Metabolic studies of γ-polyglutamic acid production in Bacillus licheniformis by small-scale continuous cultivations

Biochemical Engineering Journal 73 (2013) 29–37

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Metabolic studies of ␥-polyglutamic acid production in Bacillus licheniformis by small-scale continuous cultivations Anja Wilming a , Jens Begemann a , Stefan Kuhne a , Lars Regestein a , Johannes Bongaerts b , Stefan Evers b , Karl-Heinz Maurer b,1 , Jochen Büchs a,∗ a b

Aachener Verfahrenstechnik – Biochemical Engineering, RWTH Aachen University, Sammelbau Biologie, Worringer Weg 1, 52074 Aachen, Germany Henkel KGaA, Henkelstraße 67, 40191 Düsseldorf, Germany

a r t i c l e

i n f o

Article history: Received 2 September 2012 Received in revised form 17 December 2012 Accepted 13 January 2013 Available online 19 January 2013 Keywords: Bioreactors Viscosity Fermentation Protease Bacillus licheniformis Polyglutamic acid

a b s t r a c t For the cultivation of microorganisms shake flasks are frequently used. The development of the broth viscosity during the cultivation is usually not paid special attention to. However, changes in viscosity may have a severe impact on shake flask cultivations, ranging from effects on mass and heat transfer to the occurrence of out-of-phase conditions. Significant changes in viscosity were observed during batch cultivations with Bacillus licheniformis on defined mineral medium. The formed biopolymer was identified to be ␥-polyglutamic acid (␥-PGA), a polymer, Bacillus species are known to excrete as a side-product. ␥-PGA leads to increasing viscosity and, thus, has critical influences on shake flask cultivations. Consequently, the oxygen transfer is strongly influenced by viscosity and, therefore, ␥-PGA formation. Furthermore, ␥-PGA has chelating characteristics, which might affect nutrient and trace element availability, possibly resulting in substrate limitations. In total, these influences lead to undefined conditions in the fermentation. Therefore, the trigger for ␥-PGA production was investigated. By performing continuous cultivation experiments as well as pulse experiments, catabolite controlled overflow was identified as one of the key triggers for ␥-PGA production. The results suggest, that catabolite controlled overflow leads to inhibition of the 2-oxoglutarate complex, resulting in accumulation of 2-oxoglutarate and a redirection of metabolic flux toward glutamic acid and finally to ␥-PGA. Additionally, oxygen limitations were determined to further potentiate ␥-PGA production. As a result, fermentation conditions favoring catabolite controlled overflow as well as oxygen limitation need to be strictly avoided in order to assure defined conditions at low viscosity throughout the fermentation. © 2013 Elsevier B.V. All rights reserved.

1. Introduction Metabolic studies of Bacillus licheniformis gain importance because derivatives of the gram-positive, spore forming soil bacterium are industrially used for the bulk production of enzymes [1,2]. As B. licheniformis, such as other Bacillus species, is able to secrete large amounts of extracellular enzymes [2], they are heavily used for protease production in detergent industry. Especially proteases of the subtilisin family are produced and used in detergents nowadays. In 2002, proteases accounted for roughly 40% of total enzyme sales in various industrial market sectors [3]. Just within the European Union, about 900 tons of pure enzyme were produced and used in 2002 [2]. During fermentation processes of Bacillus strains, high amounts of ␥-polyglutamic acid (␥-PGA) can

∗ Corresponding author. Tel.: +49 241 80 24633; fax: +49 241 80 22570. E-mail address: [email protected] (J. Büchs). 1 Present address: AB Enzymes GmbH, Feldbergstraße 78, 64292 Darmstadt, Germany. 1369-703X/$ – see front matter © 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.bej.2013.01.008

be formed as an undesired side-product [4]. As the polymeric ␥PGA leads to an increase in broth viscosity, it may cause severe problems during fermentation. Consequently, conditions favoring high viscosities should be avoided for a defined and well-controlled process resulting in high productivity. In order to achieve this, the production mechanism of ␥-PGA must be well understood. Despite these challenges, B. licheniformis is nowadays frequently used for the production of ␥-polyglutamic acid (␥-PGA) [5,6]. ␥PGA is a water soluble, anionic, non-toxic polymer used in drug release [7], as moisturizing agent in cosmetics, and as thickener, cryoprotectant or aging inhibitor in food [4]. Its versatile applications make it interesting for the industrial use. Several groups have investigated the formation of ␥-PGA in much detail [8–14]. The most frequently used method for the ␥-PGA production in lab scale is the one first published by Leonard [15]. The used medium E contains glycerol, citrate, as well as glutamic acid as carbon and energy sources in high concentrations (80 g/L, 12 g/L and 20 g/L, respectively). High ␥-PGA product concentrations (∼12 g/L–23 g/L) were reported when using the medium E in shake flasks [10,11] and fermenter [12]. Cromwick et al. [12] for example, investigated ␥-PGA

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Nomenclature  d0 n D cS YO/S

growth rate (h−1 ) shaking diameter (m) shaking frequency (rpm) dilution rate (1/h) amount of substrate metabolized by the cells (mmol/L) yield coefficient (mol oxygen/mol substrate)

