You-sheng Wang, Beijing Higher Institution Engineering Research Center of Food Additives and Ingredients, Beijing Technology and Business University, Beijing 100048, China. E-mail: firstname.lastname@example.org
Aim: To provide the observation that sodium citrate induced apoptosis in biocontrol yeast Cryptococcus laurentii.
Methods and Results: The viability of the yeast cells was evaluated using the percentage of colony-forming units (CFU) of treated cells. The induction of cell death was dependent on the concentration of sodium citrate and exhibited typical apoptotic markers such as phosphatidylserine (PS) translocation as shown by annexin V coupled with fluorescein isothiocyanate (FITC) labelling and DNA fragmentation as detected by TdT-mediated dUTP-biotin nick end labelling (TUNEL) assay. The annexin V-positive cells reached the maximum (14·8%) on the third day, whereas TUNEL-positive cells increased gradually from 5·92 to 27·9% within 5 days of incubation in sodium citrate. In addition, confocal laser microscopy and flow cytometric analysis revealed that the induction of apoptosis was associated with the production of reactive oxygen species (ROS) that reached the highest intracellular level in the first day, before the peak of the early event (PS exposure) in apoptosis. The apoptosis was delayed by the addition of antioxidant glutathione (GSH), suggesting that ROS generated in this process plays a key role in the regulation of the apoptosis in C. laurentii cells.
Conclusions: This study indicated that the apoptotic signals in C. laurentii are dependent on citrate ions and/or sodium ions, the concentration and initial acidity of sodium citrate. Induction of ROS in response to sodium citrate plays a significant role in apoptosis.
Significance and Impact of the Study: Yeast Cryptococcus laurentii has been selected as an effective biocontrol indicator for the postharvest diseases because of its competition for nutrients and space with the pathogen in the wound of fruits. This study presents a convenient method for commercial production of yeast as biocontrol agent.
Yeast has been studied as an alternative biological approach to synthetic fungicides for controlling postharvest decay because wounds are the main entry through which pathogens attack fruits (Wilson and Wisniewski 1989; Filonow 1998). The yeast Cryptococcus laurentii has been shown to display a significant inhibitory activity on the main diseases of pear (Roberts 1990), peach (Zhang et al. 2007) and sweet cherry (Wang and Tian 2008). In addition, this yeast showed its abilities to colonize on fruit surfaces under field conditions and adapt to postharvest storage conditions of low temperature, low O2 and high CO2 concentrations (Tian et al. 2004), and to serve as antagonistic yeast when used together with some exogenous substances such as ammonium molybdate (Wan et al. 2003), sodium bicarbonate (Yao et al. 2004) and low amount of preservatives (Qin et al. 2004).
The main obstacle for commercialization of yeast as biocontrol agents is the formulation of shelf-stable product that retains activity after storage (Janisiewicz and Jeffers 1997). Torres et al. (2003) reported that the liquid product of antagonist Candida sake using 10% lactose as protectant retains >70% activity after storage at 0°C for 3 months, and no significant difference was observed between fresh and stored products. However, Granot and Snyder (1991) observed glucose-induced death of the yeast Saccharomyces cerevisiae, indicating that not all carbon sources could be used as protectant for yeast. In addition, organic acids such as acetic acid (Ludovico et al. 2001), valproic acid (Mitsui et al. 2005) and formic acid (Du et al. 2008) could also induce apoptosis-like cell death in yeast. Citric acid, which is often in the form of sodium citrate, is the major organic acid of fruits and the most abundant in berries citrus and tropical fruits (Belitz and Grosch 1987). Until now, no work on the liquid form of antagonist has been carried out, especially the influence of sodium citrate on the viability of biocontrol yeast C. laurentii.
Yeast apoptosis was initially described using temperature-sensitive cdc48 mutant of S. cerevisiae cultured at the appropriate restrictive temperature (Madeo et al. 1997). The expression of the human pro-apoptotic protein Bax in yeast led to accumulation of reactive oxygen species (ROS) and an apoptotic phenotype that could be alleviated by trapping free radicals and anaerobiosis (Perrone et al. 2008). In addition, the presence of wounds and fungal development constitutes another stress source that produces lots of ROS (Bolwell and Wojtaszek 1997). However, there is no previous report on the direct role of ROS during the yeast apoptosis induced by sodium citrate.
The object of this study is to investigate the influence of sodium citrate on the viability of C. laurentii and the role of ROS, to provide insight into the production of liquid formulated biocontrol yeast.
Materials and methods
Yeast strain and growth conditions
The biocontrol yeast, Cryptococcus laurentii, was obtained from Chinese Center for Industrial Culture Collection. The yeast was grown in 250-ml conical flasks containing 50 ml of yeast extract, peptone and dextrose (YPD: 10 g of yeast extract, 20 g of peptone and 20 g of dextrose in 1000 ml water) and incubated in an orbital shaker at 25°C for 48 h. Yeast cells were harvested by centrifugation at 4000 g for 10 min and resuspended in sterile deionized water.
