Immobilization of Bacteria in Macro- and Microparticles

David B. Knaebel, Keith E. Stormo, and Ronald L. Crawford

1. Introduction Microencapsulation of bacteria is a technique that offers some advantages

to the scientist interested m bioremediation applications. Several studies have demonstrated the utility of using immobilized cells in btoreactor (see Chapter 6) or bioremediation settings (1-4). In this chapter, we describe three protocols for the immobilization of bacteria in macroparticles (200 pm-3 mm diameter) and microparticles (2-200 pm diameter) composed of different polymers. We provide information on the choice of polymers and particle sizes and the envi- ronmental applications of rmmobilized cells. Other sources provide informa- tion about immobilization and encapsulation that is beyond the scope of this chapter, such as the encapsulation of enzymes or fungi (see Chapter 7) or eukaryotic cells. Kolot (5) provides protocols for immobilizing yeast, bacteria, and enzymes for industrial apphcations. Akin (6) (and references therem) pro- vide a review of how rmmobrlization may enhance several catalytic processes The ACS symposium proceedings, Immobilized Microbial Cells (7), provides summary accounts of tmmobilizing bacteria in several natural and synthetic polymers. Saher (8) also provides some recent developments in the immobi- lization of cells.

Encapsulation, or more properly, particle immobilization, is a process by which cells are entrapped in semipermeable polymer spheres, with the cells uni- formly distributed throughout. The process is accomplished by mixing cells with a prepolymeric solutron, applying a force that separates the polymer/cell mixture mto generally spherical particles, and allowing the prepolymeric mate- rial to solidify. Polymers that are commonly used include algmate, carrageenan, agarose, polyurethane, polyacrylamide, and methacrylate. Depending on the

From Methods m B~ofechnology, l/o/ 2 Eboremed/abon Protocols Edited by D Sheehan Humana Press Inc , Totowa, NJ

67

68 Knaebel, Stormo, and Crawford

polymer and formation technique used, the partrcles range m size from very small (approx 2-10 pm drameter) to very large (approx 3 mm). Both polymer type and particle stze influence cellular actrvtty, retention, nutrient drffusion, and parttcle stability

The mam benefit of particle tmmobrlizatlon for envu-onmental apphcations IS that mn-nobihzed cells often have enhanced or more stable activity than free cells. This is because particle-immobilized bacteria:

1. Are protected from bacterrophage (9), protozoan (or other) predators, or toxins (IO),

2 Can be supplemented with additional, cotmmoblhzed nutrients (II), 3 Can survive long periods of time and retain physiological actrvuy (1,12), and 4 Can be tmmobrlized at very hrgh cell densities (13)

Furthermore, bacteria may preferentially grow m certain zones (e.g , mrcroaerophihc) wtthm the particle, which will lead to prolonged activity com- pared to that of free cells (14,15). Because of these benefits, particle immobr- lizatron can sometrmes lead to better stability and activity characterlsttcs than those offered by surface rmmobilizatron (16), where the cells are immobtlrzed only on the surface of a particle (I 7).

Besides being stabilized by tmmoblhzation, cells that are particle-rmmobl- hzed usually retam physlologtcal activities similar to those of free cells. Most commonly used polymers are porous, and permit adequate diffusion of nutri- ents, dissolved gases, and metabohc byproducts, However, higher polymer con- centrations or cell loadings, or larger particle sizes can create drffusronal hmitattons (18) (e.g., decrease oxygen availability to the immobilized cells) (19-21), which can lead to a reduction m some aspects of cellular activity. This problem can be overcome by use of smaller particles (- 200 pm diameter) or a more porous polymer, where transport and diffusronal limrtattons have less impact. For example, some types of mtcrocapsules (tc-carrageenans) have a limit of oxygen transport of approx 100 pm (19), a particle size >200 pm dram- eter should therefore be unaffected by O2 limitation. A slightly greater limit of activity and growth (-200 mm) has been observed m other matrices, where cells grew primarily at the outer edge of larger capsules (I4,22,23). This dlffusional barrier, however, may also protect the cells from high concentrations of a toxic chemical that 1s being degraded (24).

