in Fig. 1 for Nile Red. In the emission spectra shown in Fig. 3, there are no significant differences between the spectra of oil rich cells and the spectra of oil lean cells, except for the high intensity auto-fluorescent peak of chlorophyll present in the oil lean cells. Furthermore, when algal suspensions varying in biomass concentrations were stained with equal amounts of BODIPY 505/515 following a

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Fluorescence intensity counts (a.u.)

3.5

179

x 106 BIODIPY emission at 513 nm

3 2.5 Chlorophyll autofluorescence peak

2 1.5 1 0.5 0 500

550

600

650

700

750

800

Wavelength (nm) Fig. 3. Emission spectra of A. protothecoides cells stained with BODIPY 505/515, with excitation at 485 nm. [‐‐‐] Oil-rich sample (50.2 ± 0.9 % w/w oil content); [—] oil-lean sample (22.8 ± 0.4 % w/w oil content), culture conditions were the same as indicated in Fig. 1. The cell concentration, as cell dry weight, in each sample was set at 5 g/L. Note that there was no a significant difference between the emission counts for the two algal samples, except for the autofluorescence peak at 685 nm corresponding to chlorophyll.

Fig. 2. Microphotographs of A. protothecoides, with an average oil content of 43 ± 2 % (w/w), stained with the lipid probe BODIPY 505/515. (a) Bright field image; (b) fluorescence image. The BODIPY dye preferentially stains the liposomes, which exhibit a bright green fluorescence. The red fluorescence observed in one of the cells in the sample is due to chlorophyll autofluorescence. Microphotographs were taken using a Leica DMRXA compound light microscope with a Nikon (DXM 1200) digital camera, a bandpass filter with an excitation range of 450–490 nm, and a longpass suppression filter with an edge wavelength of 515 nm.

microplate high-throughput fluorometric technique, no difference in the total fluorescence count was observed. These results suggest that even though BODIPY 505/515 preferentially binds to neutral lipids, the binding is either not directly proportional to the amount of lipid present in the cells or the fluorescence background of the samples is too high to allow quantification. Govender et al. [28] have recently reported that the fluorescence intensity count of identical microalgal samples stained with varying concentrations of BODIPY 505/515 reached the maximum at a specific concentration of BODIPY after which the fluorescence intensity count decreases, more likely due to an increase in the fluorescence background caused by the dye. Further research is required to determine if it is possible to quantitatively correlate the lipid content of algal cells to the BODIPY fluorescence count at this optimal stain concentration using a high-throughput, microplate-based, fluorometric method. 3.3. Factors affecting the linearity and variability of fluorescence measurements The high-throughput technique proposed by Chen et al. [8] has the advantage of requiring a small sample volume and a short total

analysis time, as several samples can be analyzed at the same time in a multiwell microplate. The use of this technique, however, is limited by the lack of reproducibility of fluorescence measurements and by the non-linear response of fluorescence intensity to oil content in a given sample. Moreover, for certain algal strains even the use of DMSO does not result in an effective distribution of the Nile Red stain in the algal samples [24]. To address the variability in the fluorescence intensity measurements, and to remove the need for the use of a calibration standard, Bertozzini et al. [29] have proposed the use of the standard addition method in combination with Nile Red staining. This modified method requires the serial addition of known amounts of an internal standard (i.e. triolein) to a sample of unknown oil content. It has been successfully applied to Bacillariophyceae and Dinophyceae algal strains. The standard addition method, however, might not be applicable to Chlorophyceae and Trebouxiophyceae algal species as they have thicker cell walls that restrict the diffusion of the Nile Red stain into the cells [8]. Moreover, the use of triolein, might result in over or underestimation of the actual neutral lipid content in the cells due to differences in the relative fluorescence of Nile Red in the different lipid classes [24], as well as differences in the distribution of Nile Red between the triolein globules and the cells. Chen et al. [24] have used microwaves in combination with DMSO to permeate the cells and deliver the Nile Red dye into the lipid bodies, resulting in effective staining, and increased fluorescence intensity, in samples of S. dimorphus, thick-walled Chlorophyceae microalgae. However, the use of dimethyl sulfoxide (DMSO) as a carrier solvent is inconvenient for routine applications due to the suspected toxicity of DMSO [30] and its capacity to carry other chemicals and permeate the skin and other physiological barriers, including the blood–brain barrier. In this work, DMSO is replaced by ethanol as the carrier solvent, as ethanol poses a lower health risk than DMSO while there is only a minor difference in the fluorescence intensity of stained cells using either ethanol or DMSO. Among the different treatments tested by Chen et al. [8], permeation with ethanol ranked second, after DMSO permeation, in terms of total fluorescence intensity count. When samples of C. vulgaris were stained with Nile Red with the assistance of either 20% v/v DMSO or 20% v/v ethanol, Chen et al. [8] reported that the fluorescence intensity count was 430± 15 (standard deviation) for DMSO and 370 ±15 (standard deviation) for ethanol.

