Appl Microbiol Biotechnol(1991) 35:10-13 017575989100080T

Applied Microbiology Biotechnology © Springer-Verlag 1991

Studies on the production of lipase from recombinant Staphylococcus carnosus M. P. F. Falk, E. A. Sanders, and W.-D. Deckwer GBF - Gesellschaftfor Biotechno|ogischeForschungmbH, Mascheroder Weg 1, D-3300 Braunschweig,Federal Republic of Germany Received 7 September 1990/Accepted 10 December 1990

Summary. Production of lipase from recombinant S t a phylococcus carnosus pLipPS1 was studied in standard stirred tank bioreactors. Only low lipase activity was obtained under conventional operating conditions, i.e., moderate to high stirring speeds and aeration rates for keeping the dissolved oxygen concentration at high levels. Additional targetted experiments indicated that the reason for the observed low lipase activity is lipase inactivation due to surface forces and shear stress at the gas/liquid interface. Therefore, a cultivation strategy is proposed that minimizes gas/liquid interfacial area and maximizes the driving concentration for O2 mass transfer by controlling the dissolved oxygen to low values by gentle stirring and low aeration rates. Thus, high lipase activities can be obtained even in larger scale standard stirred tank bioreactors.

Introduction Lipases (glycerol ester hydrolases EC 3.1.1.3) have been traditionally obtained from animal pancreas as a digestive aid for human consumption. Initial interest in microbial lipases was generated as a result of shortage of pancreas and difficulties in collecting available materials. The yeasts Candida lipolytica and C. cylindracea have been identified as the most important microorganisms to produce lipases. However, there is also an interest in using bacterial strains as they generally offer higher activities compared to yeasts (Frost and Moss 1987). It has been known for many years that certain S t a phylococcus spp. produce extracellular lipases (Arvidson and Holme 1971; Sztajer et al. 1988). Some species are pathogenic whereas others have been extensively used as starter cultures for the food industry. In this study a strain of S. carnosus, generally regarded as safe, was used as host for the plasmid pLipPS1 with a lipase

Offprint requests to: W.-D. Deckwer

gene from S. hyicus (G/Stz et al. 1985), which encodes for an 86-kDa lipase. Most of the staphylococcal lipases show a wide substrate specifity, i.e., hydrolysis of both water-soluble and insoluble triglycerides as well as polyoxyethylene-sorbitan fatty acids (Tweens) can be used as a test for lipase activity (Lechner et al. 1988). Lipase production from S. carnosus rec was investigated by Lechner et al. (1988). The authors compared fermentations in a special bioreactor consisting of two compartments separated by a dialysis membrane with conventional batch technology. A remarkable increase in product concentration from 3.7 mg enzyme/l in the case of a 2-1 standard laboratory fermentor up to 20 mg/l in the inner compartment of the dialysis system was observed. Furthermore, a huge increase in product concentration in the inner part of the fermentor could be obtained in semi-continuous cultivation with concentrated media in the outer compartment. The dry cell mass reached 60 g/1 and the lipase concentration was about 225 mg/1 after 46 h cultivation. S. carnosus grows only on complex media, which leads

to excessive foaming by aeration. Therefore, a centrifugal bioreactor has been used to cultivate S. carnosus for lipase production (Mersmann, personal communication). This paper reports on studies to produce lipase from the same strain of S. carnosus (Grtz et al. 1985) in conventional equipment, i.e., standard type stirred tank bioreactors. The aim was to provide sufficient amounts of lipase for its structural analysis and applicability studies. It turned out that lipase production in traditional bioreactors requires special cultivation strategies.

Materials and methods Bacterial strain, plasmid and media. S. carnosus TM300::pLipPS1

was kindly provided by F. G/Stz, University of TObingen, FRG. The pLipPS1 plasmid (4.16 kbp) carries the lipase gene of S. hyicus and gives lipase overproduction in S. carnosus (Liebl and Grtz 1986). Precultivationas well as all fermentationstudies were

11 carried out on 37 g/1 brain-heart-infusion medium (BHI) obtained from Difco (Detroit Mich., USA), with 10 mg chloramphenicol (Serva, Heidelberg, FRG) per litre. A selective agar to discover the best lipase-producing colonies was prepared with 1% (v/v) Tween 20 (Merck, Darmstadt, FRG).

