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Liu, Z. J., Liu, L. P., Wen, P., Li, N., Zong, M. H., and Wu, H. (2015). "Effects of acetic acid and pH on the growth and lipid accumulation of the oleaginous yeast Trichosporon fermentans," BioRes. 10(3), 4152-4166.

Abstract

Acetic acid, one major inhibitor released during the hydrolysis of lignocellulosic biomass, can be utilized by the oleaginous yeast Trichosporon fermentans without glucose repression. The effect of acetic acid on the cell growth and lipid accumulation of T. fermentans under controlled pH conditions was investigated in a 5-L fermentor. Undissociated acetic acid with concentrations of 0.026, 0.052, and 0.096 g L-1 in media contributed to approximately 12-, 24-, and 48-h lag phases, respectively, indicating that undissociated acetic acid is the inhibitory molecular form. The inhibition of cell growth was correlated with undissociated acetic acid concentration. However, acetic acid had little influence on the lipid accumulation of T. fermentans at different pH conditions. The specific glucose consumption rate decreased with increasing acetic acid concentration, but the impact of acetic acid on the specific xylose consumption rate was not pronounced. In addition, the variation of pH and acetic acid concentration had no significant influence on the fatty acid composition of the lipids. Acetic acid showed more severe inhibition under low pH conditions. The reduction of intracellular pH partly explains this inhibitory effect.


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Effects of Acetic Acid and pH on the Growth and Lipid Accumulation of the Oleaginous Yeast Trichosporon fermentans

Zong-jun Liu,a, # Li-ping Liu,a,# Peng Wen,b Ning Li, b Min-hua Zong,b,* and Hong Wu b,*

Acetic acid, one major inhibitor released during the hydrolysis of lignocellulosic biomass, can be utilized by the oleaginous yeast Trichosporon fermentans without glucose repression. The effect of acetic acid on the cell growth and lipid accumulation of T. fermentans under controlled pH conditions was investigated in a 5-L fermentor. Undissociated acetic acid with concentrations of 0.026, 0.052, and 0.096 g L-1 in media contributed to approximately 12-, 24-, and 48-h lag phases, respectively, indicating that undissociated acetic acid is the inhibitory molecular form. The inhibition of cell growth was correlated with undissociated acetic acid concentration. However, acetic acid had little influence on the lipid accumulation of T. fermentans at different pH conditions. The specific glucose consumption rate decreased with increasing acetic acid concentration, but the impact of acetic acid on the specific xylose consumption rate was not pronounced. In addition, the variation of pH and acetic acid concentration had no significant influence on the fatty acid composition of the lipids. Acetic acid showed more severe inhibition under low pH conditions. The reduction of intracellular pH partly explains this inhibitory effect.

Keywords: Lignocellulosic biomass; Acetic acid; Lipid production; Intracellular pH; Trichosporon fermentans

Contact information: a: College of Biosciences and Bioengineering, South China University of Technology, Guangzhou 510640, PR China; b: State Key Laboratory of Pulp and Paper Engineering, College of Light Industry and Food Sciences, South China University of Technology, Guangzhou 510640, PR China;

* Corresponding authors: btmhzong@scut.edu.cn; bbhwu@scut.edu.cn;

These authors have the same contribution and are co-first authors.

INTRODUCTION

Biodiesel, as a renewable and green alternative to traditional fossil fuels, has attracted increasing attention in recent years. The most commonly used feedstocks in biodiesel production are waste oils and vegetable oils. However, widespread use of vegetable oils may contribute to a reduction of food supply, and the limited amount of waste oils cannot meet the increasing demand for biodiesel production (Adamczak et al. 2009). Microbial oils, namely singlecell oils (SCOs), produced by oleaginous microorganisms, have long been considered as substitutes for value-added oils with rare fatty acid composition or structure, such as cocoa-butter and those containing polyunsaturated fatty acids (Ratledge 1993; Papanikolaou and Aggelis 2010, 2011). However, in recent years attention has focused on adoption of SCOs as feedstocks for biodiesel production because of their similarity to vegetable oils in fatty acid composition (Li et al. 2008). Presently, although the high cost of the fermentation process (e.g. aeration and agitation) makes SCOs less competitive compared with vegetable oils (Koutinas et al. 2014), their beneficial impacts in the production process, such as non-use of arable land, high productivity, and easy scaling-up, makes them a promising candidate for future utilization. Moreover, the use of renewable and low-cost lignocellulosic biomass for SCOs production seems to be a sustainable development strategy (Huang et al. 2009).

Up to now, the bioconversion of lignocellulosic sugars (mostly xylose, xylose/glucose blends, or lignocellulosic hydrolysates) into SCOs has been extensively studied with various oleaginous microorganisms (Fakas et al. 2009; Economou et al. 2011; Zikou et al. 2012; Ruan et al. 2012, 2015). However, the use of lignocellulosic biomass as feedstock for SCO production still presents many technical hurdles. One challenge is that oleaginous microorganisms should be capable of utilizing mixed sugars and tolerant to inhibitors (Hahn-Hägerdal et al. 2007) because lignocellulosic hydrolyzates contain a variety of sugars (mostly glucose and xylose) and inhibitory compounds.