Cryptococcus laurentii cells were grown in YPD broth. Briefly, 200 ml of medium (in 500 ml Erlenmeyer flasks) was inoculated with C. laurentii cells from fresh YPD plate cultures and incubated in an orbital shaker at 28°C for 48 h. Cells were then collected, washed three times with sterile deionized water and resuspended in 200 ml of the treatment medium at 2 × 107 cells ml−1. Yeast cells were resuspended in sodium citrate (100 mmol l−1, pH 5·8) with or without 6·7 g l−1 yeast nitrogen base (YNB) with amino acids and ammonium sulfate. To determine the influence of concentration and initial acidity, the C. laurentii cells were incubated in treatment media containing 25 mmol l−1 sodium citrate at pH 5·8 or 100 mmol l−1 sodium citrate at pH 3·8. To investigate the effect of citrate and sodium ion, yeast cells were also resuspended in sterile deionized water with 50 mmol l−1 sodium citrate plus 50 mmol l−1 sodium phosphate at pH 5·8 or 100 mmol l−1 potassium citrate at pH 5·8. To verify the influence of antioxidant, the C. laurentii cells were resuspended in sodium citrate (100 mmol l−1, pH 5·8) with 10 mmol l−1 glutathione (GSH). All the treated cells were cultured at 25°C for 5 days with shaking.
Growth and viability assays
Yeast cells were diluted with water to proper concentration and spread on YPD plates. Cell viability was determined by plating serial dilutions of the treated yeast cells onto YPD plates. The percentage of colony-forming units (CFU) of treated cells was obtained by relating the CFU counts of treated cells to those of cells at time point 0, which were considered to be 100%. The survival rates were expressed as mean values with standard error of mean (SEM) of at least three independent experiments.
Analysis of apoptotic markers
Cells were stained using the annexin-V-FITC kit (Bender Co., San Diego, CA, USA) and propidium iodide (PI) according to manufacturer’s instructions with minor modifications (Du et al. 2008). Briefly, cells were washed once with PBS and then incubated in annexin binding buffer containing 5 μl ml−1 annexin reagent and 20 μg ml−1 PI for 20 min in dark. For confocal laser microscopy, cells were concentrated by a short centrifugation step. Analysis was performed using an LSM 510-META confocal laser scanning microscope (Carl Zeiss, Jena, Germany). For detection of annexin V fluorescence signals, a 505–550-nm band-pass emission filter was used with an excitation at 488 nm (argon laser). PI fluorescence (excitation at 543 nm, HeNe laser) was ascertained using a 585-nm long-pass emission filter. To avoid crosstalk between the fluorescence channels in the case of co-staining, probes were scanned sequentially.
TdT-mediated dUTP-biotin nick end labelling (TUNEL) staining was performed as described by Kitagaki et al. (2007). Briefly, harvested cells were fixed with 3·7%(v/v) formaldehyde at 4°C for 1 h in 200 μl of sorbitol buffer, digested with 15 U lyticase at 28°C for 2 h, washed twice with PBS, applied to poly-lysin-coated glass slide, permeablized with 0·1% Triton-X 100 and 0·1% sodium citrate on ice for 2 min, washed once with PBS, stained with TUNEL reaction mixture and observed under Axio Imager.A1 fluorescence microscope (Carl Zeiss).
Detection of reactive oxygen species
Intracellular ROS were detected using the oxidant-sensitive probe 2′,7′-dichlorodihydrofluorescein diacetate (H2DCF-DA, Molecular Probes) (Balzan et al. 2004). Cells were harvested by centrifugation at 3290 g for 10 min, resuspended in 10 μg ml−1 H2DCF-DA that was diluted from a stock of 2·5 mg ml−1 in methanol and incubated for 2 h in dark, washed three times with sterile deionized water and incubated in 50 mmol l−1 sodium citrate solution containing 5 μg ml−1 PI for 20 min in the dark.
For confocal laser microscopy, cells were concentrated by a short centrifugation step. Analysis was acquired using an LSM 510-META confocal laser scanning microscope (Carl Zeiss). For detection of H2DCF-DA fluorescence signals, a 505–550-nm band-pass emission filter was used with an excitation at 488 nm (argon laser). PI fluorescence (excitation 543 nm, HeNe laser) was analysed using a 585-nm long-pass emission filter. To avoid crosstalk between the fluorescence channels in the case of co-staining, probes were scanned sequentially.