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Chen et al. [8] optimized the staining conditions of algal cells based on the fluorescence intensity of the stained cells, allowing the detection of neutral lipids in green algae with thick cell walls. The aim of this report, is to develop a measuring technique that minimizes the relative error associated with the fluorometric estimation of oil content in algal cells, rather than to maximize the total fluorescence intensity. Several factors have been previously identified as affecting the fluorescence intensity response of stained cells, including staining agent concentration, cell concentration, staining temperature, and staining duration. In addition to these factors, the effect of the mixing parameters on fluorescence variability is also investigated in this study. 3.3.1. The effect of biomass and carrier solvent concentration To determine the effect that the cell biomass concentration has over the intensity and variability of fluorescence measurements, aliquots of a suspension containing algal cells with an oil content of 50.2± 0.9 % (w/w), as determined gravimetrically after hexane extraction, were diluted to a variable biomass concentration and stained using a carrier solution with a varying concentration of ethanol. Fluorescence measurements were performed in triplicate, and the results are reported in Fig. 4. In Fig. 4, it is observed that fluorescence intensity increased non-linearly with increasing biomass concentration. Therefore, it is advisable to perform fluorescence measurements at a constant biomass concentration, or to include the effect of biomass concentration in the calibration model. Furthermore, there was a continuous decrease of fluorescence intensity as ethanol concentration increased. This result differs from that reported for DMSO by Chen et al. [8], where fluorescence intensity initially increased with increasing DMSO concentration and reached a maximum at a DMSO concentration of 20–25 % v/v. This maximum was not observed for ethanol. The effect of solvent addition on the correlation coefficient between neutral lipid content and fluorescence intensity was examined by preparing 27 samples of A. protothecoides with varying oil contents, at a constant biomass concentration of 5 g/L, and staining them with an ethanolic Nile Red solution (1.0 μg/mL). The algal samples were prepared by culturing the cells at different C:N:Fe molar ratios in order to achieve different oil contents in the cells. In Fig. 5, it is shown that the use of ethanol improved the correlation coefficient between fluorescence intensity and neutral lipid content, as determined gravimetrically. At the highest tested ethanol concentration (30% v/v) the correlation coefficient was higher while the relative fluorescence intensity was lower.

Fluorescence intensity counts (a.u.)

45 40

0% v/v Ethanol 20% v/v Ethanol

30 25 20 15 10 5 0 0

10

20

30

40

50

60

Neutral lipid content of algal cells (% w/w) Fig. 5. Correlation between Nile-Red fluorescence intensity and neutral lipid content of A. protothecoides cells determined gravimetrically, after hexane extraction. Each sample was prepared to have a cell dry weight equal to 5 g/L, and was stained using a 10 μL aliquot of a Nile Red ethanolic solution, containing 10 μg/mL of Nile Red. [‐‐‐, ○] 0% v/v EtOH, r2 =0.272; [—, □] 20% v/v EtOH, r2 =0.824; [‐ ‐, Δ] 30% v/v EtOH, r2 =0.888; [—, ◊] unstained cells in 20% v/v EtOH, showing that background fluorescence is negligible at the excitation/emission wavelengths used. Algal cells were cultured in batch mode for 200 h, with the initial nutrient concentration ranging as follows: glucose (15–45 g/L), glycine (0.2–0.6 g/L), iron sulfate (24–48 mg/L), total phosphates (2–6 g/L, 30% molar K2HPO4 and 70% molar KH2PO4).