Analytical methods, Cell dry mass (CDM) was determined by filtration (0.1 ~tm, Sartorius, Grttingen, FRG) of 10 ml culture, washing with water and drying at 105°C until constant weight. Lipase activity, concentration of sugars, ammonia and acetate were measured in the supernatants of centrifuged samples (5 min, 8200 g), which were stored at - 2 0 ° C until use. Lipase activity was estimated with p-nitrophenyl palmitate, pNPP (Sigma), as substrate (Winkler and Stuckmann 1979). The test buffer was prepared by emulsifying 1 ml solution A (90 mg pNPP dissolved in 30 ml isopropanol) in 9 ml solution B (2 g Triton-X-100, Merck, 0.5 g gum arabic, Serva, in 450 ml 50 mM TRIS/HCI, pH 8.0). The enzymatic activity was determined by mixing 900 p.1 test buffer with 100 ktl of the sample. Liberation of p-nitrophenol at 37°C was detected at 410 nm over a period of 90 s with an Ultrospec K spectrophotometer (Pharmacia-LKB, Uppsala, Sweden). One unit is defined as the cleavage of 1 ~tmol pNPP into p-nitrophenoi and palmitate per minute at 37 ° C. The molar adsorption coefficient of pNPP was taken as 14.9.10 -3 cm 2 ixmol-l: 1 mg lipase corresponds to 150 units (U) of enzyme activity (Erdmann, personal communication). To check the purity of the enzyme and its behaviour in some inactivation experiments sodium dodecyl sulphate-polyacrylamide gel electrophoresis (SDS-PAGE) (Laemmli 1970) with a gradient of 8-18% was carried out with a PhastSystem or Multiphor II (both Pharmacia-LKB) followed by silver staining (Heukeshoven and Dernick 1988). The concentration of acetate, ammonia, glucose and fructose were assayed by enzymatic analysis (t~est kit 148.261, 542.946, 716.251, 139.106, Boehringer, Mannheim, FRG). Protease activity in the supernatant was measured according to Schleuning and Fritz (1976). Precultivation. When a standard 2-1 laboratory fermentor was inoculated from an overnight shake-flask culture that was started from normal BHI agar petri dishes, an unacceptable scatter in the experimental results was obtained. This problem was overcome by using fresh nutrient agar slants containing 1% (v/v) Tween 20 and picking only clones that were surrounded by visible deposits of precipitated, insoluble fatty acid.

Results and discussion

Growth and product formation were studied in several batch fermentations in a 2-1 stirred bioreactor. Sugars (glucose, fructose) and oligopeptides both present in the BHI medium served as the main substrates. A typical time course of a cultivation carried out at 37 ° C, pH 7.4, a stirrer speed of 300-500 rpm and an aeration rate of 0.2 vvm (air) is given in Fig. 1. To avoid oxygen lim-. itation the dissolved oxygen (DO) was maintained at around 50% saturation by appropriately adjusting the stirrer speed stepwise. After 12 h a maximal cell density of about 2.6g CDM/I was reached, the maximum growth rate being 0.9 h -1. Lipase activity appeared after 6 h cultivation and passed through a maximum of about 0.25 U/ml (1.7 mg/l). At the end of the cultivation the lipase activity was less than 0.1 U/ml (0.67 mg/ 1) whereas in shake flasks a nearly stable maximum of 2.1-2.7 U/ml (14-18 mg/1) was obtained reproducibly. The decline in lipase activity during cultivation in the bioreactor as a result of protease formation could be excluded as protease tests were negative. To clarify the low level of lipase production and its decline during cultivation in the stirred bioreactor, additional experiments were carried out in shake flasks with and without baffles at 30 and 37 ° C. The different cultivation conditions resulted in only moderate deviations in the biomass concentration, which on average was about 2.4 g/1 after 36 h cultivation. However, as shown in Fig. 2, pronounced differences in lipase concentration (always measured as activity) were observed. o