Dtfferent polymers have different environmental stabilities. Some carbohy- drate-based polymers are unstable under some environmental conditions or m the presence of some buffers, and may be degraded by some mrcroorganisms. These factors may allow for cell escape into the medium or environmental matrix, which may be either undesirable or advantageous. For example, Ca- algmate requires calcmm ions for maintenance of the solid polymer, the poly-

Bacteria in Macro- and Micropartlcles 69

mer is drsrupted if phosphate or citrate ions are present. Agar, agarose, chitosan, and tc-carrageenans are all biodegradable, albeit slowly. Some of these poly- mers can be strengthened by secondary treatments. Hertzberg et al. (25) devel- oped a method to crosslink algmate beads wrth a polyvmyl alcohol/strlbazolmm matrix that should be more stable than alginate alone. Birnbaum et al. (26) used a polyethylenimme/glutaraldehyde treatment to stabrlize alginate beads to pre- vent dissolution by phosphate buffers. Iijima et al (27) coated alginate beads with polyurethane to prevent cell leakage and improve the particle’s strength. Knaebel and Crawford (unpublished results) developed a similar procedure for coating agarose microcapsules with polyurethane to prevent cell leakage or phage susceptibility.

We have used both macroencapsulation (capsules >0.5 mm diameter) and mrcroencapsulatron (capsules co.5 mm diameter) to rmmobrlize bacteria to study or enhance biodegradative activities in broreactor and envtronmental set- tings. We have used several polymers (e.g., agarose, alginate, and polyurethane 11,12,28; see also 17,27,29,30), but several other matrix materials have been applied m other studies; for example, carrageenans (15,18,19,31), polyacry- lamides (18), methacrylates and polyacrylates (32-35), chitosan (18), agar (29), and polyamides (36). The main criteria for determining whrch particular polymer to use were the condmons required for cellular activity and ease of production.

We designed our particle immobthzation protocols to be relatively easy and consistent (reproducible), and to provide cells that retam high acttvtty following tmmobrhzation. Since we work with defined bacterial systems, we also required that the materials be amenable to stenhzation. This chapter will describe some general methods that we have used to encapsulate cells, one method is based on an emulsion technique to form small (0.0542 mm diameter) agarose mtcropar- titles (37), another method is based on the crosslinkmg of alginate by Ca2+ to form macroparticles (2-3 mm diameter), and another uses a low pressure fog- gmg nozzle to produce much smaller alginate microcapsules (2-100 pm diame- ter, II). The techniques produce cells entrapped in either calcmm algmate or agarose particles that are formed when the polymer/cell mixture IS dispersed mto an appropriate receiving bath. The low pressure nozzle microencapsulatton entraps the cells m very small mrcrobeads as they exit a fogging nozzle orrftce. These microbeads are also solidified when they contact an appropriate recetving bath. The agarose mrcroparttcle method and the alginate macropartrcle method are simpler, can produce great quantities of entrapped cells, and require only minimal equipment. The low-pressure nozzle method can also produce great quantmes of entrapped cells and produces very small mtcrobeads, but requires specialized, although generally avatlable, equipment. The methods are adaptable to multiple polymers and cell types.

70 Knaebel, Stormo, and Crawford

2. Materials

2.1. Agarose/Oil Emulsion Technique for Microparticle (50-200 ,um Diameter) Immobilization

Any equipment or materials that will come m contact with the cells and poly- mer material should be sterile Aseptic techmque should be used throughout these procedures.