formation for B. licheniformis ATCC9945A using medium E in a pHcontrolled fermenter. An increased ␥-PGA formation was observed for cultures at pH 6.5 and high aeration (800 rpm, 2 vvm) in contrast to low aeration (250 rpm, 0.5 vvm). In the 1980s, Frankena et al. [16,17] reported that citric acid acts as carbon catabolite repressor already in a concentration of 3.8 g/L in B. licheniformis. Therefore, the above-mentioned studies were most probably performed under carbon catabolite repression (CCR) as the medium E contains three times more citrate. Dauner et al. investigated the influence of carbon-limiting and excess-carbon conditions on the metabolism of the close relative Bacillus subtilis [18]. While performing continuous cultivation experiments, Dauner et al. [18,19] observed a highly active tricarboxylic acid (TCA) cycle under carbon-limiting conditions. Furthermore, they observed a decrease in TCA activity with an increasing dilution rate. Low dilution rates result in high level of carbon limitation and, therefore, no carbon catabolite repression. Due to accumulation of carbon source, higher dilution rates close to washout indicate carbon catabolite repression. Therefore, the observed decrease in TCA activity with increasing dilution rate is probably connected to carbon catabolite repression. In addition to continuous cultivation experiments, batch experiments were performed with a high initial carbon concentration. The TCA activity observed was close to complete repression due to very strong CCR. Moreover, Thorne [13] observed a 2-oxoglutarate accumulation in B. subtilis cultures producing ␥-PGA, which is an obvious indication that 2-oxoglutarate acts as precursor for ␥-PGA. In 1973, Troy et al. [8,9] performed a comprehensive characterization of the biosynthesis as well as a characterization of structural aspects of the B. licheniformis ␥-PGA in batch experiments with the above mentioned medium E. During these studies, 2-oxoglutarate was confirmed as precursor for ␥-PGA. As a result to the high carbon concentration within the medium, especially of citrate, carbon catabolite repression most probably prevailed until citrate was completely metabolized. Thus, carbon catabolite repression is likely to be connected to the observed repression of the TCA, and results in a catabolite controlled accumulation of 2-oxoglutarate at high carbon concentrations, here termed catabolite controlled overflow. The observations made by Cromwick et al. [10–12], Dauner et al. [18,19] and Troy et al. [8,9] led to the hypothesis that catabolite controlled overflow caused by CCR in B. licheniformis results in redirection of metabolic fluxes to the production of ␥PGA. In Escherichia coli, oxygen limitation has a similar effect on the TCA as observed for CCR and catabolite controlled overflow in Bacillus [20]. Consequently, also the influence of oxygen limitation on ␥-PGA production was investigated in batch and continuous cultures. For batch experiments, the Respiration Activity Monitoring System (RAMOS) was used. This system enables online monitoring of oxygen transfer rate (OTR), carbon dioxide transfer rate (CTR) and respiration quotient (RQ) by measuring the oxygen partial pressure in the gas phase of specially modified Erlenmeyer flasks (RAMOS flask) [21,22].

Continuous cultivation strategies and pulse experiments were applied in this work since they provide for steady-state conditions, simplifying the deduction of kinetic parameters [23–24]. In contrast to batch experiments media composition, viscosity as well as the metabolic cell state do not vary over time. By introducing pulses to a carbon-limited continuous steady-state culture, the impact of different stress signals on the cells can be studied [25]. In order to perform continuous cultivation and pulse experiments, the continuous parallel small scale shaken bioreactor system (COSBIOS) developed by Akguen et al. [23] was used. For further details refer to the work of Akguen et al. [23,24]. Especially in shake flask cultivations, the change in medium viscosity by produced biopolymers such as ␥-PGA can have significant influences. High viscosities not only affect heat and mass transfer but also power input, coalescence, bulk mixing and process control in stirred tank reactors [26]. In addition, the hydromechanical conditions in the flask are influenced. An increase in viscosity can lead to the so-called out of phase phenomenon. As a consequence, high viscosities may result in a breakdown of liquid rotation and, therefore, in a breakdown of oxygen transfer [27]. Because the viscosity has such a severe influence on fermentation processes this paper aims at elucidating the trigger for ␥-PGA production as an undesired side-product in the industrially highly relevant protease production by B. licheniformis. 2. Materials and methods 2.1. Strains and media The protease producing B. licheniformis strain and a ␥-PGA deficient mutant were kindly provided by Henkel KGaA (Düsseldorf, Germany). All chemicals were purchased from Carl Roth GmbH & Co. KG (Karlsruhe, Germany), Sigma–Aldrich Chemie GmbH (Taufkirchen, Germany), or Fluka (Buchs, Switzerland) and were of analytical grade. The modified V3 MOPS medium for continuous cultures contained per liter: glucose 20 g, MOPS – acid 52.3 g (0.25 mol), MnCl2 ·4H2 O 0.05 g, CoCl2 ·6H2 O 0.53 mg, ZnCl2 0.26 mg, H3 BO3 0.01 mg, NiSO4 ·6H2 O 0.66 mg, CuSO4 ·5H2 O 0.31 mg, Na2 MoO4 ·2H2 O 0.65 mg, MgSO4 ·7H2 O 1.01 g, CaCl2 ·H2 O 0.026 g, FeSO4 ·7H2 O 0.05 g, (NH4 )2 SO4 7.0 g, K2 HPO4 3.4 g, Na3 citrate 3.8 g, kanamycin 50 mg. Every component was added separately from sterile stock solutions in the order of appearance. Na3 citrate was used as chelating agent to prevent precipitation and to enable unhindered feeding through the pump hoses of the COSBIOS system. The initial pH was set to 7.5 with 5 M NaOH. The V3 MOPS medium for batch cultivation was identical to the modified V3 MOPS medium except for the buffer concentration 41.85 g/L (0.2 M), the (NH4 )2 SO4 concentration of 15 g/L, and initial pH = 8.0. No Na3 citrate was used as chelating agent in the batch medium. 2.2. Cultivations Cultivations and pulse experiments were performed in duplicates which essentially showed the same result. For simplicity, only one data set is presented in this work. A two-stage preculture was used to inoculate both batch and continuous main cultures. The first preculture was inoculated from a glycerol stock and cultivated over night (250 mL Erlenmeyer flask, cotton plug, shaking frequency 350 rpm, shaking diameter 50 mm, filling volume 10 mL, 4% (v/v) inoculum, 37 ◦ C) in LB medium [28,29]. The second preculture in LB medium was inoculated from the first preculture with an initial OD0 = 0.4 and was run until the cells reached mid-exponential growth. Monitoring of the second preculture was realized using the RAMOS device (same conditions as described above). For main batch cultures a master mix was inoculated from

