48 In Vil,'o PROTEIN DEPOSITION [3] [3] Isolating Inclusion Bodies from Bacteria By GEORGE GEORGIOU and PASCAL VALAX In~oducUon In 1975 Prouty et al. i first described the formation of dense, amorphous intracellular granules in Escherichia coli cells grown in the presence of the amino acid analog canavanine. These granules, comprised primarily of polypeptide chains, could be solubilized by sodium dodecyl sulfate (SDS) and were not surrounded by any sort of membrane. For several years this observation was considered an aberrant, and rather irrelevant, cellular response induced by growth under nonphysiological conditions. It was not until much later that it became apparent that protein aggregation in vivo is a widespread phenomenon, manifested in cells overexpressing heterologous proteins or native proteins beyond a certain level, and in cells exposed to thermal or other kinds of physiological stress. In addition, mutations re- suiting in amino acid substitutions, deletions, or insertions can interfere with the folding of a polypeptide to the native state, causing the formation of protein aggregates. 2 Intracellular protein aggregates form dense, electron-refracting particles that can be distinguished readily from other cell components by electron microscopy. For this reason, protein aggregates, at least those observed in microorganisms, are usually called inclusion bodies or, less often, refractile bodies. However, it should be noted that protein misfolding and self-associa- tion may occur even when inclusion bodies cannot be detected by electron microscopy or following careful cell fractionation (H. G. Gilbert, personal communication, 1999). Nonetheless, for practical purposes, it is safe to assume that the expression of proteins susceptible to aggregation at a level 2% or greater of the total cell protein will be accompanied by the appearance of readily identifiable inclusion bodies. Inclusion bodies have been used extensively as a source of relatively pure, albeit misfolded, polypeptide chains that can be renatured to give the biologically active soluble protein. The refolding of proteins from inclusion bodies has proven of great value for analytical and preparative purposes. As of 1998, there have been over 300 reports of mammalian, plant, and microbial proteins obtained and renatured from inclusion bodies formed in E. coli. Tens of kilograms of biologically active bovine somatotropin are 1 W. Prouty, M. J. Karnovsky, and A. L. Goldberg, J. Biol. Chem. 250, 1112 (1975). 2 A. L. Fink, Folding Design 3, R9 (1998). Copyright © 1999 by Academic Press All rights of reproduction in any form reserved. METHODS IN ENZYMOLOGY, VOL. 309 0076-6879/99 $30.00
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`KASHIV EXHIBIT 1020
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`[3] ISOLATING INCLUSION BODIES 49 produced every year from inclusion bodies and even larger amounts of human hemoglobin may eventually be produced by renaturation. Irrespec- tive of the scale, the recovery of purified active proteins from inclusion bodies involves the following steps3: (1) isolation of inclusion bodies, (2) solubilization, (3) removal of protein impurities, and (4) refolding. Often the sequence of steps 3 and 4 is interchanged so that the desired protein is purified after refolding. Obviously, the success of steps 2-4 de- pends on the quality of material from step 1. Also, preparing high-quality inclusion bodies is important for structural studies. 4'5 Inclusion bodies formed in cells overexpressing a certain polypeptide are generally expected to be comprised predominantly of that polypeptide. However, this is often not the case. We have found 6 that the composition of inclusion bodies is a complex function of the mode of expression and the growth conditions. For example, purified inclusion bodies from E. coli cells expressing/3-1actamase contain between 35 and 95% intact/3-1actamase polypeptides. The rest are composed of a variety of intracellular proteins, some lipids, and a small amount of nucleic acids. Homogeneous inclusion bodies [95% (w/w)/3-1actamase] were obtained by expressing the protein without its leader peptide, in which case aggregation occurred within the bacterial cytoplasm. This gave rise to large, highly regular inclusion bodies that could be separated readily from other particulate matter in cell lysates (Fig. 1). As might be expected, the efficiency of/3-1actamase refolding was inversely proportional to the level of contaminants present in the inclusion body preparation. 7 "High-quality" inclusion bodies consist primarily of the overexpressed recombinant protein with as little contaminating material as possible. The quality of inclusion bodies obtained from bacteria overexpressing a desired protein depends on two parameters: (1) the degree to which extraneous polypeptides, and possibly other macromolecules, are incorporated within the aggregate and (2) the ability to separate inclusion bodies from other cellular particles having a similar sedimentation coefficient and from mate- rial, mostly membrane vesicles, that becomes adsorbed onto the surface of the protein particles following cell lysis. It is usually difficult to ascertain whether extraneous proteins form an integral part of the aggregates or represent copurifying contaminants. There is increasing evidence, discussed in other articles of this volume, that protein aggregation involves the associ- 3 R. Ruboph and H. Lilie, FASEB J. 50, 49 (1996). 4 K. Oberg, B. A. Chrunyk, R. B. Wetzel, and A. L. Fink, Biochemistry 33, 2628 (1994). 5 T. M. Przybycien, J. P. Dunn, P. Valax, and G. Georgiou, Prot. Engin. 7, 131 (1994). 6 p. Valax and G. Georgiou, Biotech. Progr. 9, 539 (1993). 7 p. Valax, "In Vivo and in Vitro Folding and Aggregation of Escherichia coli/]-Lactamase." Ph.D Dissertation, Univ. of Texas at Austin, 1993.