Acetic acid is one of the major inhibitors released during the solubilization and hydrolysis of hemicellulose (Palmqvist and Hahn-Hägerdal 2000). Generally, the inhibitory effect of acetic acid is attributed to uncoupling (Baronofsky et al. 1984; Herrero et al. 1985; Luli and Strohl 1990) and intracellular anion accumulation (Russell 1991, 1992). The undissociated form of acetic acid (HAc), which is liposoluble, can diffuse across the plasma membrane and dissociate into the anion (Ac) and a proton (H+) in the near-neutral cytosol. The drop in intracellular pH (pHi) caused by the dissociation can be neutralized by the action of the plasma membrane ATPase at the expense of ATP hydrolysis (Verduyn et al. 1992). At high concentrations of acetic acid, the proton pumping capacity of the cell is exhausted, resulting in depletion of the ATP content and acidification of the cytoplasm (Imai and Ohno 1995). In addition to a perturbation of pHi homeostasis, intracellular accumulation of high levels of the acetate anion may give rise to organic acid toxicity (Pampulha et al. 1989).

In a previous study, this group reported that the oleaginous yeast Trichosporon fermentans can use mixed sugars in lignocellulosic hydrolyzates for lipid production. However, T. fermentans gave a poor lipid yield on non-detoxified dilute sulfuric acid-treated rice straw hydrolyzate containing a concentration of acetic acid ranging from 1 to 3 g L-1 (Huang et al. 2009).

Although there have been many works investigating the inhibitory effect of acetic acid on ethanologenic yeasts under anaerobic conditions (Pampulha et al. 1989; Bellissimi et al. 2009; Casey et al. 2010), so far only a few reports have referred to the effect of acetic acid on oleaginous microorganisms under aerobic conditions (Chen et al. 2009; Hu et al. 2009; Huang et al. 2012). Therefore, little is known about this acid’s inhibitory mechanism.

Acetic acid can be also used to synthesize acetyl-CoA (Vorapreeda et al. 2012), which is a two-carbon unit for the synthesis of fatty acid. Hence, in this study, the impacts of acetic acid on the cell growth and lipid accumulation of T. fermentans under controlled pH conditions were examined. To further understand the inhibitory mechanism of acetic acid, the influence of acetic acid on the pHi of T. fermentans was investigated using the fluorescence probe 2′,7′-bis-(2-carboxyethyl)-5-(and-6)-carboxyfluorescein acetoxymethyl ester (BCECF-AM).

EXPERIMENTAL

Yeast Strain and Materials

The oleaginous yeast Trichosporon fermentans CICC 1368 was obtained from the China Center of Industrial Culture Collection and kept on wort agar (130 g L-1 malt extract, 15 g L-1 agar, and 0.1 g L-1 chloramphenicol) at 4 °C. The fluorescent probe BCECF-AM was purchased from Beyotime Biotec. (China). Ionophore nigericin, which was used in the in situ calibration of intracellular pH, was purchased from Enzo Life Sciences Inc. (Switzerland). The 0.22 μm pore size polyether sulfone (PES) filter used for medium filtration sterilization was purchased from Ameritech Scientific Corp. (USA). Other materials and chemicals were purchased from commercial sources at the highest purity available.

Cultivation and Media

Media

The precultivation medium (i.e. YEPD medium) was as follows (g L-1): glucose 10, xylose 10, peptone 10, and yeast extract 10. The fermentation medium contained (g L-1) glucose 20, xylose 20, yeast extract 0.2, peptone 0.7, MgSO4·7H2O 0.4, KH2PO4 2.0, MnSO4·H2O 0.003, and CuSO4·5H2O 0.0001. To investigate the metabolism of acetic acid as sole carbon source by T. fermentans, 1.5 to 30 g L-1 acetic acid were supplemented into the fermentation media instead of glucose and xylose before adjusting the pH to 6.5 by a pH meter (Sartorius, Germany) with 1 M NaOH or 1 M HCl. To avoid a pH change, the medium containing acetic acid was sterilized by filtration through a 0.22 μm pore size PES filter.

Flask cultivation

After 24 h of precultivation at 28 °C and 160 rpm in a rotary shaker, 2.5 mL of seed culture was used to inoculate 50 mL fermentation medium in a 250-mL conical flask. Then, cultivation was performed at 25 °C and 160 rpm (Zhu et al. 2008).

Batch fermentation in 5-L fermentor

The inoculum for the fermentor was prepared by precultivation of T. fermentans in a rotary shaker set at 28 C and 160 rpm in 500-mL flasks containing 100 mL of precultivation medium. Once the cell density of seed culture reached approximately 4.5 g dry cells L-1, the culture was used to inoculate the fermentor to an initial cell density of OD600 = 0.4 to 0.6. Batch fermentation was carried out at 25 C in a 5-L laboratory fermentor (Biostat A Plus Sartorius, Germany) with a working volume of 1.5 L. The concentrations of acetic acid examined were 0 g L-1, 1.5 g L-1 (25 mM), and 3 g L-1 (50 mM). The pH was adjusted to pH 6.5 or 4.5 before inoculation by automatic addition of 1 M NaOH or HCl and kept constant during the fermentation process. Dissolved O2 and pH were monitored with an autoclavable O2 electrode and pH electrode (Hamilton, Switzerland). All fermentations were performed at least in duplicate.