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the second preculture with an initial OD0 = 0.4 and the desired volume transferred to the different RAMOS flasks. For these experiments a RAMOS device made in house was utilized. Commercial versions are available from Kühner (Birsfelden, Switzerland) and Hitech Zang (Herzogenrath, Germany). Samples were drawn from ordinary shake flasks (250 mL Erlenmeyer flask, cotton plug) cultivated in parallel under identical conditions than the RAMOS flasks. RAMOS flasks and sample flasks were inoculated from the same master mix. Therefore, it is reasonable to assume that cultures run synchronously. This technique was successfully applied in previously published projects [22,30]. For continuous cultures the COSBIOS was used, equipped with six parallel bioreactors. For details on system assembly please refer to Akguen et al. [23,24]. To pump the feed medium, a multiperistaltic pump (IPC-N-8, Ismatech, Glattbrug, Switzerland) was used. Thereby, every reactor could be operated at a distinct dilution rate, or if desired, with identical ones. For the main culture, the corresponding reactor volume for the chosen shaking parameters (as described by Akguen et al. [23]) was transferred to the COSBIOS flasks from an inoculated master mix using a sterile syringe (OD0 = 0.4, filling volume 15.5 mL at 350 rpm, 25 mL at 200 rpm, shaking diameter 50 mm, 37 ◦ C). After a batch phase of 24 h, the reactor system was switched to continuous mode by starting of the multiperistaltic pump. Samples were drawn from each reactor after reaching steadystate conditions by using the inoculation port and a sterile syringe. Glucose and citric acid concentrations were determined by isocratic HPLC using 5 mM H2 SO4 as eluent and an organic acid column (60 ◦ C, flow rate 0.6 mL/min) (CS Chromatographie Service GmbH, Langerwehe, Germany). For the pulse and shaking frequency reduction experiments, a blood glucose measurement device (Accutrend Sensor, Roche diagnostics, Germany) usually used by diabetics was used for the glucose quantification, as the sample volume was too small to perform detailed HPLC analysis. 2.3. Dry cell weight Dry cell weight was determined in doublets by centrifuging 2 mL of fresh sample. The pellets were washed with 0.9% (w/v) NaCl, centrifuged (10 min, 17,000 × g) and dried at 60 ◦ C until constant weight was reached and cooled down in a desiccator before weighing.

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Fig. 1. Maximum oxygen transfer capacities (OTRmax ) in the COSBIOS system in dependency of the shaking frequency [23,24]. Determination of OTRmax with the 0.5 M sulfite system as described by Akguen et al. [23,24].

2.5. Protease activity measurements The protease activity was measured according to the method developed by DelMar et al. [35]. The artificial substrate Suc-AAPFpNA (succinate – l-alanine – l-alanine – l-proline – l-phenylalanine – p-nitroanilide) is cleaved into AAPF and pNA leading to a change in absorption, which can be measured at 405 nm. Cell free supernatant was used for the activity measurement. It was diluted accordingly with 0.1 M Tris/HCl buffer pH 8.6 containing 0.1% Brij 35%. The substrate Suc-AAPF-pNA (70 mg/mL in DMSO) was diluted 20 fold with 0.1 M Tris/HCl buffer pH 8.6 containing 0.1% Brij 35%. To 200 ␮L of the diluted sample, 25 ␮L AAPF solution was added in a microtiter plate. The same amount of buffer was used for blank measurements instead of sample. The kinetic measurement was started immediately after addition of the substrate. From the slope of the absorption changes over time, the activity could be calculated with known extinction coefficient of Suc-AAPF-pNA (ε405 = 8.48 cm2 /␮mol).

2.4. Viscosity measurements

2.6. Determination of maximum oxygen transfer capacity in the COSBIOS

As described by Kubota et al. [31], King et al. [32] and Goto and Kunioka [33], the molecular weight of the formed ␥-PGA can decrease during the fermentation due to the presence of a polyglutamyl hydrolyzing enzyme which breaks down ␥-PGA. Undisputedly, changes in molecular weight influence the viscosity of the fermentation broth. Therefore, viscosity measurements were chosen as a first, general method to follow ␥-PGA formation. Nevertheless, quantification and molecular weight analysis must be performed in future to support the results presented within this study. A MCR 301 rheometer from Anton Paar (Graz, Austria) was used for viscosity measurements using the cone-plate measurement technique (angle 0.467◦ ) and a gap width at the tip of the conus of 0.054 mm. A volume of 0.5 mL of the fresh sample was used for viscosity measurements at 25 ◦ C and a range of shear rates from 100 to 3000 s−1 was applied. As ␥-PGA solutions exhibit pseudoplastic properties, viscosity values for a distinct shear rate of  = 300 s−1 were used to compare experiments with each other. According to Peter et al. [34] the shear rates actually prevailing in a shake flask during cultivation are in the range of  = 300 s−1 [34].

