Bioprocessing Aids in the Recovery of Proteins from Biomass

Chapter 12

Bioprocessing Aids in the Recovery of Proteins from Biomass 1,3

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Ian T. Forrester , Anthony C. Grabski , Mark N. Shahan , and Kathleen Fletcher 2

Downloaded by CORNELL UNIV on September 6, 2016 | http://pubs.acs.org Publication Date: April 30, 1991 | doi: 10.1021/bk-1991-0460.ch012

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University of Wisconsin Biotechnology Center, 1710 University Avenue, Madison, WI 53705 TosoHaas, Independence Mall West, Philadelphia, PA 19105 2

The extraction of enzymes from biological tissues has been examined using a three step process involving an initial clarification, employing bioprocessing aids such as the Biocryls to remove non-protein contaminants, followed by ultrafiltration for concentration and then a final chromatographic purification. DNA, RNA, cell-wall debris and pigmented compounds in particular are eliminated by this process. The method has been used for bacterial, plant, fungal and animal tissues. The inherent complexity and chemical diversity of biomass predicts the recovery and purification of proteins from such a source will be an extremely challenging task. We have been studying this problem with a long-term research goal of identifying and establishing experimental conditions which may be applicable for protein purification from all biomass systems. Our approach has been to divide protein purification into three distinct steps. These being Step I, the collection and clarification of extracts from the original biomass; Step II, the concentration of the clarified stream; and Step III, the chromatographic purification of the specified protein. It has been our experience that Step I, in the removal of either non-protein contaminants or superfluous proteins, is critical to the design and effectiveness of the latter two Steps. As to be expected, different types of biomass contain widely varied non-protein contaminants. However, these components are commonly; particulate debris, pigmented organics, and biochemicals such as oligosaccharides, lipids, pyrogens and nucleic acids (1-3). The downstream effects of these types of contaminants assumes greater importance when the biomass processes are transformed into production-level systems. Various chemical agents, including the Biocryls, polyethylenimine and chitosan, have been under study in our laboratory as so-called bioprocessing aids in Step I, to remove the contaminating species found in biomass. In this paper we report on the application of the three step approach to protein purification in two different biomass systems (i) a bacterial homogenate and (ii) a plant homogenate. We also discuss the advantages which accrue, in each case, from the incorporation of bioprocessing aids into Step I, an experimental strategy for evaluating the adaptation of bioprocessing aids into a protein purification protocol, and the general utility of these techniques to protein recoveryfrombiomass. 3

On leave from the Department of Biochemistry, University of Otago, Dunedin, New Zealand 0097-6156/91/0460-0152$06.00/0 © 1991 American Chemical Society

Leatham and Himmel; Enzymes in Biomass Conversion ACS Symposium Series; American Chemical Society: Washington, DC, 1991.