Determination of Biomass, Lipid Content, and Fatty Acid Composition of Lipid

Biomass, lipid content, and fatty acid composition of lipids were determined as described in this group’s previous work (Zhu et al. 2008). Biomass was harvested by centrifugation, and the weight was measured in lyophilized form. Extraction of lipids was carried out according to the modified method of Dyer and Bligh (1959). After extraction of the oil with a mixture of methanol and chloroform (1/2, v/v) for 60 min, the mixture was separated by centrifugation and the chloroform phase was collected and evaporated by a vacuum rotary evaporator (EYELA, Japan) at 100 rpm and 50 °C. The fatty acid profile of lipid was determined by gas chromatography. Lipids were converted to fatty acid methyl esters (FAMEs) according to the method of Morrison and Smith (1964). Then, the FAMEs were determined by gas chromatography (GC-2010) with an ionization detector and a DB-1 capillary column (0.25 cm × 30 m, Agilent Technologies Inc., USA). The column temperature was increased from 170 to 220 °C at a rate of 3 °C min-1 and kept at 220 °C for 3 min. Nitrogen was used as the carrier gas at 0.80 mL min-1. The split ratio was 1:50 (v/v). The injector and the detector temperatures were set at 250 and 280 °C, respectively (Zhu et al. 2008).

Metabolite Analysis

Cell optical density was recorded at 600 nm with a spectrophotometer (Shimadzu UV-2550, Japan). The supernatant obtained following centrifugation of culture sample was analyzed for glucose, xylose, xylitol, and acetic acid via HPLC with a Waters 2410 refractive-index detector and an Aminex HPX-87H column (Bio Rad Corp., USA) at 50 °C. The mobile phase was 0.005 M H2SO4 at 0.5 mL min-1(Huang et al. 2009).

Determination of Intracellular pH Change

Cell loading

Cells grown in YEPD were diluted to an OD600 of approximately 0.8, then centrifuged at 1500 g for 5 min and resuspended in an equal volume of 100 mM citric/phosphate buffer (potassium dihydrogen orthophosphate and citric acid) at pH 4.5. BCECF-AM is a fluorescent substance which displays pH-dependent excitation, thus allowing the implementation of pHi measurement. Here, it was added at a final concentration of 5 μM, and then in the loading process yeast cells were incubated at 28 °C for 1 h.

Calibration curve and determination of intracellular pH change

Loaded cells were diluted with citric/phosphate buffer at pH values between 6.0 and 8.0 (increments of 0.5 pH units) to OD600 of approximately 0.4. To equilibrate the pHi (intracellular pH) with the controlled extracellular media, H+/Kionophore nigericin was added at a concentration of 30 μM in the presence of 100 mM potassium in citric/phosphate buffer, and incubated for 60 min.

Fluorescence measurements were acquired on a SpectraMax M5 (Molecular Devices, USA) fluorescence spectrophotometer using 96-well black microplates (Corning, USA) with a working volume of 200 μL. A calibration curve was established for BCECF-AM by plotting the ratio of fluorescence intensities (emission wavelength 530 nm) at the pH-dependent excitation wavelengths of 490 and 440 nm as a function of pHi. It was essential to determine the levels of fluorescence in the culture supernatant separately to remove the background fluorescence interference. Thus, for each sample, total fluorescence was examined, followed by a fluorescence determination of the supernatant only (cells were removed by filtering through 0.22-μm filters).

To examine the effect of acetic acid on the pHof T. fermentans, acetic acid was added to the culture at a buffered pH of 4.5 and an OD600 of 0.4 with a final concentration of acetic acid of 50 mM (3.0 g L-1). The control was made using the same procedure without addition of acetic acid. Fluorescence determinations were made as described previously. The pHchange was calculated based on the ratio of fluorescence intensities and the calibration curve.

To confirm the distribution and the level of the loaded BCECF-AM within the cytosol of the yeast cells, loaded cells were examined with a confocal laser scanning microscope (CLSM). The cells were visualized using a Leica TCS SP5 CLSM (Leica, Germany) equipped with a 100-mW argon laser and objective magnification of 100× (Leica 100× oil). Split screen images were obtained using the dual-channel collection mode. One channel was an illumination phase-contrast image, and the other channel was an epi-fluorescent image of intracellular BCECF-AM (Bracey et al. 1998).

Statistical Analysis

All the experiments were performed in duplicate, and their average values with standard deviations were used for statistical analysis. SPSS software (Version17.0, Chicago, USA) were used to analyze the resulting data. One-way analysis of variance (ANOVA) and Turkey’s Honestly Significant Differences (HSD) Test were used to determine the significant differences of data at a 95% confidence interval.

RESULTS AND DISCUSSION

Metabolism of Acetic Acid by T. fermentans

To understand the role of acetic acid in the fermentation by oleaginous yeast T. fermentans, acetic acid was used as the sole carbon source for a culture of T. fermentans. As shown in Fig. 1, in the concentration range of 1.5 g L-1 to 30 g L-1, acetic acid could be assimilated by T. fermentans. When 20 g L-1 acetic acid was used as the sole carbon source, the biomass and lipid content were 5.82 g L-1and 34.1%, respectively, after shake-flask cultivation for 60 h at an initial pH of 6.5.

Fig. 1. Utilization of various concentrations of acetic acid by T. fermentans in shake-flask cultivation. Cultures were incubated at initial pH 6.5, 25 °C, and 160 rpm. Data are the mean and standard error of two duplicate fermentations.