For flow cytometric analysis, cells from different treatments were resuspended in 500 μl of phosphate-buffered saline, and 10 000 cells were probed. Analysis was performed using the BD FACSCalibur and Summit ver. 3.1 software (Becton, Dickinson & Co., Franklin Lakes, NJ). H2DCF-DA fluorescence signals were determined using a 510–550-nm band-pass filter; for PI, a 610–650-nm band-pass filter was used (excitation at 488 nm, argon laser).
Results and analysis
Cell death of Cryptococcus laurentii induced by sodium citrate
The yeast C. laurentii grew rapidly in the presence of sodium citrate (100 mmol l−1, pH 5·8) plus YNB and reached its peak at 48 h, as shown by the typical growth curve of yeast cells (Fig. 1). However, cells underwent rapid death and reached below 1% of the initial concentration after 72 h when incubated in sodium citrate at 100 mmol l−1, pH 5·8).
In contrast to pH 5·8, C. laurentii grew quickly when incubated in 100 mmol l−1 sodium citrate at pH 3·8, indicating that the low pH prevented the yeast cells from death (Fig. 1). Similarly, when the salt concentration decreased from 100 to 25 mmol l−1, the yeast cells apparently grew normally. These results indicated that the cell death was dependent on the concentration and initial acidity of sodium citrate.
Inhibitory effects of the cell growth by sodium citrate alone (100 mmol l−1, pH 5·8) were reversed by incubation in 50 mmol l−1 sodium citrate plus 50 mmol l−1 PBS or by use of potassium citrate instead of sodium citrate. In contrast, cell growth incubated in 100 mmol l−1 PBS remained at 88–158% of initial level within 5 days. The results indicated that both citrate ions and sodium ions could induce cell death.
Cell death induced by sodium citrate (100 mmol l−1, pH 5·8) was delayed by GSH, suggesting that ROS may play an important role in the cell death induced by sodium citrate.
PS exposure examined in Cryptococcus laurentii by annexin V
PS is distributed asymmetrically in the lipid bilayer of the plasma membrane in yeast and mammalian cells (Phillips et al. 2003; Fannjiang et al. 2004; Yang et al. 2006). Translocation of PS from the inner leaflet to the extracellular side of the plasma membrane is an early event in apoptosis and can be examined by annexin V, a protein with strong affinity to PS (Martin et al. 1995; Madeo et al. 1997). The C. laurentii cells treated with 100 mmol l−1 sodium citrate at pH 5·8 were green in colour, indicating that the cells were stained by FITC-coupled annexin V but not by PI, demonstrating the intactness of the plasma membrane of the apoptotic (a) protoplasts. Viable (v) cells could not be stained by either FITC-coupled annexin V or PI. Yeast cells in the late apoptotic stages (l) stain green and red because of the inability of the cell membrane to exclude PI. In contrast, the PI-positive PI cells (n) may represent cells that became secondary necrotic or died by an unknown mechanism (Fig. 2a, sodium citrate). No annexin V staining was observed in protoplasts obtained from control cells treated with water (Fig. 2a, water).
It can be seen that the annexin V-positive cells exposed to 100 mmol l−1 sodium citrate at pH 5·8 reached a peak value of approximately 14·8% on the third day (Fig. 2b, sodium citrate). In contrast, annexin V-positive cells incubated in water showed weak staining in the cell lumen or no detectable fluorescence (Fig. 2b, water).
Nuclear changes associated with cell death in Cryptococcus laurentii
DNA damage was detected by TUNEL assay for the apoptotic cells (Madeo et al. 1997; Herker et al. 2004). After 3-day culture in 100 mmol l−1 sodium citrate liquid medium at pH 5·8, yeast cells became TUNEL-positive and 16·3% of the cells exhibited an intense black nuclear staining (Fig. 3a, sodium citrate and 3b). The TUNEL-positive cells suggested that condensed fragmented chromatin occurred as the consequence of apoptosis in C. laurentii cells.
In addition, the percentage of TUNEL-positive sodium citrate-treated cells showed that DNA fragmentation in these cells was time dependent. It was found that cells with DNA cleavage increased from 5·92% in 1-day cultures to 27·9% in 5-day cultures (Fig. 3b). In contrast, water had no/little effect on DNA damage of cells (Fig. 3a,b).
Dying cells produce reactive oxygen species
The appearance of apoptotic markers in yeast is accompanied by the production of ROS (Madeo et al. 2002). Some cells in the C. laurentii suspensions exposed to sodium citrate (100 mmol l−1, pH 5·8) displayed significant green fluorescence staining with H2DCF-DA and red fluorescence labelled with PI after 3 days (Fig. 4a, sodium citrate). In contrast, no detectable H2DCF-DA fluorescence was observed in the cells incubated in water (Fig. 4a, Water).