1 Approprtately grown and prepared cells 2 Autoclaved agarose (Low EEO; Gibco-BRL, Gatthersburg, MD) solution (that is

twice the final percentage desired) m the same buffer used for cell resuspenston 3 Canola oil (food grade quality) (600-800 mL) m a large capacity glass bottle (1 L),

sterthzed by heating (autoclavmg) with lid tightly sealed. 4. Sterile centrifuge bottles (250 or 500 mL, Nalgene) 5 Ice bath. 6 Two-liter autoclaved and oven-dried glass beaker 7 Autoclaved impeller three-blade propeller type or high-efficiency impeller (e.g ,

Aa- 10, Cole Parmer, Ntles, IL) and stainless steel shaft 8. Mixer Lightmn Labmaster (Cole-Parmer) or equivalent 9 Lammar flow hood

10 Four (4) liters of sterile phosphate-buffered salme (PBS) at appropriate pH con- taining 0.1% Tween 80 or other surfactant that is compatible with the cells used.

11 Two (2) liters of sterile at appropriate pH

2.2. Calcium Alginate “Drop” Technique for Macroparticle (-2 mm Diameter) Immobilization

1. Appropriately grown and prepared cells (e g , Pseudomonas putzda mt-2, Flavobactenum sp., Eschenchra ~011, and so on).

2 Sterile sodium algmate at appropriate concentration (e g., 6%) made up m non- phosphate buffer [e g., 50 mM HEPES (N-(2-hydroxyethyl)piperazme-N-ethane- sulfonic acid); Sigma, St Louts, MO] at pH 7 3 (see Note 3).

3 Sterile 50 mM CaCl,, m HEPES or water, chilled or at room temperature, m a large- volume beaker contammg a stenle stir bar

4 Stertle storage bottles, 5. Variable speed Peristaltic pump and autoclaved peristaltic pump tubing A stenle

capillary may be attached to the end of the tubing to make slightly smaller droplets, which will result m smaller particles

2.3. Low-Pressure Fogging Nozzle Technique for Microparlicle (5-100 ,um Diameter) Immobilization in Alginate (1,ll)

1 Appropriately grown and prepared cells (see Note 3 for appropriate buffers) 2 Sterile algmate (4-6%) m HEPES buffer 3. Four (4) liters of 50 mM CaCl, (autoclaved, in spray chamber, #4)

Bacteria in Macro- and Microparticles 71

-E

F

Fig. 1. The spray chamber used to make very small microparticles (see refs. 1,ll). (A) Vent line to glass wool-packed syringe “filter”; (B) Gas line (from nitrogen or ah-tank and regulator) that delivers pressure at -10 psi; (C) low pressure fogging nozzle; (D) stir bar; (E) gas line (from nitrogen or air tank and regulator; note in-line 0.2~pm filter); (F) peri- staltic pump tubing that delivers alginate/cell suspension at -15 mL/min; (G) peristaltic pump tubing that delivers alginate/cell suspension to the low-pressure fogging nozzle.

4. Sterile 20-L Pyrex carboy (Corning) that can tolerate slight pressurization. (This is the “spray chamber”; Fig. 1).

5. Low-pressure fogging nozzle, model #052H (Sonic Environmental, Parsippany, NJ) (Fig. 1C).

6. Peristaltic pump and tubing and fittings: connectors, sterile 0.2qm in-line air filter for the nitrogen or air gas tubing).

7. Pressurized nitrogen gas or air, with appropriate regulator. 8. Sterile large-volume separatory funnels (2 L).

72 Knaebel, Stormo, and Crawford

9 Stenle HEPES lmmoblhzatlon buffer (HIB)* 50 n-&I HEPES, pH 7 3, 1 mM CaCl,, 1 mA4 MgS04, 20 /.U4 FeSO, This buffer IS used for Flavobactenum sp , the required components are the HEPES and CaClz

10 Lammar flow hood 11 Sterile l- or 2-L beakers 12 Sterile water (100 mL) in sterile container. 13 Three-hole stopper to fit the mouth of the carboy. Holes should allow for a snug fit

of the two pieces of tubing to the foggmg nozzle and the tubing for the exhaust vent 14 Magnetic stmer 15 Large sterile funnel that fits mslde mouth of separatory funnel (item 8 above) 16 Sterile nylon mesh (-1 mm) for removing clumps of microbeads This mesh IS

placed mslde the large funnel (Item 15). Several sterile pieces of this may be useful m case one or more get clogged.