The maximum oxygen transfer capacity of the COSBIOS was investigated. As can be seen in the work of Akguen et al. [23,24], the maximum oxygen transfer capacity (OTRmax ) in the COSBIOS system mainly depends on the applied shaking frequency. Akguen et al. [23,24] determined the OTRmax for shaking frequencies up to 300 rpm with a chemical system, the 0.5 M sulfite system. For details on this method please refer to Akguen et al. [23,24]. In Fig. 1 the OTRmax is plotted over the shaking frequency. An increasing shaking frequency leads to a nearly exponential increase in the OTRmax . It should be noted that increasing shaking frequencies results in decreasing filling volumes [23,24]. Therefore, both changes result in increasing OTRmax values. Since the COSBIOS was operated at 280 rpm and 350 rpm in this study, this diagram was extended with additional measurements for these frequencies. At 280 rpm and 350 rpm, the COSBIOS can supply 34.5 mmol/L/h and 62.5 mmol/L/h of oxygen to a sulfite solution in the flasks, respectively. Because the OTRmax was determined with the sulfite system, the measured values need to be multiplied with a correlation factor for the used medium [23,24]. This correlation factor was determined to be 1.84 for the used modified V3 MOPS medium (data

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Fig. 2. Batch cultivation of Bacillus licheniformis. V3 MOPS medium, 20 g/L glucose, pH0 = 8.0, 200 mM MOPS, n = 350–400 rpm, d0 = 50 mm, 37 ◦ C, VL = 10–55 mL. (A and B) The oxygen transfer rate, (C and F) the glucose concentration, (D and E) the measured viscosities at a shear rate of  = 300 s−1 . For better comparability, the lag-phases of the cultures were adjusted as the cultivations were performed in several experiments.

not shown). Therefore, the maximum oxygen transfer capacities for 280 and 350 rpm in culture medium are 63.5 and 115 mmol/L/h, respectively.

amount of carbon source initially added to the medium, the yield coefficient YO/S can be derived as YO/S = 1.8 mol oxygen/mol substrate. With Eq. (1) the oxygen demand can, thus, be calculated for the specific dilution rates.

2.7. Calculation of oxygen demand According to Akguen et al. [24] the oxygen demand of a culture, OTRdemand , can be calculated according to OTRdemand = D · cS · YO/S

(1)

The yield coefficient YO/S resembles the amount of oxygen needed for the full metabolization of the available carbon source. It was determined through batch experiments using a RAMOS device monitoring the oxygen transfer rate (OTR) (data not shown). Once the carbon source in the experiment is depleted, a characteristic drop in the OTR is visible [21,22]. For this time point, the total oxygen transfer was calculated by integrating the oxygen transfer rate over time. By dividing the calculated total oxygen transfer with the

3. Results and discussion 3.1. Batch cultivation The influence of oxygen limitation on ␥-PGA production and the characteristics of ␥-PGA formation throughout batch fermentations were investigated. The results of the respective batch cultivations in the RAMOS device are depicted in Fig. 2. In Fig. 2A and B, the measured oxygen transfer rates (OTR) are plotted over time, in Fig. 2C and D the measured viscosity at a shear rate of  = 300 s−1 is shown. The glucose concentration in the medium is depicted in Fig. 2C and F. Because the presented data were received from several experiments, lag-phases were adjusted for better

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comparability of the cultivation progresses. In order to impose varying degrees of oxygen limitation onto the cultures, the filling volumes in the flasks were varied. As generally known, increasing the filling volume results in a reduced maximum oxygen transfer capacity. When the oxygen demand of the microorganisms exceeds the maximum oxygen transfer capacity an oxygen limitation occurs, visible in a more or less horizontal plateau in the oxygen transfer rate [22,36]. In Fig. 2A and B all experiments show a very uniform exponential growth phase until the cultures enter oxygen limited conditions. The expected plateau for the oxygen limitation is observed in filling volumes higher than 15 mL. As expected, the maximum OTR decreases with increasing filling volume from ∼ 60 mmol/L/h (10 mL, 400 rpm) to ∼ 20 mmol/L/h (55 mL, 350 rpm). It is furthermore expected, that the length of the plateau lengthens with increasing filling volume. For higher filling volumes, more absolute carbon source is available in the shake flask, while at the same time the amount of available oxygen decreases due to increasing oxygen limitation. Therefore, it takes longer for cultures with high filling volume to fully metabolize the available carbon source. Surprisingly, all filling volumes show the sharp decrease in oxygen transfer rate (OTR) at approximately 18 h, indicating the depletion of the carbon source at the same time. At 18 h, glucose is completely metabolized and not measurable in the cultures supernatants as seen in Fig. 2C and F. As the OTR drops at the same time for all cultures, an oxygen limitation clearly does not affect the glucose uptake rate in B. licheniformis. Furthermore, this first growth phase is followed by a phase with more or less constant breathing activity until the end of the fermentation. With increasing filling volume, a second peak builds up around t = 30 h. This second peak is an indication for the metabolization of an additional carbon source (diauxic growth). As only glucose was added to the medium, this additional carbon source must have been produced during the exponential growth phase. A residual breathing activity, with an OTR ∼ 10 mmol/L/h, remains until 50 h. Also this extended residual breathing activity is a strong hint for the metabolization of considerable amounts of an additional carbon source. Furthermore, the long time of elevated breathing activity (from 30 to 50 h) is pointing to a more complex substrate e.g. a biopolymer as this needs to be hydrolyzed by the organism prior to uptake. Because B. licheniformis is known to produce the biopolymer ␥-PGA the viscosity was measured in samples drawn throughout the cultivation. Fig. 2D and E shows the results. As described above, samples were drawn according to characteristic points in OTR. Thus, maximal viscosities were measured at maximum OTR for all filling volumes, except the experiment with 10 mL at 400 rpm and the 35 mL filling volume. Here, the maximum viscosity was measured at the second OTR peak. In all preparations the viscosity increases to the maximum and decreases to that of an aqueous solution (∼1 mPa s) at the end of fermentation. The formed biopolymer causing the viscosity increase obviously acts as storage compound. Furthermore, with increasing filling volume, and, therefore, increasing oxygen limitation, an increase in viscosity was observed. The maximum viscosity measured for oxygen nonlimited conditions (filling volumes 10 mL, n = 350–400 rpm) was  ∼ 22 mPa s, while the maximum viscosity measured for oxygen limited conditions (filling volumes 15–55 mL) was  ∼ 60 mPa s. Thus, a viscosity increase by the factor three is caused by applying an oxygen limitation to batch cultivations with B. licheniformis in defined mineral medium. To confirm the biopolymer being ␥-PGA, a ␥-PGA-deficient mutant was cultivated under identical conditions. This mutant did not show any viscosity increase at all (data not shown). In consequence, the formed biopolymer by the original strain is most probably ␥-PGA.