Fluorescence-activated cell sorting measurement of the number of ROS-positive sodium citrate (100 mmol l−1, pH 5·8)-treated cells was 65·0%. Parallel assessments of these cells with PI indicated a much smaller increase (only 11·7%) in the number of cells that were stained with both H2DCF-DA and PI after 3 days (Fig. 4b). The number of ROS-producing cells exposed to sodium citrate decreased after a transient increase (80·1%) on the first day, reaching a minimum (40·1%) on the fifth day (Fig. 4c). In contrast, water-treated cells did not show a significant increase in the amount of ROS; only 0·15–1·5% water-treated cells had increased ROS amount.
Granot and Snyder (1991) reported that the exposure of stationary-phase cells to sugar in the absence of additional nutrients caused loss of viability within a few hours and cell death, a phenomenon termed to be sugar-induced cell death (SICD). Our present data show that sodium citrate at 100 mmol l−1, pH 5·8 plus YNB stimulated growth of yeast C. laurentii, but induced cell death in the absence of YNB (Fig. 1). Our results seem to support the suggestion that the exposure of stationary-phase cells to a stimulating nutrient, in the absence of additional nutrients to support growth, will force the cells out of the stationary phase (Granot and Synder 1993). However, further study in our experiment showed that the induction of apoptosis in C. laurentii by sodium citrate depends on the culture acidity and concentration of sodium citrate. Yeast cells grew normally when incubated in 100 mmol l−1 sodium citrate at pH 3·8 or 25 mmol l−1 sodium citrate at pH 5·8 (Fig. 1), supporting the observation that the exposure of stationary-phase cells to carbon stimulates the growth of C. laurentii. Thus, the phenomenon, as observed by Torres et al. (2003), that survival rate of the liquid product of antagonist Candida sake was more than 70% after storage at 0°C for 3 months using 10% lactose as protectant might be due to the slow growth during storage.
This study showed that the sodium citrate (100 mmol l−1, pH 5·8) induced cell death in biocontrol yeast C. laurentii cells and this process exhibits typical markers of apoptosis-like PS flipping (Fig. 2a) and DNA fragmentation (Fig. 3a), which were also the landmarks of apoptosis in S. cerevisiae induced by acetic acid (Ludovico et al. 2001) and Candida albicans induced by amphotericin B (Phillips et al. 2003). However, the present study showed that the maximum number of annexin V-positive cells, which often serves as a sensitive marker for early stages of apoptosis (Martin et al. 1995; Madeo et al. 1997), was observed at the third day, whereas the percentage of intense TUNEL-positive cells increased from 3- to 5-day cultures. Thus, it could be deduced that the DNA damage (TUNEL staining) was time dependent and sodium citrate could induce the progression of yeast cells from early apoptosis to late or necrotic apoptosis. On the other hand, apoptosis has been defined as a highly regulated form of programmed cell death (PCD) in yeast cells (Gourlay et al. 2006). PCD might play an important physiological role in the self-destruction of virus-infected, damaged or old cells. The self-destruction may consume dwindling nutrients and contribute to the viability and reproductive success of healthier members of the community harbouring similar genomes (Büttner et al. 2007). Our results suggested that the altruistic apoptotic death induced by sodium citrate (100 mmol l−1, pH 5·8) could promote the regrowth of a subpopulation of better-adapted mutants (data not shown). Consequently, sodium citrate-induced PCD might be a new research field to discover biocontrol yeast that can tolerate limited nutrients against pathogen attack.
It has been reported that the apoptosis in S. cerevisiae was induced by both low concentration of H2O2 and deletion of GSH1, during which an accumulation of ROS was observed (Madeo et al. 1999). The present results also revealed that the number of ROS-producing cells was much more than that of yeast cells stained with annexin V (Fig. 2b) and TUNEL (Fig. 3b) on the first day, indicating that the production of intracellular ROS might initiate the apoptotic programs in C. laurentii cells induced by sodium citrate. This can be further supported by the observations that the addition of GSH, an effective antioxidant that can be absorbed by the cell, could delay the viability loss of C. laurentii (Fig. 1), and that much less ROS production was detected in C. laurentii cells which can survive in water for 5 days (Fig. 4c). These results were consistent with the observations of Longo et al. (1997) and Laun et al. (2001), who agreed with the fact that ROS induced and regulated apoptotic death of yeast cells. Similar result was also presented by Ludovico et al. (2001, 2002), who observed that acetic acid induced apoptosis in S. cerevisiae via ROS generation. However, the mechanism(s) by which sodium citrate induced the production of ROS needs further study.
In conclusion, this study indicated that the apoptotic signals in C. laurentii are dependent on citrate ions and/or sodium ions, the concentration and initial acidity of sodium citrate. ROS generated in response to sodium citrate play significant roles in apoptosis.
This work was supported by the National Natural Science Foundation of China (30901009) and the grants from Beijing Nova Programme (2007B011) and Beijing Natural Science Foundation (6122003).