Ethanol, being soluble in both water and lipids, acts as a carrier solvent improving the transport of Nile Red and other compounds into the cells [24,31]. The improved transport of the staining agent linearizes the relationship between neutral lipid content and fluorescence intensity, as observed in Fig. 5. Even though the total fluorescence count decreases with increasing carrier solvent concentration, the use of a 30% v/v ethanol concentration resulted in a higher correlation coefficient. Therefore, ethanol concentration was maintained at 30% v/v for all other measurements and in the recommended staining protocol. In Fig. 5, the background fluorescence due to the algal biomass without Nile Red staining is also reported. It can be observed that at the excitation and emission wavelengths selected (530/604 nm), there is no detectable background fluorescence coming from the unstained algal sample. In Figs. 4 and 5, it can also be observed that at zero biomass concentration, i.e. no algal cells in the sample, there is a residual fluorescence originating from the Nile Red staining. This non-zero fluorescence at zero oil content will result in a non-zero intersect in the calibration line between oil content in the samples and the fluorescence intensity count.

30% v/v Ethanol

35 30 25 20 15 10 5 0 0

35

Fluorescence intensity counts (a.u.)

180

5

10

15

20

Biomass concentration (g/L, dry weight) Fig. 4. Effect of biomass and carrier solvent concentrations on the intensity and variability of fluorescence measurements of Nile Red stained cells of A. protothecoides. Error bars indicate ±2σ, or approximately 95% confidence intervals on the reported measurement, as determined from triplicate samples.

3.3.2. The effect of dye concentration and staining time After identifying the appropriate concentration of carrier solvent (ethanol) it is still necessary to evaluate the effect that other factors have over the correlation between fluorescence intensity and neutral lipid content. For this purpose, the relative standard error for repeated fluorometric measurements (5 replicates) was evaluated at two different Nile Red concentrations (1.0 μg/mL and 10 μg/mL), and for incubation times of 0, 10, 20, and 30 min at 40 °C. As can be seen in Fig. 6, for incubation times shorter than 20 min there were significantly more errors in the measurements made at the lower concentration of Nile Red. Staining time, on the other hand, did not affect the measurement error, except for the longest incubation time. Kinetic characterization of the interactions between Nile Red and microalgal cells has been recently reported by Pick and Rachutin-Zalogin [32]. It was found that there is a sequential transfer of Nile Red from the cellular membrane, where Nile Red interacts with the phospholipids, to the lipid bodies. This sequential transfer results in a time dependency for the fluorescence intensity at a given emission

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4.5

Nile Red conc. = 10 µg/mL Nile Red conc. = 1.0 µg/mL

Relative standard error (%)

4

Table 1 Effect of mixing on the inter-sample variability of oil estimates based on Nile Red fluorescence measurements for a sample of A. protothecoides containing 50.2 ± 0.9 % (w/w) oil content, as determined gravimetrically with hexane extraction.

3.5 3 2.5 2 1.5 1 0.5 0

0

5

10

15

181

20

25

30

Treatment

STD

RSD

MAD

Range

IQR

N

Mix-in-well, 600 rpm, vw = 200 μL Premixed, vw = 200 μL Mix-in-well, 1200 rpm, vw = 200 μL Mix-in-well, 1200 rpm, vw = 120 μL

4.1 2.4 3.2 2.2

8.2 4.9 6.4 4.4

3.2 1.9 2.0 1.6

24.2 12.3 21.7 12.2

5.2 3.3 2.7 2.3

96 96 47 47

Reported values correspond to a dispersion measurement, in percentage (%) on weight basis, around the mean oil content value of 50.2 ± 0.9 % (w/w) for the sample. All measurements were performed on a sample containing 10 g/L of biomass, using ethanol as carrier solvent (30% v/v), and staining with Nile Red at a concentration of 10 μg/mL. STD: sample standard deviation; RSD: relative standard deviation (%); MAD: median absolute deviation, a robust measure of the variability of quantitative data; IQR: interquartile range or mid‐spread, a robust measurement of the statistical dispersion; N: number of replicates; vw: total volume of the sample in the well.