17. Fifty-mllhhter plastic syrmge with plunger removed. The chamber of the synnge is packed loosely with glass wool. The Luer lock end 1s attached to the vent line tub- ing. The syrmge 1s then placed m a large Erlenmeyer flask with the chamber open- mg pointing down, and glass wool IS packed gently around the syrmge body. This provides a system to remove aerosohzed microbeads that escape the chamber through the vent lme.

3. Methods

3.1. Agarose/Oi/ Emulsion Technique for Microparticle (50-200 ,um Diameter) Immobilization

1 Grow bacterial cells (e g., Pseudomonas putzda mt-2, Flavobactenum sp., E cob) to mid- or late log phase, centrifuge, and resuspend m an appropriate buffer Wash the cells a second time and resuspend them in a volume of buffer equal to that of the molten agarose solution. To lessen heat shock problems, slowly warm the cells to the temperature of the agarose solution (see step 2)

2 Prepare and autoclave an agarose solution m a 125mL Erlenmeyer flask that con- tams a stir bar The agarose IS made up to a 2X concentration in the buffer used for cell resuspension. Autoclave the solution and keep it molten (45-50°C) or reliqulfy It by heating Immediately before use, but cool It to 45-50°C before use A target concentration of l-2% agarose 1s commonly used

3 Autoclave the large volume (600-800 mL) of food-grade canola oil m large- capacity bottle (1 L) (see Section 2.1 3 ). This is placed water bath at 45°C

4 Assemble the mixer, impeller, and shaft m the lammar flow-hood using sterile tech- nique. Arrange the impeller/shaft assembly m the beaker so that the impeller IS 3-5 cm above the bottom of the beaker Place the bottom of the beaker m a plastic tub, which will be carefully filled with Ice after step 8

5. Pour the warm canola 011 into the beaker and set the mixer motor speed to approx 700-800 rpm (see Note 1) Ensure that the temperature of the 011 bath 1s mamtamed at 45-50°C so that the agarose/cell suspension does not solidify on contact when It 1s added (see step 7)

Bacteria in Macro- and Micropartdes 73

6. Slowly warm the cell suspension to the same temperature (45-50°C) as the agarose, and mix the two solutions.

7 Pour the agarose/cell suspenston mtxture mto the oil bath; adlust the speed of the impeller to promote the formation of particles of the desired size (-50-100 pm) Stze can be momtored by sampling and microscopic examination.

8. After approx 10 min, carefully place ice around the beaker; add chilled water (if desired) to improve heat transfer. As the oil solution is chilled, the agarose/cell particles harden. Contmue mixing until the desired temperature is reached for the sohdificatton of the agarose particle. Generally, the process is stopped when the oil bath reaches -lO-15°C The viscosity of the oil/agarose suspension will increase as the temperature decreases. Therefore, increase the speed of the mixer to mamtain adequate shear,

9 Allow the particles to settle and then decant the oil aseptically The 011 can be fur- ther removed from the agarose microbeads by gentle suctton-filtermg through a glass fiber filter. Aseptically transfer the particles to 500-mL centrifuge bottles Add the PBS/Tween 80 solution and vortex gently Centrifuge the suspension at low RCF Aspirate the supernatant and repeat the PBS/Tween 80 washes until no oil sheen is visible on the surface of the supematant Usually, wash twice more with PBS/Tween 80 to ensure removal of the oil, and then wash wtth PBS alone to remove the Tween 80 (see Note 2).

3.2. Calcium Alginate “Drop” Technique for Macroparticle (-2 mm Diameter) Immobilization

1 Dissolve the 6% sodium algmate (type VII) in an appropriate buffer (e.g , HEPES buffer, phosphate and citrate buffers should not be used) Stertlize the solution by autoclavmg. Generally, the alginate will solubdize completely following extended heating (steammg) or autoclavmg.