Interestingly, the initial utilization rate of acetic acid did not correlate with the concentration, remaining at approximately 0.1 g L-1 h-1 within the concentration range tested. This result indicates that acetic acid can be metabolized by T. fermentans without substrate inhibition. It was also reported that Cryptococcus curvatus and Yarrowia lipolytica could use acetic acid as substrate (or co-substrate) for efficient SCO production (Christophe et al. 2012; Fontanille et al. 2012). As shown in Fig. 2, T. fermentans can metabolize acetic acid and glucose simultaneously at a pH of 6.5, suggesting that there is no glucose repression and that the enzymes responsible for acetic acid metabolism are non-inducible. This behavior is similar to Zygosaccharomyces bailii in the acetic acid metabolic pathway (Sousa et al. 1998).

Fig. 2. Time course profile for the fermentation of glucose and acetic acid by T. fermentans in shake-flask cultivation. Cultures were incubated at initial pH 6.5, 25 °C, and 160 rpm. Data are the mean and standard error of two duplicate fermentations.

Effect of Acetic Acid and pH on Cell Growth and Lipid Accumulation

Acetic acid showed more serious inhibition on ethanologenic yeast under low pH conditions (Pampulha et al. 1989), indicating that the inhibition of acetic acid is correlated with culture pH. To study the combined effects of acetic acid and pH on the cell growth and lipid accumulation of T. fermentans, three concentrations of acetic acid (0, 1.5, and 3 g L-1) and two levels of pH (6.5 and 4.5) were tested for their influence on biomass, sugar utilization rate, and lipid yield. The time course profiles for the fermentation are shown in Fig. 3, and corresponding results are summarized in Table 1.

As can be seen in Fig. 3 and Table 1, the inhibition of acetic acid on the cell growth of T. fermentans is closely related to the concentration of undissociated acetic acid. The undissociated acetic acid concentration was related to the culture pH. Increasing the concentration of undissociated acid prolonged the lag phase, with 0.026, 0.052, and 0.096 g L-1 in culture media contributing to approximately 12, 24, and 48 h lag phases, respectively. Less than 0.052 g L-1 of undissociated acid did not affect the growth of T. fermentans heavily, allowing glucose and acetic acid to be consumed simultaneously. When the undissociated acetic acid concentration was as high as 1.92 g L-1, the growth of T. fermentans ceased, suggesting that the undissociated acetic acid is the inhibitory molecular form.

Fig. 3. Time course profiles for the co-fermentation of glucose and xylose by T. fermentans under different acetic acid concentrations and pH values. (a) 0 g L-1 acetic acid, pH 6.5; (b) 0 g L-1 acetic acid, pH 4.5; (c) 1.5 g L-1 acetic acid, pH 6.5; (d) 1.5 g L-1 acetic acid, pH 4.5; (e) 3 g L-1 acetic acid, pH 6.5; (f) 3 g L-1 acetic acid, pH 4.5. ● Glucose; ■ xylose; □ biomass; ○ lipid; △ acetic acid; ◊ OD600. Data are the mean and standard error of two duplicate fermentations.

Interestingly, varying the concentration of acetic acid and pH had little influence on the lipid accumulation of T. fermentans. Specifically, at pH 6.5 a greater lipid content of 57% in comparison to the control of 51% was obtained for T. fermentans in the presence of low concentrations of acetic acid (1.5 g L-1). Lipid yield (gram lipid per gram sugar) under these conditions (pH 6.5, 1.5 g L-1 acetic acid) was also a little greater than that of the control (0.22 vs. 0.19). One possible reason is that the acetic acid may be involved in the lipid synthesis because acetate ions can be used in the formation of acetyl-CoA via the action of acetyl-CoA synthetase, which is subsequently applied in the fatty acid synthesis.

It was reported by Sousa et al. that the specific enzyme activity of acetyl-CoA synthetase increased when the spoilage yeast Zygosaccharomyces bailii was cultivated in media containing acetic acid (Sousa et al. 1998). Vorapreeda et al. also found that acetic acid is one of the acetyl-CoA sources for fatty acid synthesis (Vorapreeda et al. 2012).

Table 1. Summary of Results Obtained from Co-fermentation of Glucose and Xylose by T. fermentans under Varying pH Values and Acetic Acid Concentrations

N.D.= Not Detected.

Values shown are the mean and standard error of two duplicate fermentations for each condition. Specific growth rate was calculated by dividing 0.693 by doubling time of the cell concentration. Specific sugar consumption rate was calculated by dividing the slope of steepest portion of sugar consumption curve by the average cell concentration over that period. The concentrations of undissociated acid were calculated according to Henderson–Hasselbalch equation, using a pKa of 4.75 for acetic acid, the pH value, and total acid concentration for each fermentation condition.

The fatty acid compositions of the lipids obtained under different pH and acetic acid concentrations are summarized in Table 2. The majority of the fatty acids present were oleic acid, palmitic acid, linoleic acid, and stearic acid, with oleic acid accounting for about 60% of the total fatty acids. The variation of pH and acetic acid concentrations had no significant influence on the fatty acid composition of lipids (p>0.05). The possible reason is that the activities of key enzymes responsible for lipid synthesis were not seriously affected during fermentation. However, the related mechanism needs to be further investigated in our ongoing research. It is worth noting that a low pH was beneficial for oleic acid synthesis, while the synthesis of palmitic acid was stimulated by acetic acid at both pH conditions.