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Bioprocessing Aids The researcher wishing to study and characterize the mix of enzymes associated with the conversion of a specific biomass is often confronted with a familiar problem. The key enzymes of interest are present only in very minute amounts and are extremely intractable to separationfromthe overwhelming mass of chemically-complex biomass. In our laboratory, we confronted this problem during the recovery and characterization of the extracellular enzymes produced by a white-rot fungus, Lentinula edodes, grown on a commercial wood medium. More complete details of the issues involved in that program are presented in this monograph (4) and elsewhere (7,5-7). Suffice to say, the key to the success of the L. edodes program was the utilization of chemical agents, such as the Biocryl BPA-1000 or polyethylenimine (PEI), to remove tannins, polyphenols and debris from the wood extracts. We now have extended this line of research to examine a broad range of biomass systems, using various chemical agents for enhancement of bioprocessing and enzyme recovery. Selective precipitation of a protein from a biomass stream, although a desirable method for purification of an enzyme (8), is often an impracticable option due to the high concentrations of non-protein contaminants. In considering that problem, we have adopted a different approach, that is to introduce a procedure -ideally very early in the biomass processing system - which may remove selectively the non-protein components. The procedure we sought to satisfy those criteria was the addition of a chemical agent to the crude biomass in Step I, resulting in flocculation and selective precipitation of the undesirable components, but at the same time leaving the target protein (enzyme) in the resultant supernatant phase. This general class of chemical agent is defined as a bioprocessing aid. Some of the chemical and physical properties of compounds which have been investigated as potential bioprocessing aids are summarised in Figure 1. Other materials which could be classified as bioprocessing aids include; polyvinylpyrrolidone, the Whatman CDR product, as well as Chitoplex and Evalsan (9,70). The application of PEI as a bioprocessing aid in protein purification has been covered in a comprehensive review by Jendrisak (77). Although in some situations PEI does initiate the flocculation of non-protein contaminants in Step I, without simultaneous precipitiation of the target protein, in most cases the PEI causes coincident precipitation of the protein with nucleic acid, from the crude biomass. Further careful processing is then necesssary to selectively dissociate the protein from the complex nucleic acid aggregation. This approach does not satisfy the conditions we sought for Step I. Chitosan, which is derivedfromchitin by deacylation, has been of limited use in bioprocessing (72), it is available commercially in various size ranges. The form we have found most useful has an average molecular weight of -750,000. Chitosan like PEI however, is an example of a water-soluble, cationic polymer of high molecular weight and charge density which initiates the flocculation of nucleoprotein complexes. The need for chitosan to be solubilized in acidic conditions may also restrict its widespread application for biomass processing. Chemical agents which more closely approach the criteria we sought for use in Step I, are the Biocryl bioprocessing aids (BPAs). These materials, are crosslinked acrylic or styrenic polymer particles with diameters of 0.1 or 0.2 microns. Surface functional groups are strongly basic quaternary ammonium (BPA-1000), weakly basic tertiary amine (BPA-1050) or strongly acidic sulfonic acid (BPA-2100). These charged particles adsorb oppositely charged species such as cell debris, nucleic acids, contaminant proteins, color bodies and enzymes in a wide variety of fermentation broths, cell or tissue extracts and homogenates. A high molecular weight, water soluble formulation is also available. This material, BPA-5020, has a weakly basic tertiary amine functionality. In some situations it may act synergistically with another polycationic bioprocessing aid such as BPA-1000 (3).



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Unlike water soluble polymers such as PEI or chitosan, the particulate BPAs are shear stable and pumpable. The particles, along with the adsorbed materials, can be effectively removed from the suspending medium by either membrane filtration or centrifugation. The BPAs have been designed to have specific functional groups on the surface and in the case of the strong anionic and cationic species, maintain their charge over a broad pH range. When used in conjunction with membrane filtration (13) they reduce the development of a concentration polarization layer on the membrane surface resulting in enhanced flux rates. When an appropriately sized membrane is chosen, the particulate BPAs are maintained in the retentate producing a product stream free of the filter aid. Given the broad pH range over which the BPAs can be used, it is also possible to specifically target a product protein for removal with the BPA, leaving the contaminants behind. This is accomplished by adjusting the pH of the medium so that the target protein alone (or with few contaminants) is bound. The product bound BPA can then be removed by centrifugation or in ultrafiltration it can be eluted and concentrated in a single step. Materials and Methods Growth and Lysis of Escherichia coli. The E. coli strain used in this work was RAJ201 harboring the plasmid pRAJ255 (14). This plasmid contains the gene for βglucuronidase, uidA, under the control of the lac promoter and operator (74). Cultures of E. coli were grown in a 10 L fermenter on 2YT, induced with IPTG at 8 hours (O.D.550 = 5), harvested at 23.5 hours (O.D.550 = 8), and the cell paste stored frozen at -70° C until they were to be lysed. Upon thawing of the cell paste the material was suspended in phosphate lysis buffer A [20 mM potassium phosphate (pH 7.1), 10 mM 2-mercaptoethanol] at a concentration of 1.0 g wet weight cells in a total volume of 5 ml. The bacteria were lysed by a double passage through a Gaulin homogenizer at 10,000 psi, with cooling of the cellular material in a dry icerethanol bath after each passage. Lysis was estimated to be >95% by phase contrast light microscopy. The resulting E. coli homogenate was stored, until required, at -70° C. Alfalfa Lysis. Whole alfalfa (Medicago sativa) plant tissue was broken and cell rupture was by shear in a rotary device. Juice expression was by a batch type press, as described elsewhere (77). The resulting alfalfa extract was stored, until required, at -20°C. Fractionation and Analysis of Extracts Extract Preparation. The E. coli extract was diluted four-fold with buffer A and the alfalfa extract was diluted 2.5 fold with 50 mM sodium phosphate buffer pH 7.1. The pH of the diluted alfalfa juice was adjusted with 1 M NaOH to a final pH of 7.1 before proceeding with the next bioprocessing step. Step I: Standardized BPA Assessment Protocol. A 1.0-1.2 ml aliquot of extract was added to each of a series of plastic microfuge tubes (No. 72.690 Sarstedt, W. Germany). The series was titrated with BPA-1000 or BPA-1050[TWS-3088B] (TosoHaas, Philadelphia, PA). Addition of 5-100μ1 of BPA is usually sufficient but greater quantities may be required depending on the nature of the extract and degree of contamination. The BPA treated extracts were mixed thoroughly, allowed to react for 5 minutes [0°C for E. coli extract, 22°C for alfalfa extract], and centrifuged (700-1000 g for 2 min). Pellets were then discarded and the supernatant was assayed for turbidity, enzyme activity [β-glucuronidase (GUS) or peroxidase activity], protein, RNA, and DNA. This standardized BPA assessment protocol is shown schematically in Figure 2.