Incubation time (min) Fig. 6. Effect of Nile-Red concentration and incubation time on the relative standard error of fluorescence measurements. Relative standard error was determined for triplicate measurements. For each measurement, a 10 μL aliquot of an A. protothecoides suspension, containing 5 g/L (cell dry weight), was stained with 10 μL of a Nile Red ethanolic solution, and diluted with 180 μL of an aqueous ethanol solution (30% v/v). Plates were incubated at 40 °C.

wavelength. Moreover, as there is also a fluorescence quenching, the combined effect is an optimum time at which fluorescence intensity reaches a maximum value, as reported by Elsey et al. [11] and Chen et al. [8]. The results from Pick and Rachutin-Zalogin [32] indicate that the quenching rate is dependent on the Nile Red concentration, being more pronounced at higher concentrations. Pick and Rachutin-Zalogin [32] hypothesized that the fluorescence quenching is the result of Nile Red oligomerization and redistribution at the lipid/water interface. The increased relative error observed after 30 min of incubation at the higher Nile Red concentration is consistent with the observations by Pick and Rachutin-Zalogin [32], as the quenching is expected to occur faster at the highest Nile Red concentration, and this fluorescence decay does not occur at the same rate in all the cells [32].

volume, from 200 μL to 120 μL, prevented this spillover. The variability for these two datasets was comparable in terms of the fluorescence counts. However, given that volume reduction results in a larger fluorescence count due to a lower dilution, the volume reduction is beneficial as it translates a reduced relative error in the oil measurements, as shown in Table 1. In order to maximize the number of samples per plate, while preserving the error bounds reported in Table 1, it is necessary to determine the minimum number of replicates that statistically will provide a similar variation (e.g. relative standard deviation or IQR) as the variation calculated when 47 or 96 replicates were performed. For this purpose, repeated simulated resampling of the replicate fluorescence measurements, followed by a t-test hypothesis test was performed. When five measurements were selected at random from the original dataset containing either 47 or 96 replicates, the mean and variance of the selected five measurements were statistically equal to the mean and variance of the full dataset, more than 96% of the time. Therefore, by performing only 5 replicate measurements, it is expected to achieve the error bounds presented in Table 1. 3.4. Improved staining protocol for neutral lipid quantification

3.3.3. Mixing effects In addition to evaluate the effect of staining conditions and concentrations on the variability of fluorescence measurements, in this report it is investigated whether appropriate mixing occurs in the microplate wells and if there is any other source of variability. The inter-sample variation in fluorescence count was determined by preparing either a full or a half 96-well microplate with replicates of the same algal sample and measuring the fluorescence count. In Table 1, a statistical analysis of the inter-sample variability is presented. Statistically, the variation in a population is quantified by the standard sample deviation, or by its robust counterpart the median absolute standard deviation (MAD). Another measure of variability is the range, which is the length of the smallest interval containing all the measurements. Given that the range is highly influenced by the presence of an outlier, a robust estimation of the spread is the interquartile range (IQR). Variations in the fluorescence count were found to be due to both mixing effects, and inherent variability of fluorescence readings. Improvement in the mixing of the sample and staining reagents, either by premixing or by modifying mixing parameters caused a reduction in the variability of fluorescence measurements. The sample standard deviation was reduced by 56% when all the reagents were mixed before distributing the sample into the microwells, as shown in Table 1. This implies that 56% of the inter-sample variation was due to volumetric measurement error or mixing effects. By modifying mixing parameters (doubling the rpm) a reduction in both the sample standard deviation and the spread range was observed, as presented in Table 1. When the mixing rate was increased to 1200 rpm, spillover of the content of the microwells was observed. A reduction in the total well