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Culture conditions. A 2-1 glass fermentor (Setric Grnie Industrial, Toulouse, France) with 1.2 1 BHI medium was inoculated with 30 ml of the overnight culture, resulting in a start concentration of 0.03 g CDM/1. The cells were grown at various temperatures (2537 ° C) and pH 7.4 controlled by the addition of 1 M NaOH or 0.33 M H3PO4. Dissolved oxygen concentration was controlled by aeration or agitation rate. Further cultivations under optimized conditions were carried out in fermentors of 100 1 (B50, Giovanola Frrres S.A., Montey, Switzerland), and 1501 (P150, Bioengineering, Wald, Switzerland) with a culture volume of 701 and 1201, respectively. Inoculation was carried out as described in the case of the 2 1 fermentor. The volume of the overnight culture was 1% of the final culture volume. Temperature and pH was controlled at 30°C and 7.4, respectively. Inactivation experiments. Separate studies on the stability of the lipase were carried out at 30 ° C with a solution of purified lipase in a small bubble column reactor (chromatographic column X K 16/40, Pharmacia-LKB). A sintered plate was used as sparger for humidified air and helium while the column was filled with 20 ml of a lipase solution (3 U/ml, 50 mM TRIS/HC1, pH 8.0). Samples of 0.5 ml were taken from the top of the column.

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It was also found in the shake flasks that the lipase concentration passed a maximum under all conditions provided the observation time was long enough. Even more remarkable are however the differences in lipase activity obtained from cultivations with baffled and unbaffled shaking flasks. Obviously, the unbaffled system yielded considerably higher lipase activities. In addition, there was a temperature effect on lipase productivity. A cultivation temperature of 30 ° C gave better lipase productivity than 37 ° C. From the results shown in Figs. 1 and 2 one has to conclude that lipase is subjected to some inactivation mechanism. The deactivation of the lipase produced presumably depends on turbulence intensity and/or 02 mass transfer rate as the unbaffled system gives better results. Mozhaev and Martinek (1982) summarized the major phenomena that may cause enzyme inactivation, such as aggregation and alterations in primary structure, for example, chemical modification of functional groups, cleavage of S-S bonds, dissociation of oligomeric proteins into subunits and conformational changes in the macromolecule. In the particular case of lipase, inactivation due do photooxidation (Sugiura and Oikawa 1980) and shear (Lee and Choo 1989) can apparently play an important role. Since lipase contains SH groups inactivation by oxidation may also be considered.

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To discriminate among the rival deactivation mechanisms a solution of purified lipase (3 U/ml) in TRIS/ HCI buffer (pH 8.0) was aerated with either air or helium in a small bubble column at 30 ° C. In both cases, the same drastic decrease in lipase activity (kinetic test with pNPP) was detected. As shown in Fig. 3 the inactivation seemingly depended on the gassing rate. If no gas passed through the lipase solution, which is moderately stirred in the column, only a small decrease in enzyme activity was observed (1.7%/h). In the deactivation experiments the lipase concentration was additionally determined by SDS-PAGE. Typical results are shown in Fig. 4. No weakening of the lipase band can be detected and other bands due to degradation into subunits cannot be observed. Hence, oxidative effects and cleavage of the lipase can be excluded as prevailing inactivation mechanisms. Therefore, it is suspected that the observed lipase deactivation is due to changes in protein folding caused by surface forces and shear stress at the gas/liquid interfacial area. A similar interpretation has been given for lipase inactivation from C. cylindracea by Lee and Choo (1989). From these findings it is understood that production of lipase from S. carnosus rec should be carried out under conditions where the gas/liquid interfacial area (a) can be minimized or, at least, kept at low values. One way of overcoming the problem could be 02 transfer

Fig. 4. Sodium dodecyl sulphate-polyacrylamide gel electrophoresis with silver staining, indicating no change in lipase concentration during gassing with helium

13

into the microbial culture by making use of bubble-free aeration with hydrophobic membranes. This method is extensively used in mammalian cell cultivation (Lehmann et al. 1988). However, an estimation of the achievable 02 transfer rates indicated that they are too small to be applied in the present cultivation of S. carnosus. In general, the 02 transfer rate (OTR) is given by OTR = kLa(c~ -- cz.)