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Physical Form

Functionality

Structure

Size

Stable Suspension 10% Solids Water Soluble Polymer 2% Solids Stable Suspension 10% Solids Stable Suspension 10% Solids Stable Solution 10% Solids Stable Solution 1% Solids

Quaternary Amine

~0.1 micron

Tertiary Amine

High MW >200,000

Tertiary Amine

~0.1 micron

Sulfonic Acid

0-@-SO H

~0.2 micron

Secondary Amine

^N-J-C-C-N-j-C-Ç-

High MW 30-40,000

Primary Amine

3

Ιψ

G—f-G—4-G [NHj NH n

2

High MW 750,000

Figure 1. Characteristics of bioprocessing aids.

Extract Prepare Titration Series with BPA Incubate

1 Supernatant

Pellet

Quantitate Purification Parameters Figure 2. Schematic of the standardized BPA assessment protocol.


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Step II: Diafiltration. Extracts to be fractionated by HPLC were diafiltered into buffer A using Centricon 10 microconcentrators (Amicon Div., Grace & Co., Danvers, MA) and then passed through 0.22 um filters.


Step ΠΙ: HPLC. Samples (500 μΐ) of the concentrated, filtered extracts (1.7-3.6 mg protein) were applied (1 ml/min.) to a 7.5 mm χ 75 mm TSK DEAE-5PW column coupled to a gradient HPLC system consisting of a #210A sample injection valve, a #126 dual pump, a #165 dual-channel variable wavelength detector, and an IBM-PCbased data system/controller running System Gold software (Beckman Instruments, Inc., Waldwick, NJ). Elution buffers were buffer A and buffer A plus 1.0 M KC1 (buffer B). Protein elution and column re-equilibration was achieved with a gradient of buffer A/B [% (v/v)A/min]: 100/0; 100/5; 20/45; 0/46; 0/48; 100/50; 100/65; using a flow rate of 1 ml/min and collecting 1 ml fractions. Protein elution was monitored at 280 nm. Assays. The turbidity of E. coli extracts and alfalfa extracts was quantitated spectrophotometrically at 600 nm and 680nm respectively. GUS activity was assayed as outlined by Novel and Novel (75), except that activity was monitored at 400 nm (ε° = 9.6 χ 10 ). Peroxidase activity was determined at 37°C with 0-tolidine using a final concentration of 0.67 mM 6>-tolidine*HCl (Sigma Chemical Co., St. Louis, MO) and 1 mM H2O2 in 20 mM sodium phosphate buffer pH 7.1. The change in absorbance was measured at 600 nm, ε ° = 6.34 χ 10 (18). Protein was determined by the biuret reaction; RNA by the orcinol method; and DNA by the diphenylamine method (76). 3