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Because experiments with the different filling volumes suffer from carbon catabolite repression, caused by the high initial glucose concentration, the induced oxygen limitation is not necessarily the sole trigger for ␥-PGA formation. Even the oxygen non-limited experiments (400 rpm, 10 mL and 350 rpm, 10 mL) showed an increase in viscosity during the exponential growth phase. In consequence, the results presented in Fig. 2 indicate a possible carbon catabolite repression being one of the key triggers for ␥-PGA formation. Additionally, an oxygen limitation obviously has a potentiating effect on ␥-PGA formation, as a three-fold increase in viscosity was observed with exposure to oxygen limitation. Cromwick et al. [12] observed higher ␥-PGA production with increased aeration for B. licheniformis ATCC9945a cultures grown on medium E. The used strain and medium differ from the ones used within this work regarding carbon sources and overall composition and, therefore, results cannot be directly compared. However, the authors state that even for high aeration conditions, the partial oxygen pressure dropped to values <1.0%. Thus, oxygen limited conditions even prevailed for high aeration conditions, used by these authors, suggesting that the cultures at low aeration conditions were exposed to such severe oxygen limitation that resulted in severely decreased ␥-PGA formation. Thus, also for these experiments oxygen limited conditions as well as citrate mediated carbon catabolite repression prevailed. In B. subtilis, carbon catabolite repression is mediated by a complex interaction of the HPr-kinase and, among others, the global regulator CcpA. Preferred sugars, such as glucose, are transported into the cell by the phosphotransferase system (PTS) and are thereby phosphorylated. The phosphorylated glucose6-phosphate enters the glycolytic pathway and is processed to fructose-1.6-bisphosphate (FBP). High FBP concentrations induce the HPr-kinase, which in turn phosphorylates the HPr or Crh proteins [37–39]. Both, HPr–P as well as Crh–P bind to CcpA. The Hpr–CcpA and Crh–CcpA complexes bind to the cre sites of carbon catabolite regulated genes. Thereby, the carbon catabolite regulation takes place on the transcriptional level [37–39]. On the metabolic level, for the catabolite controlled overflow, the Pta–AckA pathway as well as the production of glutamic acid by GltAB is directly regulated by CcpA. In the presence of glucose, both pathways are induced, resulting in a strong acetate and glutamic acid formation. In consequence, catabolite controlled overflow metabolism takes place. In addition, the succinyl-CoA synthetase is repressed leading to accumulation of succinyl-CoA. Consequently, under catabolite controlled overflow inducing conditions metabolic fluxes are expected to be redirected toward glutamic acid and finally to ␥-PGA, visible in an increase in viscosity. For detailed reviews on CCR, refer to Fujita [39] and Sonenshein [38]. 3.2. Continuous cultivations To investigate the effect of catabolite controlled overflow on ␥PGA formation, continuous cultures in the COSBIOS system were performed. One characteristic of continuous cultivations is that defined carbon-limited conditions can be employed. Therefore, no ␥-PGA formation was expected in steady state carbon-limited conditions. Results of a continuous cultivation performed with the COSBIOS are depicted in Fig. 3. Up to a dilution rate of D = 0.34 h−1 , the carbon sources are limiting as neither glucose nor citrate are quantifiable. When increasing the dilution rate above D = 0.34 h−1 , unutilized citrate accumulates in the culture indicating carbon nonlimited but glucose-limited conditions. The progress of glucose and citrate concentrations illustrates the simultaneous metabolization of both carbon sources for lower and medium dilution rates. With increasing dilution rate, the carbon source glucose that permits

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Fig. 3. Continuous cultivation of Bacillus licheniformis at oxygen non-limited conditions. Modified V3 MOPS medium, 20 g/L glucose, pH0 = 7.5, 250 mM MOPS, n = 350 rpm, d0 = 50 mm, 37 ◦ C, VL = 15.5 mL.

the higher growth rate is preferably metabolized, which is well in accordance with literature for mixed substrate growth in chemostat cultures [40–45]. At a dilution rate higher than D = 0.5 h−1 , the glucose concentration increases and dry cell weight decreases sharply. Thus, it is reasonable to assume that the washout point Dcrit equals about 0.5 h−1 . The maximum oxygen transfer capacity (OTRmax ) as well as the theoretical oxygen demand (OTRdemand ) of the culture was determined, as described above. Since the OTRmax is independent of the dilution rate, it is a constant. The oxygen demand, on the other hand, is dependent on the growth rate, and, therefore, on the dilution rate under steady state conditions. Hence, there is a linear dependency between the dilution rate and the OTRdemand . The boundary condition is found at a dilution rate where OTRmax and OTRdemand correspond [24]. Cultures cultivated at higher dilution rates and, therefore, higher growth rates would be oxygen-limited. As illustrated in Fig. 3B, no oxygen limitation is affecting the culture at any dilution rate since there is no intersection of the dotted OTRmax line and the curve representing the theoretical OTR demand. Thereby, oxygen non-limited conditions are guaranteed when choosing these fermentation parameters in the COSBIOS system. Unexpectedly, the pH is almost constant throughout all dilution rates. A slight decrease to pH ∼ 7.0 with increasing dilution rate is recognizable followed by an increase to pH 7.5 at the point of washout. Surprisingly, no clear correlation between pH and citrate concentration is visible because a decrease in pH with accumulation of citrate would have been expected.