The improved staining protocol is summarized in Fig. 7. As previously indicated, to achieve the error bounds indicated in Table 1, five replicates of each measurement must be performed. After reducing the sources of variability, following the improved staining protocol, a correlation between Nile Red fluorescence measurements and oil content determined gravimetrically after hexane extraction gave a correlation coefficient, r 2, greater than 0.99 as can be observed in Figure S1 in the supplementary materials available on-line. The linearity of the correlation between fluorescence intensity count and neutral lipid content has been verified for a lipid content ranging between 9 and 60% (w/w), which corresponds to the typical range observed in algal cultures. There are, however, no evident reasons to suspect that the linearity will not be preserved at lower or higher values of the range tested. Nonetheless, the linearity in the response should always be verified before accepting a value that falls outside the range at which the calibration curve was generated. Due to variations in the excitation light intensity, caused by lamp aging as well as other sources of day to day variations in the fluorometer, the use of an experimental standard for recalibration is required with each plate that is run. The need for this frequent recalibration can be avoided by using a photon counting spectrofluorometer, which eliminates the effect of variations on light intensity. A periodic recalibration is, nonetheless, recommended to assure that calibration curves remain valid. Freeze-dried algal samples, of known oil content, can be used as standards for recalibration. Freeze-dried algal samples are stable for relatively long-term storage periods, provided they are kept in a dry

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Fig. 7. Recommended staining protocol for the fluorometric quantification of neutral lipids in microalgal suspensions.

and airtight container. Ryckebosch et al. [33] measured the total lipid, free-fatty acid, and carotenoid contents of spray-dried and freeze-dried samples of Phaeodactylum tricornutum, stored at atmospheric pressure or in vacuum and at −20, 4, and 20 °C, and found no significant differences in composition after 14 or 35 days and with respect to storage conditions. In our lab, the neutral lipid content, as determined gravimetrically after hexane extraction, of freeze-dried algal samples that have been stored for up to six months was determined to be the same as the one determined in freshly dried samples. When algal samples were stored for longer times (2–3 years), however, gum was formed during hexane extraction, which reduced the amount of oil extracted. It is recommended, however, to always verify the stability of the standards used for calibration, as conditions will vary for each algal strain. Freeze-dried cells can be resuspended in a sterile phosphate buffer solution to a known biomass concentration (same as the one for the samples of interest). For recalibration purposes a two-point calibration is usually satisfactory. Assuming that 10 wells in a 96-well microplate are used at each time for recalibration (5 replicates are also used for the experimental standards), it is possible to test simultaneously up to 17 algal samples. Based on the reports by Chen et al. [24] and Pick and Rachutin-Zalogin [32], the use of triolein or other lipids as a standard must be avoided, as the Nile Red probe will interact differently with “free” lipid globules and lipid bodies contained inside a rigid cell wall. Moreover, the fluorescence intensity count will be different depending not only on the total neutral lipid content, but also on the lipid composition (i.e. fatty acid profile) of the sample [24]. 3.5. Application of the improved protocol to different microalgal systems To verify the applicability of the improved fluorometric protocol, four different algal strains were cultured at varied C:N molar ratios, under heterotrophic conditions, in order to achieve a varied oil content in the cells. Intracellular oil content was quantified gravimetrically on freeze dried samples, and the fluorescence intensity of Nile Red stained samples was measured following the protocol indicated in Fig. 7. To further verify the robustness of the improved protocol, a technician not previously involved in the protocol development was trained on the use of the staining protocol, and was instructed to perform all the fluorescence measurement for the four algal strains. The correlation