3

Electrophoresis. Electrophoresis was performed on a Pharmacia Phast System (Piscataway, NJ). Protein purity was determined by sodium dodecyl sulfate (SDS) polyacrylamide gel electrophoresis (PAGE). A Phast System 10-15% gradient polyacrylamide gel was run and stained with Coomassie brilliant blue G250 according to standard Phast System methods. Protein mobility was compared to an SDS-7 standard (Sigma Chemical Co., St. Louis, MO), containing seven proteins in the molecular weight range of 14,000 to 66,000. Results BPA Treatment of E. coli Extracts. The purpose of these experiments was to develop a general approach to Step I of our protein purification scheme, as applied to bacterial homogenates. This goal was considered particularly relevant given the importance of bacterial systems as a biomass as well as a valuable resource for enzyme recovery. In our particular case, we sought to develop a simple and efficient clarification procedure for bacterial homogenates (using E. coli as the test biomass) that would remove, ideally, cell debris and nucleic acids while leaving the enzyme of interest in the solublefractionof the clarified extract. For our program GUS was used as the target enzyme because of the ease with which it is assayed, because of its stability, and because of the common use of GUS as a genetic marker in plant transformation systems. Previously, PEI had been used to precipitate proteins such as RNA polymerase along with the nucleic acids (77). In that approach to bioprocessing the desired protein was differentially releasedfromthe nucleoprotein pellet by step-wise batch extractions with buffers of increasing salt concentration. It would appear that such an extraction adds a time consuming and potentially expensive step to a large scale protein purification. Therefore, it was considered desirable to develop a method that eliminated the salt extraction step yet still resulted in a clarified extract suitable for use directly in subsequent purification steps. Our approach to the experimental problem was based on the concept of varying the


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pH and/or ionic strength to prevent binding of the target enzyme to the BPAs while simultaneously inducing aggregation and precipitation of the cell wall debris and nucleic acids. The first step in this process was to titrate the E. coli extract with the BPA at neutral pH, in the absence of exogenous salt, to determine the extent of separation of GUSfromnucleic acids and cellular debris which could be achieved. It was determined that at a BPA-1000 concentration of 4000 ppm, nucleic acids were almost completely removed and turbidity was reduced by approximately 50% (Figure 3). Unfortunately in these experiments, the GUS activity was found to coprecipitate with the nucleic acids as did 75% of the total cellular protein. Increasing the amount of added BPA-1000 continued to reduce the turbidity and the amount of protein. The clearing effect of increasing amounts of BPA-1000 on the E. coli extract is shown in Figure 4. Also of note was the steady increase in the size of the pellet with increasing additions of BPA1000. Similar results were obtained with BPA-1050 except that a final concentration 2000 ppm higher than for BPA-1000 was required. Effect of pH, salt and mixing sequence. It was ascertained that changes in pH were unsuccessful in achieving differential precipitation of GUS and nucleic acids. At pH 5.5, GUS activity and nucleic acids were less well separated than at pH 7.1. Utilization of more alkaline conditions was of restricted value since at pH 9.0 GUS was unstable. Only at pH 8.0, and only with BPA-1050, was some separation possible between GUS and nucleic acids precipitation. In view of the limited benefits of this experimental approach, we decided to investigate the effect of exogenous salt on the flocculation by the Biocryls of GUS in the E. coli extracts. At pH 7.1, using a constant BPA-1000 concentration of 4000 ppm, but varying exogenous KC1 between 300 mM and 500 mM, it was observed that nucleic acids were almost completely removed from the extract while greater than 95% of the GUS activity remained in the supernatant (Figure 5). Almost identical results were obtained with BPA-1050 except that the optimum BPA concentration was higher, requiring 6000 ppm BPA-1050. Subsequently a sample of the untreated E. coli extract and the extract processed with BPA-1000 and exogenous 300 mM KC1 were analyzed separately by HPLC. The GUS activity in both samples was found to elute from the DEAE column at about 300 mM KC1 and furthermore the SDS-PAGE analysis showed that samples contained approximately 50% of protein as GUS (Figure 6). The SDS-PAGE gel also showed that in the unfractionated extracts, prior to HPLC, the BPA-1000 treatment (in the presence of 300 mM KC1) does not decrease the amount of GUS compared to the crude E. coli extract untreated with BPA. It was also noted during the course of the exogenous KC1 addition studies that the sequence in which the salt, dilution buffer, extract, and BPA were added had a profound effect on the turbidity of the resulting extracts. Initially, experiments were performed by adding the BPA to the diluted extract. We found subsequently that a five fold decrease in turbidity could be achieved by mixing diluting buffer, salt, and BPA first, and then adding extract to this mixture (Figure 7). These results were surprising and suggest that dilution and ionic environment may influence the ability of positively charged BPA to react with particulate material such as the cell wall and lipid components present in bacterial cell debris. Further studies will be required to fully investigate this interesting observation. By comparison the altered mixing protocol decreased protein content slightly but had no effect on GUS activity or nucleic acid content (Figure 7). Using the new protocol we studied the effect of centrifugal force on achieving clarification of the untreated and BPA-1000 (or BPA-1050) treated E. coli extracts containing 300 mM KC1. As centrifugal force increased, the turbidity of the resultant supernatant phase for both the untreated and BPA treated samples decreased (Figure 8A). However, the turbidity in the BPA treated samples was 5-10 fold lower than in the untreated sample at all centrifugal forces tested. In fact, the turbidity in the treated samples at the lowest centrifugal force (-80 g) was less than that in the untreated