A clear decrease in protease activity is observable with increasing dilution rate due to catabolite repressed protease formation. A decrease with increasing dilution rate is typical for catabolite repressed product formation as the cells face higher carbon source concentration with higher dilution rates. Frankena et al. [16,17] also reported decreasing protease production with increasing dilution rates in a mineral medium with glucose and citrate. No significant increase in viscosity is observable as long as carbon-limited conditions prevail. Once citrate accumulates at a dilution rate of D = 0.36 h−1 , the viscosity increases to  = 10 mPa s. This corresponds to an increase of a factor two compared to carbonlimited conditions at lower dilution rates. When the dilution rate is further increased, a decrease in viscosity is observable. In summary, under oxygen non-limited and carbon-limited conditions no viscosity increase, and, therefore, no significant ␥-PGA formation occurs. If catabolite controlled overflow induces ␥-PGA formation, a clear viscosity increase should have been observed with the onset of accumulation of a carbon source. In agreement to this hypothesis, a twofold increase in viscosity is observable in the presented data once citrate accumulates. Therefore, it is very likely that citrate acts as inducer to catabolite controlled overflow. This hypothesis is further supported by the results of Frankena et al. [16,17]. They stated that citrate acts as carbon catabolite repressor in B. licheniformis. As citrate is an organic acid and no carbohydrate, the regulatory pattern for citrate mediated catabolite controlled overflow should differ from glucose mediated CCR. Consequently, the detailed molecular mechanism needs to be elucidated in future. Since it was observed in batch experiments (Fig. 2) that oxygen limitation has a potentiating effect on ␥-PGA formation, another continuous cultivation experiment was performed with a reduced shaking frequency of 280 rpm to induce oxygen limitation. Viscosity, glucose and citrate concentration are plotted over dilution rate in Fig. 4A. The theoretical OTRdemand , the maximum oxygen transfer capacity (OTRmax ) and the protease activity are shown in Fig. 4B. By reducing the shaking frequency, the volume in the flask was increased from 15.5 to 25 mL (Fig. 1), reducing the maximum oxygen transfer capacity from 115 mmol/L/h (Fig. 3) to 63.5 mmol/L/h (Fig. 4B). As expected and differing from the oxygen non-limited experiment, OTRmax and OTRdemand intersect at D ∼ 0.29 h−1 , indicating oxygen limited conditions for dilution rates higher than D = 0.29 h−1 . Due to the strong viscosity increase with rising dilution rate (Fig. 4A), dry cell weights could not be determined, since the polymer adheres strongly to the cells. Therefore, dry cell weight determination is only possible with high error. Only for the dilution rate of D = 0.11 h−1 , carbon-limited conditions exist (Fig. 4A) as neither glucose nor citrate were measurable. When increasing the dilution rate to D = 0.19 h−1 , citrate is accumulating in the culture liquid. Consequently, citrate mediated catabolite controlled overflow is imposed on the culture [16,17]. Carbon non-limited but glucose-limited conditions prevail in the reactor for dilution rates higher than D = 0.19 h−1 . For the highest dilution rate, D = 0.49 h−1 , 8.75 g/L glucose are detectable, indicating the critical dilution rate for washout being close by. In accordance with the oxygen non-limited experiment, the viscosity increases once citrate accumulates at a dilution rate of D = 0.19 h−1 . The maximum viscosity of  ∼ 34 mPa s was measured at a dilution rate of D = 0.23 h−1 . With a further increase in dilution rate, the viscosity decreases due to stronger dilution and washout of the formed ␥-PGA. Hence, once catabolite controlled overflow is imposed on the culture, ␥-PGA formation is induced. Moreover, the oxygen limitation imposed on the culture clearly has a further potentiating effect on ␥-PGA formation. In fact, 1.9-fold higher viscosities (max = 23.7 mPa s) (Fig. 4A) were measured under clearly oxygen-limited conditions when compared to the maximum

A. Wilming et al. / Biochemical Engineering Journal 73 (2013) 29–37

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Fig. 4. Continuous culture of Bacillus licheniformis under oxygen limited conditions. Modified V3 MOPS medium, 20 g/L glucose, pH0 = 7.5, 250 mM MOPS, n = 280 rpm, d0 = 50 mm, 37 ◦ C, VL = 25 mL.

viscosity for oxygen non-limited conditions (max = 12.5 mPa s) (Fig. 3A) and approximately the same dilution rate (D = 0.34 h−1 ). The pH of the culture decreases with increasing dilution rates. After a pH minimum of ∼6.5 at D = 0.29 h−1 it increases with higher dilution rate to remain constant at ∼6.7 at D > 0.34 h−1 . Cromwick et al. [12] observed an increase in ␥-PGA formation when the pH was decreased from 7.4 to 6.5. This observation is in accordance with the progress of pH and viscosity presented in Fig. 4. However, the conformation of ␥-PGA is pH-dependent according to Borbely et al. [46], Konno et al. [47] and He et al. [14]. At low pH, the unionized polymer shows a helical conformation whereas the ionized polymer acts as in a random-coil state. At high pH, a transition into ␤-sheet conformation takes place. The pH-dependent conformational changes also influence the viscosity. It is reasonable to assume that the viscosity is low for highly organized ␣-helical conformation at low pH. With increasing pH, the transition into random coil results in an increase in viscosity with its maximum at complete random coil conformation of the ␥-PGA in solution. When further increasing the pH the transition into the highly organized ␤-sheet conformation takes place as described by He et al. leading to a decrease in viscosity. It is very likely that conformational changes also occurred in the experiment presented in Fig. 4. Thus, not only the amount of ␥-PGA but also molecular weight, the pH and the conformation influence the measured viscosity. Further experiments need to be performed to clarify which effect affects the viscosity to which extend. Another undesired side effect of the ␥-PGA formation is based on its chelating ability of positively charged ions and molecules [4].