Effect of Acetic Acid and pH on Glucose and Xylose Consumption

Generally, under nitrogen-limited condition, glucose is metabolized by oleaginous microorganisms through the glycolysis pathway and tricarboxylic acid cycle to generate large amounts of acetyl-CoA, which is then used for subsequent fatty acid synthesis. The metabolism of xylose into acetyl-CoA was considered to have two pathways: the pentose phosphate pathway (1 molar xylose generates 1.67 molar acetyl-CoA) and the phosphoketolase reaction (1 molar xylose generates 2 molar acetyl-CoA) (Evans and Ratledge 1984; Fakas et al. 2009). As depicted in Fig. 3, T. fermentans can assimilate glucose and xylose simultaneously. A similar phenomenon was also observed when glucose and xylose were used as co-substrate for lipid production by Thamnidium elegans (Zikou et al. 2012).

Table 2. Effect of Acetic Acid and pH on the Fatty Acid Composition of Lipid from T. fermentans

N.D.= Not Detected

Values shown are the mean and standard error of two duplicate fermentations for each condition.

As shown in Table 1, at each pH condition the specific glucose consumption rate decreased with an increase in acetic acid concentration. No acceleration was observed in the glucose consumption rate at low acetic acid concentrations and pH 6.5. In contrast, under such conditions the glucose consumption rate of the ethanologenic yeast Saccharomyces cerevisiae increased (Pampulha et al. 1989). As for xylose, the specific xylose consumption rate declined slightly as the acetic acid concentration increased at pH 6.5. When the pH was maintained at 4.5, the specific xylose consumption rates in the presence of 0 g L-1 and 1.5 g L-1 acetic acid were similar. There was no xylose and glucose consumption at 3 g L-1 acetic acid, which explains the cessation of cell growth under this condition. Interestingly, the lipid yield of SCO produced per mixture of sugars consumed at most of the conditions was quite high (0.19 to 0.24 g/g), which is higher than or comparable to the one achieved by other oleaginous yeast and fungi on blends of xylose and glucose (Yu et al. 2014; Fakas et al. 2009), indicating that T. fermentans is a robust strain for SCO production with lignocellulosic sugars as substrate.

Previous works on ethanologenic yeasts found that undissociated acid led to a more severe inhibition of xylose metabolism than glucose consumption (Casey et al. 2010; Hasunuma et al. 2011). For ethanol fermentation, the inhibition of xylose consumption by acetic acid may be linked to a lower capacity of ATP generation during xylose metabolism under anaerobic conditions (Bellissimi et al. 2009; Casey et al. 2010) that cannot meet the ATP requirement for expelling excess protons and anionic species. However, in this study, the specific xylose consumption rate was not reduced substantially by the addition of acetic acid at pH 6.5 or 4.5, probably because of the higher ATP regeneration rate in aerobic fermentation. It is worth noting that the concentration of xylitol detected in the fermentation broth was less than 0.01 g L-1, indicating that acetic acid did not influence the stoichiometry of the xylose metabolic pathway. In contrast, some yeasts utilize xylose via xylose reductase (XR) and xylitol dehydrogenase (XDH), thus increasing the acetic acid concentration in the culture medium and favoring the production of xylitol (Felipe et al. 1995; Helle et al. 2004).

In addition, low concentrations of acetic acid have been shown to stimulate ethanologenic yeasts’ glucose consumption rate to accelerate ATP regeneration under anaerobic conditions (Pampulha et al. 2006; Keating et al. 2006). However, no increase in the glucose consumption rate was observed with the addition of acetic acid in this study. This may be attributed to the different energy-producing route in aerobic fermentation, which does not merely rely on the glycolytic pathway.

Effect of Acetic Acid on Intracellular pH

To better understand why 3 g L-1 acetic acid caused serious inhibition on cell growth and lipid accumulation of T. fermentans at pH 4.5, the effect of acetic acid on the pHi of T. fermentans cells was investigated using fluorescent probe BCECF-AM. As described previously, the ratio value of fluorescence intensity between 490 and 440 nm was plotted as a function of pHi to establish a calibration curve,

Ratio value (x) = (tF490bF490) / (tF440bF440) (1)

where tF490 and tF440 are the total fluorescence intensities at 490 and 440 nm, respectively, and bF490and bF440 are the background fluorescence intensities at 490 and 440 nm, respectively.

As shown in Fig. 4, the calibration curve of the ratio values of fluorescence intensities of BCECF-AM against intracellular pH was established. A polynomial function can be fitted to calculate pHi (y) from the ratio value (x). The equation of best fit to the calibration curve was:

= –0.1457 x2 + 1.4714 x + 4.2734 (2)

The regression coefficient (R2) for this equation was 0.9768.