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BPA1000 (ppm) Figure 3. Titration of E. coli extract with BPA-1000. Results are expressed as a percentage of those for the untreated extract and are plotted against the added BPA-1000 expressed in parts per million (ppm). Legend: • = turbidity; Ο = GUS activity; • = protein; A = RNA; A = DNA.



Figure 4. Photograph of E. coli extract titrated with BPA-1000 as in Figure 3. The BPA-1000 concentrations from left to right are 0, 1000, 2000, 3000, 4000, 5000, 6000, and 7000 ppm.



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KC1 (mM) Figure 5. Clarification of E. coli extract in the presence of increasing concentrations of KC1. Results are expressed as a percentage of those for the untreated sample (0 mM KC1). Legend: Ο = GUS activity; A = RNA; A = DNA.


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Figure 6. 10-15% SDS polyacrylamide gel of: lane 1 — untreated E. coli extract (0 mM KC1); lane 2 — the peak GUS fraction from the HPLC separation of untreated extract; lane 3 — Sigma SDS-7 molecular weight standards (approximate molecular weights of 4 standards are indicated on the right hand edge of the figure); lane 4 — BPA-1000 treated E. coli extract (4000 ppm BPA1000 and 300 mM KC1); lane 5 — the peak GUS fraction from the HPLC separation of BPA-1000 treated extract. GUS migrates slightly slower than the 66,000 molecular weight standard.




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o

Turbidity Protein GUS Assay

RNA

DNA

Figure 7. The effect of the mixing order in the BPA treatment protocol on the clarification parameters. Results are expressed as a percentage of those for the untreated sample. All samples contained 300 mM KC1. BPA-1000 was used at a concentration of 4000 ppm and BPA-1050 was used at a concentration of 6000 ppm. Legend: • = BPA-1000 treated extract, BPA added last; m = BPA-1050 treated extract, BPA added last; • = BPA-1000 treated extract, extract added last; • = BPA-1050 treated extract, extract added last