Fig. 5. Influence of glucose pulses on ␥-PGA production. Modified V3 MOPS medium, 20 g/L glucose, pH0 = 7.5, 250 mM MOPS, n = 350 rpm, d0 = 50 mm, 37 ◦ C, VL = 15.5 mL, D = 0.2 h−1 . Carbon-limited and oxygen unlimited conditions were chosen. By injecting a high concentrated (600 g/L) glucose solution, glucose pulses of 10–60 g/L were applied. The concentrations refer to the condition in the fermentation vessels just after the pulse.

As the produced protease subtilisin is positively charged [48], there is a high chance of it being bound to the ␥-PGA and, therefore, not accessible for activity measurements. This could explain the fact that no protease activity was measurable for the dilution rates with highest viscosity, whereas protease activity was measurable in this range of dilution rates for oxygen non-limited conditions (Fig. 3A). 3.3. Pulse experiments To further investigate the influence of catabolite controlled overflow on ␥-PGA production, glucose pulse experiments were performed. One feature of the COSBIOS system is the possibility to run six reactors at the same dilution rate and perform pulse experiments with six different pulse concentrations at once. At a dilution rate of D = 0.2 h−1 and a shaking frequency of 350 rpm, carbon-limited and oxygen non-limited conditions are adjusted (Fig. 3). One glucose pulse was introduced into each of the six reactors. In fact, a high concentrated (600 g/L) glucose stock solution

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presented in this work (Fig. 2). To sum up, a sudden catabolite controlled overflow resulting from high glucose concentrations obviously has induced ␥-PGA production, visible in the viscosity increase after the glucose pulse. As reported, an increase in viscosity was observed with increasing oxygen limitation in batch experiments (Fig. 2). Therefore, an experiment was performed in which an oxygen limitation was applied to a steady-state culture in the COSBIOS system. By reducing the shaking frequency from 350 rpm to 200 rpm, the filling volume is increased to 68 mL and the oxygen transfer is reduced due to a decreased surface-to-volume ratio. Thereby, an oxygen limitation is induced onto the culture. At t = 0 h, the shaking frequency was reduced to 200 rpm. After establishment of new steady-state conditions, samples were drawn at t = 24 h and analyzed for glucose content and viscosity as depicted in Fig. 6. Three parallel experiments at dilution rates of D = 0.218 h−1 , D = 0.233 h−1 and D = 0.247 h−1 were performed. In each experiment, the glucose concentration indicated a carbon-limited cultivation at the point of shaking frequency reduction. The viscosity was identical in all three experiments at 2.9 mPa s at t = 0 h. Exposure to oxygen limitation for 24 h leads to an accumulation of glucose in the reactors, a clear indication for an oxygen limitation in continuous cultures. The viscosity increased three to fivefold compared to the oxygen unlimited conditions before shaking frequency reduction. As demonstrated above, the OTRdemand is increasing with increasing dilution rate. Thus, the higher the dilution rate, the stronger the oxygen limitation as the OTRmax is constantly low at ∼23.18 mmol/L/h at a filling volume of VL = 68 mL in this experiment after reducing the frequency to 200 rpm. Therefore, the rise in viscosity increase at t = 24 h with increasing dilution rate is in line with the explanation derived above. Furthermore, this is another proof of the potentiating effect an oxygen limitation has on the ␥-PGA formation. 4. Conclusions

Fig. 6. Influence of oxygen limitation and carbon catabolite repression on ␥polyglutamic acid formation. Modified V3 MOPS medium, 20 g/L glucose, pH0 = 7.5, 250 mM MOPS, n0 = 350 rpm, d0 = 50 mm, 37 ◦ C, VL0 = 15.5 mL. By reducing the shaking frequency from 350 rpm to 200 rpm at t = 0 h, the filling volume in the bioreactors is increased and, thereby, an oxygen limitation is induced onto the culture.

was injected using a sterile syringe. Thus, pulses with varying glucose concentrations of 10–60 g/L (Fig. 5) in the reactor after the pulse were introduced. Three samples were drawn from each reactor at t = 0, t = 3.5 and t = 7 h. Glucose concentration and viscosity are plotted over the pulse concentration in Fig. 5A and B for the three samples drawn. After 3.5 h, glucose was measurable in all reactors, but the one exposed to the 10 g/L glucose pulse. After 7 h, glucose-limited conditions, as before the pulses, were restored in all reactors. Upon induction of catabolite controlled overflow by the glucose pulse, a viscosity increase was detectable 3.5 h after the pulse in all reactors. As expected, the viscosity increase rises with increasing glucose pulse concentration up to 40 g/L glucose. For the two pulses with 50 and 60 g/L glucose, the viscosity increase is clearly within the range of the 10 g/L pulse, and, therefore, no further increase took place as would have been expected. The reason for this observation is unclear. In all vessels, the viscosity decreased significantly from t = 3.5 h to t = 7 h. As B. licheniformis is able to use ␥-PGA as substrate, this was expected from the experiments

For the first time, the continuous parallel shaken bioreactor system (COSBIOS) was used to perform small-scale continuous cultivations with B. licheniformis in a mineral medium (modified V3 MOPS medium). In continuous culture, B. licheniformis is able to utilize two carbon sources simultaneously for low to intermediate dilution rates. With increasing dilution rate the carbon source enabling the highest growth is preferably metabolized. Therefore, unutilized citrate accumulates in the culture liquid as glucose is the preferred carbon and energy source for the B. licheniformis strain used in this work. The results presented in this article illustrate that the viscosity, and, therefore, the ␥-PGA production, correlates strongly with catabolite controlled overflow. A sudden initiation of catabolite controlled overflow results in an increasing viscosity. In addition, it was demonstrated that ␥-PGA formation also correlates with the accumulation of citrate. Thereby, the hypothesis of citrate inducing catabolite controlled overflow in B. licheniformis and subsequently leading to ␥-PGA formation is strongly supported. Differences in the regulatory patterns of citrate and glucose mediated catabolite controlled overflow need to be elucidated in future. Our observations indicate a possible variation in the regulatory system of B. licheniformis compared to B. subtilis. As 2-oxoglutarate is a precursor for ␥-PGA, the citrate synthase CitZ cannot be strongly repressed under catabolite controlled overflow. Otherwise, citrate would not enter the TCA, be processed to 2-oxoglutarate and last but not least, ␥-PGA could not be produced to such an extent as it was observed repeatedly for ␥-PGA producing cultures in our experiments. Accumulation of 2-oxoglutarate is a strong indicator for a repression of the 2-oxoglutarate dehydrogenase complex under CCR. Therefore, a redirection of metabolic fluxes toward glutamic acid and next toward ␥-polyglutamic acid occurs.