T. fermentans cells in the exponential phase were transferred to media containing no acetic acid (the control) and 50 mM (3 g L-1) acetic acid, respectively, in pH 4.5 citric/phosphate buffer. Unlike 31P-NMR or the distribution of radio labelled weak acid technique, BCECF-AM analysis is a non-invasive in vivo method that is based on the pH-dependent fluorescence of BCECF-AM in the cytosol of the cells. It was reported that loading of the fungus Neurospora crassa with BCECF-AM resulted in accumulation of the fluorescent indicator in vacuoles instead of even distribution throughout the cytosol (Slayman et al. 1994). Hence, it is essential to confirm the level of fluorescent dye loading within the cells and the distribution of the loaded indicator within the cytosol of T. fermentans cells. As shown in Fig. 5, the loaded cells were visually analyzed using confocal laser scanning microscopy. Most cells showed high levels of intracellular fluorescence, which appeared to be in an even cytoplasmic distribution. As indicated in Fig. 6, the initial intracellular pH before addition of acetic acid was approximately 6.25, similar to the result of approximately 6.2 measured with the fluorescent probe carboxyfluorescein diacetate succinimidyl ester (CFDA-SE) (Bracey et al. 1998). After adding acetic acid, the intracellular pH of T. fermentans cells declined to about 5.7. This pHi change is similar to the result determined by Roe et al. (1998) by approximately 0.4 pH units (Roe et al. 1998). The severe inhibition of acetic acid at low pH on lipid production is partly attributed to the reduction of the intracellular pH of T. fermentans.

Fig. 4. Calibration curve of BCECF-AM in citric/phosphate buffer with nigericin-treated cells of T. fermentans. Data are the mean and standard error of two duplicate fermentations.

Fig. 5.Images of T. fermentans cells loaded with BCECF-AM taken using confocal laser scanning microscopy: (left) an illumination phase contrast image; (middle) the corresponding fluorescent image; (right) the fluorescent image super-imposed onto a phase image.

Fig. 6. Intracellular pH of T. fermentans with or without acetic acid in 100 mM citric/phosphate buffer, pH 4.5. Solid circle, the control without acetic acid; open circle, addition of 50 mM (3 g L-1) acetic acid. The dotted line and arrow show the time of addition. Data are the mean and standard error of two duplicate fermentations.

CONCLUSIONS

  1. Oleaginous yeast T. fermentans was able to metabolize acetic acid without glucose repression.
  2. The effect of acetic acid on the cell growth and lipid accumulation of T. fermentans was related to the culture pH, and undissociated acetic acid is the inhibitory molecular form. The reduction of intracellular pH partly explains the inhibitory effect of acetic acid.
  3. The change of pH and acetic acid concentration had no significant influence on the fatty acid composition of the lipids.

ACKNOWLEDGMENTS

We acknowledge the Fundamental Research Funds for the Central Universities (2014ZZ0048), the Science and Technology Project of Guangdong Province (2013B010404005), the New Century Excellent Talents in University (Grant No. NCET-11-0161), the National Key Basic Research Program of China (2013CB733500), and the State Key Program of National Natural Science Foundation of China (21336002) for financial support.

REFERENCES CITED

Adamczak, M., Bornscheuer, U. T., and Bednarski, W. (2009). “The application of biotechnological methods for the synthesis of biodiesel,” Eur. J. Lipid. Sci. Tech. 111(8), 800-813. DOI: 10.1002/ejlt.200900078

Baronofsky, J. J., Schreurs, W. J., and Kashket, E. R. (1984). “Uncoupling by acetic acid limits growth of and acetogenesis by Clostridium thermoaceticum,” Appl. Environ. Microbiol. 48(6), 1134-1139.

Bellissimi, E., Van Dijken, J. P., Pronk, J. T., and Van Maris, A. J. A. (2009). “Effects of acetic acid on the kinetics of xylose fermentation by an engineered, xylose-isomerase-based Saccharomyces cerevisiae strain,” FEMS Yeast Res. 9(3), 358-364. DOI: 10.1111/j.1567-1364.2009.00487.x

Bligh, E. G., and Dyer, W. J. (1959). “A rapid method of total lipid extraction and purification,” Can. J. Chem. Physiol. 37(8), 911-917. DOI: 10.1139/y59-099

Bracey, D., Holyoak, C. D., Nebe-von Caron, G., and Coote, P. J. (1998). “Determination of the intracellular pHi of growing cells of Saccharomyces cerevisiae: the effect of reduced-expression of the membrane H+-ATPase,” J. Microbiol. Meth. 31(3), 113-125. DOI: 10.1016/s0167-7012(97)00095-x

Casey, E., Sedlak, M., Ho, N. W. Y., and Mosier, N. S. (2010). “Effect of acetic acid and pH on the cofermentation of glucose and xylose to ethanol by a genetically engineered strain of Saccharomyces cerevisiae,” FEMS Yeast Res. 10(4), 385-393. DOI: 10.1111/j.1567-1364.2010.00623.x

Chen, X., Li, Z. H., Zhang, X. X., Hu, F. X., Ryu, D. D., and Bao, J. (2009). “Screening of oleaginous yeast strains tolerant to lignocellulose degradation compounds,” Appl. Biochem. Biotech. 159(3), 591-604. DOI: 10.1007/s12010-008-8491-x

Christophe, G., Deo, J. L., Kumar, V., Nouaille, R., Fontanille, P., and Larroche, C. (2012). “Production of oils from acetic acid by the oleaginous yeast Cryptococcus curvatus,” Appl. Biochem. Biotech. 167, 1270-1279. DOI: 10.1007/s12010-011-9507-5