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60 40 20

OU— 80 990 400016000 RCF(g) Figure 8. The effect of centrifugal force in the BPA treatment protocol on the clarification parameters. Results are expressed as a percentage of those for the untreated sample spun at 16,000 g. All treatments involved 2 min centrifugation. The KC1, BPA-1000, and BPA-1050 concentrations are the same as in Figure 7. A: turbidity versus relative centrifugal force (RCF); B: protein versus RCF; C: RNA versus RCF; D: DNA versus RCF; E: GUS activity versus RCF. Legend: • = untreated extract; • = BPA-1000 treated extract; • = BPA-1050 treated extract


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sample at the highest force (16,000 g). Protein content in the untreated samples decreased in total by approximately one third as the centrifugal force increased, but the protein content in the treated samples was largely unchanged by centrifugal force (Figure 8B). As indicated in Figures 8C, D and E, the RNA and DNA content, and the GUS activity, in contrast to turbidity, are largely unaffected by the centrifugal force. BPA-1000 Treatment of Alfalfa Alfalfa juice as expressed directly from whole M. saliva tissue is an extemely dark green viscous solution, high in protein content as well as undesirable components such as pigments, secondary products and the phenolic group of flavonoids and phenylpropanoids. Previous large-scale processes for the recovery of protein from alfalfa juice have involved heat precipitation (30-80°C) and coagulation or acid precipitation of the proteins (20). However, heat or acid treatment is not feasible in most cases where biologically active protein (enzymes in particular) are to be purified from the plant. Therefore, we examined the feasibility of using bioprocessing aids to selectively precipitate the plant tissue contaminants (consistent with the Step I processing strategy) with the added goal of leaving the desired protein(s) in solution. It was also hoped this goal could be achieved under conditions suitable for the maintenance of enzymatic activity. For our investigations, peroxidase was selected as the target enzyme since alfalfa tissue contains several peroxidase isozymes (79) and a simple procedure has been established to assay that enzyme (5,18). The results of BPA-1000 treatment of the alfalfa extract are summarized in Figure 9. Clarification of the extract could be assessed very effectively using spectrophotometry, due to the distinctive absorbance peak of the plant pigments at 680 nm. With increasing additions of BPA-1000, the absorbance at 680 nm was reduced to less than 5% of its untreated level indicating extremely efficient removal of pigment and debris (Figure 10). The concentration of nucleic acids remaining in solution after BPA-1000 treatment was also significantly reduced. At ~3000 ppm the amount of RNA remaining in solution was less than 55% of that in the untreated sample and DNA had been reduced to only 3% of its initial concentration. The differential in the effectiveness of RNA vs DNA precipitation could possibly be due to the formation of large DNA concatamers which increase the interaction and subsequent flocculation of DNA by the BPA. Although BPA-1000 effectively removed pigment, debris, DNA and to a lesser extent RNA, almost 90% of the total protein and greater than 100% of the peroxidase activity remained in solution at -3000 ppm BPA. It should be noted that our laboratory has frequently observed greater than 100% enzymic activity of a biological extract after treatment with BPA in Step I (2,5). It is possible that these higher levels of enzymatic activity could reflect the removal of specific enzyme inhibitors by the Biocryl. Conclusions We have found it possible to remove greater than 95% of the nucleic acids and 80-90% of the turbidity in E. coli extracts using BPA-1000 or BPA-1050. At the same time, greater than 95% of the target enzyme, GUS, remains in the clarified extract. This differential removal of debris, cell wall fragments and nucleic acids was accomplished by treating the extracts with BPAs in the presence of varying concentrations of exogenous salt. The obvious advantage of this procedure is a simplification over the procedure commonly used for PEI. Such a simplification could save considerable time and expense particularly when a system for the recovery of enzymes from bacterial biomass is transformed into a large scale, commercial operation. In addition, this procedure could prove generally useful for other bacterial proteins since greater than 90% of the nucleic acids are removed from the E. coli extracts over at least the concentration range of 0-500 mM KC1. Since proteins generally do not have the charge density of nucleic acids, it should be feasible to establish a salt concentration that



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BPA-1000 (ppm) Figure 9. Titration of alfalfa extract with BPA-1000. Results are expressed as in Figure 3. Legend: • = turbidity; Ο = RNA; • = DNA; Δ = protein; A = peroxidase activity.