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Oxygen limitation was shown to have a further potentiating effect on ␥-PGA production. Therefore, similar regulation patterns for carbon catabolite repression and oxygen limitation are most likely. The oxidation of 2-oxoglutarate to succinic acid in the TCA is a NADH-releasing reaction. Under oxygen limitation, the NADH cannot be regenerated in the respiratory chain, thus, 2oxoglutarate accumulates. In other words, the tricarboxylic acid cycle becomes a bottleneck under carbon catabolite repression and simultaneous oxygen limitation. Consequently, more experiments need to be performed to elucidate these regulation patterns, especially regarding the effects both have on ␥-polyglutamic acid formation. For this purpose, metabolomic, proteomic and transcriptomic analyses would be useful. References [1] A.L. Sonenshein, J.A. Hoch, R. Losick, Bacillus subtilis and its Closest Relatives, first ed., ASM Press, Washington, DC, 2002. [2] K. Maurer, Detergent proteases, Curr. Opin. Biotechnol. 15 (2004) 330–334. [3] R. Gupta, Q.K. Beg, P. Lorenz, Bacterial alkaline proteases: molecular approaches and industrial applications, Appl. Microbiol. Biotechnol. 59 (2002) 15–32. [4] I. Shih, Y. Van, The production of poly-(␥-glutatmic acid) from microorganisms and its various applications, Bioresour. Technol. 79 (2001) 207–225. [5] S.H. Yoon, J.H. Do, S.Y. Lee, H.N. Chang, Production of poly-␥-glutamic acid by fed-batch cultures of Bacillus licheniformis, Biotechnol. Lett. 22 (2000) 585–588. [6] Y.H. Ko, R.A. Gross, Effects of glucose and glycerol on ␥-poly(glutamic acid) formation by Bacillus licheniformis ATCC 9945, Biotechnol. Bioeng. 57 (1998) 430–437. [7] A. Richard, A. Margaritis, Poly(glutamic acid) for biomedical applications, Crit. Rev. Biochem. 21 (2001) 219–232. [8] F.A. Troy, Chemistry and biosynthesis of the poly(␥-d-glutamyl) capsule in Bacillus licheniformis: 1. Properties of the membrane-mediated biosynthetic reaction, J. Biol. Chem. 248 (1973) 305–315. [9] F.A. Troy, Chemistry and biosynthesis of the poly(␥-d-glutamyl) capsule in Bacillus licheniformis: 2. Characterization and structural properties of the enzymatically synthesized polymer, J. Biol. Chem. 248 (1973) 316–324. [10] A. Cromwick, R.A. Gross, Effects of manganese (II) on Bacillus licheniformis ATCC 9945A physiology and poly-(␥-glutamic acid) formation, Int. J. Biol. Macromol. 17 (1995) 259–267. [11] G.A. Birrer, A. Cromwick, R.A. Gross, ␥-Poly(glutamic acid) formation by Bacillus licheniformis 9945A: physiological and biochemical studies, Int. J. Biol. Macromol. 16 (1994) 265–276. [12] A. Cromwick, G.A. Birrer, R.A. Gross, Effects of pH and aeration on ␥poly(glutamic acid) formation by Bacillus licheniformis in controlled batch fermentor cultures, Biotechnol. Bioeng. 50 (1996) 222–227. [13] C.B. Thorne, C.G. Gomez, H.E. Noyes, R.D. Housewright, Production of glutamyl polypeptide by Bacillus subtilis, J. Bacteriol. 68 (1954) 307–315. [14] L.M. He, M.P. Neu, L.A. Vanderberg, Bacillus licheniformis ␥-glutamyl exopolymer: physicochemical and U(VI) interaction, Environ. Sci. Technol. 34 (2000) 1694–1701. [15] C.G. Leonard, R.D. Housewright, C.B. Thorne, Effects of some metallic ions on glutamyl polypeptide synthesis by Bacillus subtilis, J. Bacteriol. 76 (1958) 499–503. [16] J. Frankena, H.W. van Versefeld, A.H. Stouthamer, A continuous culture study of the bioenergetic aspects of growth and production of exocellular protease in Bacillus licheniformis, Appl. Microbiol. Biotechnol. 22 (1985) 169–176. [17] J. Frankena, G.M. Koningstein, H.W. van Versefeld, A.H. Stouthamer, Effect of different limitations in chemostat cultures on growth and production of exocellular protease by Bacillus licheniformis, Appl. Microbiol. Biotechnol. 24 (1986) 106–112. [18] M. Dauner, T. Storni, U. Sauer, Bacillus subtilis metabolism and energetics in carbon-limited and excess-carbon chemostat culture, J. Bacteriol. 183 (2001) 7308–7317. [19] M. Dauner, M. Sonderegger, M. Hochuli, T. Szyperski, K. Wüthrich, H. Hohmann, U. Sauer, J.E. Bailey, Intracellular carbon fluxes in riboflavin-producing Bacillus subtilis during growth on two-carbon mixtures, Appl. Environ. Microbiol. 68 (2002) 1760–1771.

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