coefficient between the oil content determined gravimetrically and the fluorescence intensity is reported in Table 2 for each one of the four microalgal strains analyzed and for the combined data of all strains. For each algal strain, five to six data-points were generated, each data point were reproduced at least five times. The experimental observations and corresponding calibration curves are reported in Figures S2–S5, as supplementary materials accompanying the on-line version of this article. It must be noted, however, that each curve is specific for the spectrometer, sets of filters, and age of the lamp used, and therefore calibration curves must be generated for each equipment used in order to account for any instrument variation. From the results presented in Table 2, it can be concluded that the improved staining protocol provides a consistent measurement of the fluorescence intensity of stained algal cells, with a relatively low relative standard deviation (less than 5%). Moreover, the correlation coefficient for all individually analyzed algal strains was close to unity, confirming that Nile Red can be used for the quantification of neutral lipids of algal strains. Nonetheless, in order to have confidence in the validity of the results obtained with the proposed fluorometric method, the linearity of the relationship between fluorescence intensity and neutral lipid content should always be verified for every new strain. A final comparison of the proposed improved technique is performed by verifying the relative standard deviation (RSD) achieved in the original high-throughput method proposed by Chen et al. [8] with the RSD of the modified protocol for two different algal strains. In this case, S. dimorphus and A. protothecoides were cultured photoheterotrophically over a period of two weeks, at two different C:N molar ratios. Microalgal suspensions were diluted up to an optical density at 750 nm equal to 0.06, and stained accordingly to either the protocol presented in Fig. 7 or to the original protocol proposed by Chen et al. [8]. Table 2 Correlation coefficients and relative standard deviations (RSD) among replicate measurements for the improved fluorometric protocol as applied to different microalgae strains. Microalgae strain

Correlation coefficient (r2)

Average RSD (%)

Chlorella vulgaris, UTEX 395 Scenedesmus dimorphus, UTEX 746 Scenedesmus obliquus, UTEX 2630 Auxenochlorella protothecoides, UTEX 25

0.989 0.973 0.981 0.989

4.3 1.3 3.7 3.4

H. De la Hoz Siegler et al. / Algal Research 1 (2012) 176–184 Table 3 Relative standard deviations (RSD, %) among replicate measurements for the original high-throughput technique and the improved fluorometric protocol. Microalgae strain

Protocol by Chen et al. [8]

Improved protocol

S. dimorphus, UTEX 746 A. protothecoides, UTEX 25 C. vulgaris

32.6 26.5 3.9–8.5a

4.3 4.1

a RSD values as reported by Chen et al. [8] for repeatability at two different lipid contents, equivalent to 20 and 2 μg/mL.

In Table 3, the relative standard deviation among replicates (8) of fluorometric measurements is reported for both the high-throughput technique proposed by Chen et al. [8] and the improved high-throughput technique here proposed. For both S. dimorphus and A. protothecoides, the modified method provided a relative standard deviation (RSD) six to seven times lower than the RSD in the original method. For comparison purposes, the relative standard deviation as determined by Chen et al. [8] for C. vulgaris is also reported in Table 3. The lowest RSD value reported by Chen et al. [8], 3.9% at a lipid concentration of 20 μg/mL, is consistent with the values found for the improved protocol in both S. dimorphus and A. protothecoides. It must be noticed, however, that the original protocol by Chen et al. [8] was developed and optimized using C. vulgaris as the model organism, and it was later reported that the method failed for S. dimorphus [13]. In the method here proposed, the average RSD ranged from 1.3 to 4.3 for four different strains and at lipid contents between 9 and 60%, which corresponds to a lipid content in the wells of 37–250 μg/mL. 3.6. Use of Nile Red for strain selection or screening A standard calibration curve should be generated for every new strain of interest, given that the fluorescence intensity of stained algal cells is not uniquely determined by the intracellular neutral lipid content. As an illustration, the fluorescence intensity of two algal samples from different species is compared together with their intracellular oil content. A sample of Chlorella vulgaris containing 26±0.5 % (w/w) neutral lipids, as determined gravimetrically after hexane extraction, produced a fluorescence intensity count of 9.9± 0.5 (a.u.) when stained following the protocol outlined in Fig. 7. Following the same staining protocol, a sample of Auxenochlorella protothecoides containing 39.3 ± 0.8 % (w/w) neutral lipids produced a fluorescence intensity count of 9.4 ± 0.3 (a.u.). Another sample of A. protothecoides containing 32.9 ± 0.6 % (w/w) neutral lipids produced a fluorescence intensity count of 7.6 ± 0.1 (a.u.). From the previous data, it is clear that the fluorescence intensity of different algal strains cannot be directly used to compare their relative lipid content, as there are significant differences in the staining capacity of Nile Red among different algal strains. For proper analysis of the results obtained using the high-throughput fluorometric Nile Red method, a standard consisting of an algal sample of known concentration and from the same strain as the analyte must be used. The high-throughput fluorometric Nile Red method should not be used to compare the neutral lipid content of un-standardized algal strains. In particular, the use of the proposed Nile Red fluorometric technique for screening of microalgal strains should be avoided. Once a particular algal strain has been isolated and identified, however, the proposed method can significantly reduce the cost and time required for the screening of optimal process conditions. The proposed method provides a fast and reliable way to quantify intracellular neutral lipids using very small sample volumes. 3.7. Destructive nature of the test The improved staining protocol for lipid quantification is in essence a destructive method, as cell viability after treatment with 30% (v/v) ethanol at 40 °C is very low or zero. To verify cell viability, 1 mL