Figure 10. Photograph of alfalfa extract titrated with BPA-1000 as in Figure 9. The BPA-1000 concentrations from left to right are 0, 475, 950, 1425, 1900, 2850, and 3800 ppm.



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prevents binding of the protein of interest to the BPA but facilitates binding, and flocculation, of the nucleic acids. Should salt alone not prove effective, a combination of salt plus changes in pH — either to the pi of the protein or to the pK of the BPA — may prove useful. Another advantage to the incorporation of bioprocessing aids into the Step I procedure is that the clarified extract can be used directly for subsequent purification steps even without the use of a Step II system to dewater or concentrate the process stream. These factors are especially relevant when HPLC systems are used in Step ΙΠ for the chromatographic procedures. Nucleic acids, pigmented organics and especially cellular debris can very quickly foul an HPLC column. This is an even more important consideration for large scale protein purification schemes where the volumes of material and costs of the operation are greatly increased (3). A final advantage to the use of bioprocessing aids in bacterial biomass processing is the low centrifugal forces needed to clarify the extract. Typically, untreated E. coli extracts are clarified by a 30,000 g centrifugation. After BPA treatment, the E. coli extracts are clarified better by an 80 g centrifugation than by a 14,000 g centrifugation of untreated extract. Centrifugal forces of 14,000 g or 30,000 g are easy to generate for laboratory-scale purifications but for industrial-scale purification strategies, where the processing stream is vast, such centrifugation systems would normally be impractical. The results obtained in this study indicate that bioprocessing aids in the Step I clarification of bacterial extracts could improve low speed centrifugation and therefore may offer adaptation for bacterial fractionation on an industrial scale. Based on the above advantages, we believe that incorporation of bioprocessing aids into Step I of a purification scheme could be used as a general approach to clarifying bacterial homogenates and the recovery of enzymes from bacterial biomass. Recovery of biologically active proteinsfromplant biomass is problematic due to the large quantity of cellular debris, nucleic acids, photosynthetic pigments and phenolic compounds (27). Removal of these contaminants in a Step I process is generally necessary in order to facilitate the subsequent bioprocessing of Step II and Step III, such as ultrafiltration and chromatography. The second series of experiments covered in this paper was designed to address those issues, employing a bioprocessing aid in Step I of a strategy for the recovery of enzymes from plant biomass. The experimental results which were obtained for peroxidase in alfalfa extracts indicate that bioprocessing aids are of considerable benefit in the clarification of crude plant extracts. This area of research is the subject of ongoing work by our laboratory. Although this report focuses on the processing of bacterial homogenates and plant extracts, all or part of the three step protein purification strategy, has been successfully applied by our laboratory to a variety of biomass sources. These sources include bacterial culture fluids (2), animal tissue extracts, animal secretions, animal tissue culture fluids (3), fungal culture fluids (7,5), fungal extracts (22), yeast extracts and yeast culture fluid (3). The full potential for the use of bioprocessing aids in the recovery of proteins from biomass has yet to be realized and further research in this area is continuing. Acknowledgments We thank W. Nick Strickland, Hans Liao, and Richard R. Burgess of the University of Wisconsin Biotechnology Center for their expert advice; Herbert J. Grimek of the UWBC Protein Purification Facility for assistance in the growth and breakage of the E. coli; Richard G. Koegel, USDA Dairy Forage Research Center and Richard J. Straub, UW Department of Agricultural Engineering for supply of the alfalfa extract and our colleagues in the UWBC for their ever-willing professional assistance. This research was supported by grants from the Rohm and Haas Company and USDA Midwest Plant Biotechnology Consortium.