2 Resuspend the cells m an equal volume of same buffer used for the alginate disso- lution and mix the two solutions using aseptic technique Adequate mixmg of the vrscous alginate/cell solution can be ensured by autoclavmg a stir bar with the algmate solution.

3. Arrange the autoclaved peristaltic pump tubing m the pump head, place one end of the sterile peristaltic pump tubing in the alginate solution, and suspend the other end (with the sterile capillary) at an appropriate distance above the CaC& receiving bath (usually 5-10 cm). Adjust the magnetic stirrer so that the receiving bath is mixing slowly, and adJust the pertstaltic pump flow rate so that the alginate/cell mixture exits the tubing as discrete droplets The droplets should solidify immediately on contact with the CaCl* solution. Once all the algmate capsules are made, the solu- tion should be slowly stnred for 30 mm to 1 h to ensure complete sohdificatton of the algmate capsules.

4. Wash the capsules with a fresh volume of HEPES buffer containing l-5 mM CaC& and use the microparticles for experimentation (see Note 3)

5 Determine cell loadings by directing droplets to 50 mM phosphate buffer dilution blanks, allow the parttcles to dissolve, and enumerate the cells on an approprrate medium

74 Knaebel, Stormo, and Crawford

3.3. Low-Pressure Fogging Nozzle Technique for Microparticle (2-100 pm Diameter) Immobilization in Alginate

1 Prepare a solution of 6% sodmm algmate (type VII) m a smtable buffer and auto- clave it HEPES buffer 1s recommended. (See step 1 of Section 3 2 )

2 Assemble the spray chamber (see Fig. 1) Fill the chamber with 4 L of 50 mM CaCl* and place a large stir bar m the carboy. Thread the tubing through the three-hole stopper and connect it to the fogging nozzle (Fig 1C) The gas line (with the in-line, 0 2+m filter) from the an or mtrogen tank (Fig 1B) enters the nozzle at the top port The feed line from the pertstaltic pump (Fig 1 G) enters the nozzle at the side port. Use enough tubing to ensure adequate lengths for connection to the peristaltic pump and to the nitrogen or an source. Adjust the tubing so that the nozzle IS sus- pended approx 10 cm from the surface of the CaClz solution and 1s perpendicular to the surface of the solution The vent line (Fig 1A) consists of tubing connected to a large-capacity disposable syringe body (without the plunger) that is loosely packed with glass wool (see Section 2 3 , item 17).

Autoclave the entire assembly (spray chamber, nozzle and associated tubing, and CaC&). Cover the tubing ends and the ltd assembly with aluminum foil for autoclavmg Ensure that the nozzle does not touch the CaCl, solution during auto- clavmg (see Note 6)

3 Resuspend the cells m an equal volume of the HEPES buffer and mix with the algr- nate solution Adequate mixing of the viscous algmate/cell solution can be ensured by autoclavmg a stir bar with the algmate solution

4 Place the spray chamber on a magnetic stirrer with a large surface area to support the carboy. Turn on the mixer so that the solutton is slowly mtxed (a small dimple should form above the stir bar)

5 Connect the tubing to the nitrogen or an tank 6 Connect the Luer end of the syrmge to the vent lme and direct the opening down-

ward in an empty Erlenmeyer flask Pack additional glass wool loosely around the outside of the syringe and the msrde edge of the Erlenmeyer flask mouth.

7. Using aseptic technique, place the tubing for the algmate/cell mixture in the sterile water Turned on the penstalttc pump to a flow rate of 15 mL/mm When the water is observed to approach the nozzle, turn on the nitrogen or au supply to a pressure of 10 psi, which will result m the formation of a small dimple on the surface of the CaClz solutton Turning on the nitrogen or air supply at this time prevents problems with backpressure that may occur if the gas is turned on too early The water rinses the nozzle and tubing free of any CaCl, that may have entered the nozzle during handling or autoclavmg Do this for a short penod of time (lo-20 s) Turn off the pump and an or nitrogen regulator.