Economou, Ch. N., Aggelis, G., Pavlou, S., and Vayenas, D. V. (2011). “Single cell oil production from rice hulls hydrolysate,” Bioresour. Technol. 102, 9737-9742. DOI: 10.1016/j.biortech.2011.08.025

Evans, C. T., and Ratledge, C. (1984). “Induction of xylulose-5-phosphate phosphoketolase in a variety of yeasts grown on D-xylose: the key to efficient xylose metabolism,” Arch. Microbiol. 139, 48-52. DOI: 10.1007/bf00692711

Fakas, S., Papanikolaou, S., Batsos, A., Galiotou-Panayotou, M., Mallouchos, A., and Aggelis, G. (2009). “Evaluating renewable carbon sources as substrates for single cell oil production by Cunninghamella echinulata and Mortierella isabellina,” Biomass Bioenerg. 33, 573-580. DOI: 10.1016/j.biombioe.2008.09.006

Felipe, M. G., Vieira, D. C., Vitolo, M., Silva, S. S., Roberto, I. C., and Manchilha, I. M. (1995). “Effect of acetic acid on xylose fermentation to xylitol by Candida guilliermondii,” J. Basic. Microb.35(3), 171-177. DOI: 10.1002/jobm.3620350309

Fontanille, P., Kumar, V., Christophe, G., Nouaille, R., and Christian, C. (2012). “Bioconversion of volatile fatty acids into lipids by the oleaginous yeast Yarrowia lipolytica,” Bioresour. Technol. 114, 443-449. DOI: 10.1016/j.biortech.2012.02.091

Hahn-Hägerdal, B., Karhumaa, K., Fonseca, C., Spencer-Martins, I., and Gorwa-Grauslund, M. F. (2007). “Towards industrial pentose-fermenting yeast strains,” Appl. Microbiol. Biot. 74(5), 937-953. DOI: 10.1007/s00253-006-0827-2

Hasunuma, T., Sanda, T., Yamada, R., Yoshimura, K., Ishii, J., and Kondo, A. (2011). “Metabolic pathway engineering based on metabolomics confers acetic and formic acid tolerance to a recombinant xylose-fermenting strain of Saccharomyces cerevisiae,” Microb. Cell. Fact. 10, 2. DOI: 10.1186/1475-2859-10-2

Helle, S. S., Murray, A., Lam, J., Cameron, D. R., and Duff, S. (2004). “Xylose fermentation by genetically modified Saccharomyces cerevisiae 259ST in spent sulfite liquor,” Bioresour. Technol.92(2), 163-171. DOI: 10.1016/j.biortech.2003.08.011

Herrero, A. A., Gomez, R. F., Snedecor, B., Tolman, C. J., and Roberts, M. F. (1985). “Growth inhibition of Clostridium thermocellum by carboxylic acids: A mechanism based on uncoupling by weak acids,” Appl. Microbiol. Biot. 22(1), 53-62. DOI: 10.1007/bf00252157

Huang, C., Zong, M. H., Wu, H., and Liu, Q. P. (2009). “Microbial oil production from rice straw hydrolysate by Trichosporon fermentans,” Bioresour. Technol. 100(19), 4535-4538. DOI: 10.1016/j.biortech.2009.04.022

Huang, C., Wu, H., Liu, Z. J., Cai, J., Lou, W. Y., and Zong, M. H. (2012). “Effect of organic acids on the growth and lipid accumulation of oleaginous yeast Trichosporon fermentans,” Biotechnol. Biofuels5, 4. DOI: 10.1186/1754-6834-5-4

Hu, C. M., Zhao, X., Zhao, J., Wu, S. G., and Zhao, Z. B. (2009). “Effects of biomass hydrolysis by-products on oleaginous yeast Rhodosporidium toruloides,” Bioresour. Technol. 100(20), 4843-4847. DOI: 10.1016/j.biortech.2009.04.041

Imai, T., and Ohno, T. (1995). “The relationship between viability and intracellular pH in the yeast Saccharomyces cerevisiae,” Appl. Environ. Microbiol. 61(10), 3604-3608.

Keating, J. D., Panganiban, C., and Mansfield, S. D. (2006). “Tolerance and adaptation of ethanologenic yeasts to lignocellulosic inhibitory compounds,” Biotechnol. Bioeng. 93(6), 1196-1206. DOI: 10.1002/bit.20838

Koutinas, A. A., Chatzifragkou, A., Kopsahelis, N., Papanikolaou, S., and Kookos, I. K. (2014). “Design and techno-economic evaluation of microbial oil production as a renewable resource for biodiesel and oleochemical production,” Fuel 116, 566-577. DOI: 10.1016/j.fuel.2013.08.045

Li, Q., Du, W., and Liu, D. H. (2008). “Perspectives of microbial oils for biodiesel production,” Appl. Microbiol. Biot. 80(5), 749-756. DOI: 10.1007/s00253-008-1625-9

Luli, G. W., and Strohl, W. R. (1990). “Comparison of growth, acetate production, and acetate inhibition of Escherichia coli strains in batch and fed-batch fermentations,” Appl. Environ. Microbiol.56(4), 1004-1011.

Morrison, W. R., and Smith L. M. (1964). “Preparation of fatty acid methyl esters and dimethyl acetals from lipids with boron trifluride–methanol,” J. Lipid Res. 5, 600-608.