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suspension of A. protothecoides was incubated for 10 min as described in Fig. 7, and later the cells were recovered by centrifugation at 200 ×g (RCF) for 5 min. As a control, another 1 mL suspension of cells was diluted with deionized water and centrifuged at the same conditions previously described. While the control cells were able to grow at the expected rate, meaning that there was no cellular damage due to the centrifugation, no growth was observed in the stained cells. Given that the proposed method only requires a very small sample volume, however, the destructive nature of the test is not considered critical or limiting for its use. In applications where cell viability is required after testing, the method here proposed shall not be used. The viability of algal cells using BODIPY 505/515 has been reported elsewhere [23], and in combination with fluorescence activated cell sorting it can be used as a suitable screening tool for mutants with high lipid content, where preserving cell viability is of the utmost importance. 4. Conclusion Nile Red can be used as a selective probe for neutral lipids by properly selecting the excitation and emission wavelengths. To allow the use of Nile Red for quantitative analysis the variability in the fluorescence measurements was reduced by tuning the mixing parameters and staining conditions. An improved Nile Red staining protocol that resulted in a significant reduction in the fluorescence intensity variability was proposed. The proposed method offers a low relative standard deviation among replicates and a high degree of linearity. However, its use shall be limited to the monitoring of lipid content in single strain cultures. For comparing the results among different algal strains, standard calibration curves shall be generated for each individual algal strain. The lipid probe BODIPY 505/515 was verified as being selective to neutral lipids, however there was not a quantitative relationship between the fluorescence intensity and the relative lipid content in the sample when analyzed using the proposed microplate-based fluorometric method. Acknowledgments The authors gratefully acknowledge the financial support provided by AVAC (Alberta Value Added Corporation), Natural Sciences and Engineering Research Council of Canada (NSERC), and Alberta Innovates— Technology Futures. The funding agencies did not play any role in the collection, analysis, and interpretation of data; neither in the writing of, or in the decision to publish this report. Appendix A. Supplementary data Supplementary data to this article can be found online at http:// dx.doi.org/10.1016/j.algal.2012.07.004. References [1] F. Chen, M. Johns, Effect of C/N ratio and aeration on the fatty acid composition of heterotrophic Chlorella sorokiniana, Journal of Applied Phycology 3 (1991) 203–209. [2] J. Sheehan, T. Dunahay, J. Benemann, P. Roessler, A look back at the U.S. Department of Energy's aquatic species program—biodiesel from algae, in: Close-Out Report NREL/TP-580-24190, National Renewable Energy Laboratory, 1617 Cole Boulevard Golden, Colorado, 1998, 80401–3393. [3] H. Xu, X. Miao, Q. Wu, High quality biodiesel production from a microalga Chlorella protothecoides by heterotrophic growth in fermenters, Journal of Biotechnology 126 (2006) 499–507. [4] H. De la Hoz Siegler, A. Ben-Zvi, R.E. Burrell, W.C. McCaffrey, The dynamics of heterotrophic algal cultures, Bioresource Technology 102 (2011) 5764–5774. [5] N. Bigelow, W. Hardin, J. Barker, S. Ryken, A. MacRae, R. Cattolico, A comprehensive GC–MS sub-microscale assay for fatty acids and its applications, Journal of the American Oil Chemists' Society 88 (2011) 1329–1338. [6] A. Hara, N.S. Radin, Lipid extraction of tissues with a low-toxicity solvent, Analytical Biochemistry 90 (1978) 420–426.

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