8 Aseptically place the free end of the tubing m the algmate/cell solutron. Adjust the peristaltic pump flow rate to a rate of 15 mL/min When the alginate/cell suspension 1s observed to approach the nozzle, turn on the nitrogen or au supply to a pressure of 10 psi. Once the algmate/cell solution leaves the ortfice of the nozzle, a fme mist of microbeads should form Once these settle mto the CaCl, solution, they harden and smk Some clumping will probably occur, but this should be mmimal

Bacteria m Macro- and M/cropatixles 75

Clumping can be decreased or avoided by optimizing the stirring speed, the gas pressure, and the peristaltic pump flow rate. Once all the algmate capsules are made, the solutton should be slowly starred for 5-10 mm to ensure complete sohdification of the algmate capsules

9 Transfer the spray chamber to the lammar flow hood. Aseptically remove the stop- per and nozzle from the mouth of the carboy, and aseptically decant the suspension mto the sterile beakers. Pour the suspension aseptically through a funnel (contam- mg the sterile large-mesh [-1 mm] nylon screen) into the sterile separatory funnels Permit the microbeads to settle, usually overnight at 4°C Some size fractionation of the microparttcles can be accomplished by carefully draining off the larger, more quickly settling microparticles through the funnel spigot

10. Transfer the settled beads to the sterile wide mouth bottles and rinse with HIB use for experimentation (see Notes 7 and 8)

4. Notes

4.1. Agarose/Oil Emulsion Technique for Microparticle (50-200 pm Diameter) Immobilization

1. Determine the appropriate mixing speed empirically to maximize shear, but mtm- imize air bubble formation (in the oil) That is, adJust the speed of the mixer so that the vortex created by the mixer is substantial, but so that no an is drawn mto the impeller (which will form air bubbles) Since the viscosity of the oil changes substantially with temperature, the mixing speed will need to be adjusted during the cooling. The Impeller type (propeller-like vs high-efficiency types) ~111 also influence the necessary mixing speed for vortex formation

2 The most dtfficult part of this process is the complete removal of the oil from the microparticles Other oils (mineral oils, different vegetable oils) have provided little improvement (or impairment) of oil removal It should be considered that some oils may adversely affect subsequent metabolic activittes of the cells Other biocompatible solvents may be tried.

4.2. Calcium Alginate “Drop” Technique for Macroparticle (-2 mm Diameter) Immobilization

Phosphate and citrate buffers should not be used for formulation or storage of the capsules because they disrupt the ionically crosslmked Ca-algmate polymer HEPES buffer offers some good biological compatrbility characteristics. It can be used for the suspension of the alginate and cells, and m receiving bath, it can be amended with the appropriate concentration of CaCl* to cause the algmate to gel To mamtam the solid algmate polymer, include a low concentration of CaClz (l-5 mM) m any buffer or solution used for storage of the particles. The Ca-algmate in the macroparticles may be covalently crosslmked to improve sta- bility The surface of the microcapsule (or the entire matrix) can be crosshnked using a polyethylenimme/glutaraldehyde posttreatment, based on the method of Bnnbaum et al (26) Expose the Ca-algmate (2-3%) capsules to a solution of 0 5%

76 Knaebel, Stormo, and Crawford

polyethylemmme-HCl and 50 mM CaC& pH 7 0, for 24 h at either room tempera- ture or 4°C (the suspension should be slowly stirred during this time) Wash the cap- sules brlefly m sterile water, and then expose them to 1% glutaraldehyde m 10 mM phosphate buffer, pH 7 0, for 1 mm. Decant the solution and wash the beads W&I stenle water These crosshnked capsules are more resistant to dIsruptIon by phos- phate or citrate buffers Greater crosshnkmg and less shrmkage of the beads occur at higher (500 n-&f) CaClz concentrations durmg the polyethylemmme stage (26)

5. Algmate provides a good matrix for lyophihzation and storage of the cells Rehydratlon of the capsules in a buffer contaming CaCI, permits them to be reformed, although m a deformed shape