Palmqvist, E., and Hahn-Hägerdal, B. (2000). “Fermentation of lignocellulosic hydrolysates. II: Inhibitors and mechanisms of inhibition,” Bioresour. Technol. 74(1), 25-33. DOI: 10.1016/S0960-8524(99)00161-3

Pampulha, M. E., and Loureiro-Dias, M. C. (1989). “Combined effect of acetic acid, pH and ethanol on intracellular pH of fermenting yeast,” Appl. Microbiol. Biot. 31(5), 547-550. DOI: 10.1007/bf00270792

Pampulha, M. E., and Loureiro-Dias, M. C. (2006). “Energetics of the effect of acetic acid on growth of Saccharomyces cerevisiae,” FEMS Microbiol. Lett. 184(1), 69-72. DOI: 10.1016/S0378-1097(00)00022-7

Papanikolaou, S., and Aggelis, G. (2010). “Yarrowia lipolytica: A model microorganism used for the production of tailor-made lipids,” Eur. J. Lipid Sci. Technol. 112, 639-654. DOI: 10.1002/ejlt.200900197

Papanikolaou, S., and Aggelis, G. (2011). “Lipids of oleaginous yeasts. Part II: Technology and potential applications,” Eur. J. Lipid Sci. Technol. 113, 1052-1073. DOI: 10.1002/ejlt.201100015

Ratledge, C. (1993). “Single cell oils – have they a biotechnological future?” Trends Biotechnol. 11, 278-284. DOI: 10.1016/0167-7799(93)90015-2

Roe, A. J., McLaggan, D., Davidson, I., O’Byrne, C., and Booth, I. R. (1998). “Perturbation of anion balance during inhibition of growth of Escherichia coli by weak acids,” J. Bacteriol. 180(4), 767-772.

Ruan, Z. H., Zanotti, M., Wang, X. Q., Ducey, C., and Liu, Y. (2012). “Evaluation of lipid accumulation from lignocellulosic sugars by Mortierella isabellina for biodiesel production,” Bioresour. Technol. 110, 198-205. DOI: 10.1016/j.biortech.2012.01.053

Ruan, Z. H., Hollinshead, W., Isaguirre, C., Tang, Y. J. J., Liao, W., and Liu, Y. (2015). “Effects of inhibitory compounds in lignocellulosic hydrolysates on Mortierella isabellina growth and carbon utilization,” Bioresour. Technol. 183, 18-24. DOI: 10.1016/j.biortech.2015.02.026

Russell, J. B. (1991). “Intracellular pH of acid-tolerant ruminal bacteria,” Appl. Environ. Microbiol.57(11), 3383-3384.

Russell, J. B. (1992). “Another explanation for the toxicity of fermentation acids at low pH: Anion accumulation versus uncoupling,” J. Appl. Microbiol. 73(5), 363-370. DOI: 10.1111/j.1365-2672.1992.tb04990.x

Slayman, C. L., Moussatos, V. V., and Webb, W. W. (1994). “Endosomal accumulation of pH indicator dyes delivered as acetoxymethyl esters,” J. Exp. Biol. 196(1), 419-438.

Sousa, M. J., Rodrigues, F., Côrte-Real, M., and Leão, C. (1998). “Mechanisms underlying the transport and intracellular metabolism of acetic acid in the presence of glucose in the yeast Zygosaccharomyces bailii,” Microbiology 144(3), 665-70. DOI: 10.1099/00221287-144-3-665

Verduyn, C., Postma, E., Scheffers, W. A., and Van Dijken, J. P. (1992). “Effect of benzoic acid on metabolic fluxes in yeasts: A continuous-culture study on the regulation of respiration and alcoholic fermentation,” Yeast 8(7), 501-517. DOI: 10.1002/yea.320080703

Vorapreeda, T., Thammarongtham, C., Cheevadhanarak, S., and Laoteng, K. (2012). “Alternative routes of acetyl-CoA synthesis identified by comparative genomic analysis: Involvement in the lipid production of oleaginous yeast and fungi,” Microbiology 158(1), 217-228. DOI: 10.1099/mic.0.051946-0

Yu, X. C., Zheng, Y. B., Xiong, X. C., and Chen, S. L. (2014). “Co-utilization of glucose, xylose and cellobiose by the oleaginous yeast Cryptococcus curvatus,” Biomass Bioenerg. 71, 340-349. DOI: 10.1016/j.biombioe.2014.09.023

Zhu, L. Y., Zong, M. H., and Wu, H. (2008). “Efficient lipid production with Trichosporon fermentansand its use for biodiesel preparation,” Bioresour. Technol. 99(16), 7881-7885. DOI: 10.1016/j.biortech.2008.02.033

Zikou, E., Chatzifragkou, A., Koutinas, A. A., and Papanikolaou, S. (2012). “Evaluating glucose and xylose as cosubstrates for lipid accumulation and γ-linolenic acid biosynthesis of Thamnidium elegans,” J. Appl. Microbiol. 114, 1020-1032. DOI: 10.1111/jam.12116

Article submitted: February 2, 2015; Peer review completed: April 12, 2015; Revised version received and accepted: May 15, 2015; Published: May 21, 2015.

DOI: 10.15376/biores.10.3.4152-4166