(1)

where, k~a is the volumetric gas/liquid mass transfer coefficient, cf~ is 02 saturation concentration in liquid phase and c~ is dissolved oxygen concentration. As the present task requires minimization of a (or kz.a), the driving concentration difference has to be enlarged to maintain the appropriate 02 supply to the culture. This can be done firstly by reducing c~, i.e., by keeping the DO as low as possible. Secondly, c~ can be enlarged by decreasing the cultivation temperature and by using 02 or O2-enriched air. When maintaining the DO at 5% saturation, no limitation of growth and metabolite formation was detected. Therefore, all further cultivations were done at a DO of 5% which was controlled by variation of impeller speed (50-300 rpm) and gas flow rate (0.05-0.2 vvm). The results obtained at different cultivation temperatures are given in Table 1. Lipase activities similar to that obtained in experiments with unbaffled shaking flasks were now also reached in the stirred reactor (2 1). Although at 25°C the lipase activity was somewhat higher, a clear temperature dependency of lipase activity could not be identified. At low temperatures the growth rate was low and hence lipase productivity was Table 1. Influence of temperature on lipase productivity

Temperature (°C)

Cell dry mass (g/l)

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Cultivation time

Lipase productivity (U/ml/h)

(h) 25 28 30

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also low. The best result was obtained at 30 ° C with a lipase productivity of 0.21 U m1-1 h -1 (1.4 mg h -1 1-1). When using pure 02 instead of air, the stirrer speed and aeration rate could be further reduced. However, a significant increase in lipase productivity was not found. Additional cultivations on a larger scale were carried out under control of the DO at 5% saturation and aeration by air at pH 7.4 and 30 ° C. The results from standard stirred tank reactors of 100 and 150 1 volume (working volumes of 70 and 120 1) are shown in Fig. 5. The course of biomass concentration and lipase activity was very similar to that observed in the 2-1 stirred bioreactor. Surprisingly, the lipase activity was even somewhat higher in the larger equipment (cf. Table 1). This indicates that there is no serious scale-up problem for the production of lipase from S. c a r n o s u s rec provided that the proposed cultivation strategy is applied. This strategy minimizes the gas/liquid interfacial area and maximizes the driving concentration difference for 02 mass transfer by controlling the DO to a low value (5% air saturation) using only gentle stirring and low aeration rates. It should also be mentioned that the proposed cultivation method does not lead to any foam problem. References Arvidson S, Holme T (1971) Influence of pH on the formation of extracellular proteins by Staphylococcus aureus. Acta Path Microbiol Scand Sect B 79:406-413 Frost GM, Moss DA (1987) Production of enzymes by fermentation. In: Rehm HJ, Reed G (eds) Biotechnology, vol 7a. Verlag Chemie, Weinheim, pp 65-211 Grtz F, Popp F, Korn E, Schleifer KH (1985) Complete nucleotide sequence of the lipase gene from Staphylococcus hyicus cloned in Staphylococcus carnosus. Nucleic Acids Res 13 : 5895-5906 Heukeshoven J, Demick R (1988) Improved silver staining procedure for fast staining in PhastSystem development unit. Electrophoresis 9: 28-32 Laemmli UK (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227:680-685 Lechner M, M~irkl H, Grtz F (1988) Lipase production of Staphylococcus carnosus in a dialysis fermentor. Appl Microbiol Biotechnol 28: 345-349 Lee YK, Choo CL (1989) The kinetics and mechanism of shear inactivation of lipase from Candida cylindracea. Biotechnol Bioeng 33 : 183-190 Lehmann J, Vorlop J, BOntemeyer H (1988) Bubble-free reactors and the development for continuous culture with cell recycle. In: Spier RE, Griffiths JB (eds) Animal cell biotechnology, vol 3, Academic Press, London, pp 221-237 Liebl W, Grtz F (1986) Studies on lipase directed export of Escherichia coli fl-lactamase in Staphylococcus carnosus. Mol Gen Genet 204:166-173 Mozhaev VV, Martinek K (1982) Inactivation and reactivation of proteins (enzymes). Enzyme Microb Technol 4:299-309 Schleuning WD, Fritz H (1976) Sperm acrosin. Methods Enzymol 45: 330-342 Sugiura M, Oikawa (1980) Chemical modification of tryptophan and histidine residues in lipoprotein lipase from Pseudotnonas fluorescens. Chem Pharm Bull 28:2803-2806 Sztajer H, Maliszewska J, Wieczorek J (1988) Production of exogenous lipases by bacteria, fungi, and actinomycetes. Enzyme Microb Technol 10:492-497 Winkler UK, Stuckmann M (1979) Hyaloronate and some other polysaccharides greatly enhance the formation of exolipases by Servatia marcescens. J Bacteriol 138:663-670