4.3. Low-Pressure Fogging Nozzle Technique for Micropatticle (2-100 ,um Diameter) immobilization in Alginate

6. If the nozzle IS Immersed durmg the autoclavmg, the solution will be drawn out of the carboy during the cooling stage of the autoclavmg, because of pressure changes

7 Determine cell loadings by ahquoting a known mass of the mlcrobeads to an equal volume of 200 mM phosphate-buffered saline After dtssolutlon of the algmate, mix the solution and perform enumerations Determme dry weights on the microbeads

8. The algmate mlcrobead matrix may be crosslmked in a fashion slmllar to that described m Note 4, but we have not done this Shorter exposure times are recom- mended for each of the solutions, because these particles are orders of magmtude smaller than the macrobeads Also, the glutaraldehyde used to crosslmk the poly- ethylenimine may be toxic to the cells at the recommended concentration, and If they are exposed for too long a time.

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waste cleanup and treatment with encapsulated and entrapped mlcroorgamsms, m BloEoggcal Degradation and Bloremedlatlon of Toxic Chemicals (Chaudhry, G. R , ed ), Dloscondes Press, Portland OR, pp 455-469

2 Colwell, R R., Levm, M. A, and Gealt, M A. (1993) Future directtons m blore- medlatlon, m Blotransformatlon of Industrial and Hazardous Waste (Levm, M A and Gealt, M. A., eds.), McGraw Hill, New York, pp 309-321.

3 Crawford, R L., O’Reilly, K T , and Tao, H.-L (1990) Microorganism stablhza- tlon for in sttu degradation of toxic chemicals, in Biotechnology and Biodegradatzon (Kamely, D , Chakrabarty, A, and Omenn, G S., eds), Gulf Co, Houston, TX, pp 203-211

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5 Kolot, F B (1988) Immobilized mlcroblal systems prmciples, techniques and mdustrlal apphcatlons. R. E Krueger, Malabar, FL

6 Akm, C. (1987) Biocatalysls with lmmoblhzed ceils Bzotech. Genet. Engin Rev 5, 3 19-367.

Bacteria in Macro- and MicropatTdes 77

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10 Tanaka, H , Ohta, T , Harada, S., Ogbonna, J C., and YaJima, M (1994) Development of a fermentation method using immobihzed cells under unsterile conditions. 1. Protection of tmmobihzed cells agamst anti-microbial substances Appl. Microbtol. Btotechnol 41,544-550.

11 Stormo, K. E. and Crawford, R. L (1992) Preparation of encapsulated microbial cells for envuonmental applications. Appl Environ. Mzcrobtol 58,727-730

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13 King, G A and Goosen, M F. A. (1993) Cell immobilization technology an overvrew, m Fundamentals of Animal Cell Encapsulatton and Immobtltzatton (Goosen, M. F A, ed.), CRC, Boca Raton, FL, pp. l-6.

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15. De Taxis du Poet, P., Dhulster, P., Barbotin, J -N., and Thomas, D. (1986) Plasmid mheritabihty and biomass production: comparison between free and immobilized cell cultures of Escherichza coli BZ18 (pTG201) without selection pressure 1 Bacterial 165,871-877.

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20 Omar, S. H. (1993) Oxygen diffusion through gels employed for immobihzatton 2 In the presence of microorgamsms. Appl. Mzcrobtol. Biotechnol. 40, 173-l 8 1

21 Omar, S. H. (1993) Oxygen diffusion through gels employed for immobihzation 1 In the absence of mtcroorgamsms. Appl. Microbtol. Bzotechnol. 40, I-6.

78 Knaebel, Stormo, and Crawford

22 Bettman, H and Rehm, H -J (1984) Degradation of phenol by polymer entrapped mrcroorgamsms Appl Mzcrobzol Biotechnol. 20,285-290.

23 Etkmeier, H , Westmeier, F , and Rehm, H -J. (1984) Morphologtcal development of Aspergzllus nzger tmmobrltzed m Ca-algmate and K-carrageenan Appl Mzcrobzol Biotechnol. 19,53-57

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