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Literature Cited 1. Forrester, I.T.; Grabski, A.C.; Kelley, B.D.; Strickland, W.N.; Leatham, G.F.; Burgess, R.R. In Biotechnology in Pulp and Paper Manufacture; Kirk, T.K.; Chang, H-M., Eds.; Butterworths: London, UK, 1990, pp 407-414. 2. Forrester, I.T.; Grabski, A.C.; Strickland, W.N. In Fermentation Technologies : Industrial Applications; Yu, P-L., Ed.; Elsevier Applied Science: London, UK, 1990, pp 403-406. 3. Fletcher, K.; Deley, S.; Fleischaker, R.J.; Forrester, I.T.; Grabski, A.C.; Strickland, W.N. In Separations for Biotechnology; Pyle, D.L., Ed.; Elsevier Applied Science: London, UK, 1990, Vol. 2, pp 142-151. 4. Leatham, G.F.; Forrester, I.T.; Mishra, C. In this volume. 5. Forrester, I.T.; Grabski, A.C.; Mishra, C.; Kelley, B.D.; Strickland, W.N.; Leatham, G.F.; Burgess, R.R. Appl. Microbiol. Biotechnol. 1990, 33, 359-365. 6. Forrester, I.T.; Grabski, A.C.; Burgess, R.R.; Leatham, G.F. Biochem. Biophys. Res. Commun. 1988, 157, 992-999. 7. Mishra, C.; Forrester, I.T.; Kelley, B.D.; Burgess, R.R.; Leatham, G.F. Appl. Microbiol. Biotechnol. 1990, 33, 226-232. 8. Lightfoot, E.N.; Gibbs, S.J.; Cockrem, M.C.M.; Athalye, A.M. In Protein Purification Micro to Macro; Burgess, R.R., Ed.; UCLA Symposia on Molecular and Cellular Biology New Series; Alan R. Liss: New York, NY, 1987, Vol. 68; pp 75-97. 9. Muzzarelli, R.A.A.; Weckx, M.; Filippini, O. Carbohydr. Polym. 1989, 11, 293-306. 10. Muzzarelli, R.A.A.; Weckx, M.; Fiippini, O.; Lough, C. Carbohydr. Polym. 1989, 11, 307-320. 11. Jendrisak, J.InProtein Purification Micro to Macro; Burgess, R.R., Ed.; UCLA Symposia on Molecular and Cellular Biology New Series; Alan R. Liss: New York, NY, 1987, Vol. 68; pp 75-97. 12. Persson, I.; Lindman, B. In Flocculation in Biotechnology and Separation Systems; Attica, Y.A., Ed.; Elsevier: Amsterdam, Netherlands, 1987, pp 457466. 13. Cabrai, J.M.S.; Robinson, E.M.; Cooney, C.L. U.S. Patent 4 830 753, 1989. 14. Jefferson, R.A.; Burgess, S.M.; Hirsh, D. Proc. Natl. Acad. Sci. USA. 1986, 83, 8447-8451. 15. Novel, M.; Novel, G. J. Bacteriol. 1976, 127, 406-417. 16. Hanson, R.S.; Phillips, J.A. In Manual of Methods for General Bacteriology; Gerhardt, P.; Murray, R.G.E.; Costilow, R.N.; Nester, E.W.; Wood, W.A.; Krieg, N.R.; Phillips, G.B. Eds.; American Society for Microbiology: Washington, DC, 1981, pp 328-364. 17. Koegel, R.G.; Fomin, V.I.; Bruhn, H.D. Trans. A.S.A.E. 1972, 16, 712-716. 18. Leatham, G.F.; Stahmann, M.A. J. Gen. Microbiol. 1981, 125, 147-157. 19. Quiros, C.F. Crop Sci. 1980, 20, 262-264. 20. Straub, R.J.; Basken, K.E.; Koegel, R.G.; Bruhn, H.D. Trans. A.S.A.E. 1977, 20, 649-652. 21. Loomis, W.D. In Methods in Enzymology; Fleischer, S.; Packer, L. Eds.; Academic Press: New York, NY, 1974, Vol. XXXI, pp 528-544. 22. Folgert, J.; Forrester, I.T.; Bush, R.K. J. Allergy Clin. Immunol. 1990, 85, 153. RECEIVED August 16,

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Bioprocessing Aids in the Recovery of Proteins from Biomass

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