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`50 In Vivo PROTEIN DEPOSITION [3] FIG. 1. (A) E. coli cells containing inclusion bodies and (B) inclusion bodies isolated by sucrose density centrifugation. Inclusion bodies were formed in the cytoplasm by expressing /3-1actamase (a normally secreted protein) carrying a deletion of the first 20 amino acids (-20, -1) of the leader sequence.
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`[3] ISOLATING INCLUSION BODIES 51 ation of subdomains in partially folded intermediates. 2 The degree of speci- ficity of the interactions that lead to protein aggregation varies from protein to protein. As a consequence, the extent of incorporation of cellular proteins and other extraneous macromolecules into inclusion bodies is protein de- pendent. Nonetheless, as was shown with/3-1actamase, 6,8 it is often possible to modify the expression conditions to reduce the amount of extraneous material incorporated within the inclusion bodies. Often the only step in the preparation of inclusion bodies is low-speed centrifugation of cell lysates. The effectiveness of this step depends to a large degree on the method of cell disruption. Low-speed centrifugation results in the sedimentation of membrane vesicles and cell wall fragments together with the inclusion bodies. In addition, other cellular components can become adsorbed nonspecifically onto the surface of inclusion bodies following cell lysis. The presence of contaminating substances causes a number of complications during subsequent solubilization and refolding steps: First of all, proteases may copurify with inclusion bodies during high- speed centrifugation. 9'1° Many proteases are active in the presence of high concentrations of denaturants used to solubilize the inclusion bodies and can rapidly cleave unfolded proteins under these conditions. For example, the presence of contaminating E. coli outer membrane protease OmpT was shown to cause significant reductions in the recovery of active porcine growth hormone I1 and creatine kinase 9 from inclusion bodies. Second, impurities found in inclusion bodies have to be removed eventually either prior to, or after, refolding. Third, protein as well as nonproteinaceous impurities interfere with refolding. In an interesting study, Maachupalli- Reddy and co-workers 12 examined the effects of typical inclusion body contaminants such as DNA, ribosomal RNA, lipids, and other proteins on the in vitro refolding of hen egg-white lysozyme. They found that the presence of other polypeptides prone to aggregation reduced the refolding yield significantly. The effect of RNA, DNA, and phospholipids at concen- trations up to 30% of that of hen lysozyme did not have a significant effect on refolding yields. However, Darby and Creighton 13 found that nonproteinaceous contaminants had a dramatic effect on the refolding of bovine pancreatic trypsin inhibitor (BPTI) mutants from inclusion bodies. G. A. Bowden, A. M. Paredes, and G. Georgiou, Bio/Technology 9, 725 (1991). 9 p. C. Babbit, B. L. West, D. D. Buechter, I. D. Kuntz, and G. L. Kenyon, Bio/Technology 8, 945 (1990). a0 J.-M. Betton, N. Sasson, M. Hofnung, and M. Laurent, J. Biol. Chem. 273, 8897 (1998). 11 N. K. Puri, M. Cardamone, E. Crivelli, and J. C. Traeger, Prot. Expr. Purif 4, 164 (1993). 12 j. Maachupalli-Reddy, B. D. Kelley, and E. De Bernardez-Clark, Biotech. Prog. 13, 144 (1997). 13 N. J. Darby and T. E. Creighton, Nature 344, 715 (1990).
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`52 In Vil.~o PROTEIN DEPOSITION [31 They suggested that even under the strongly denaturing conditions used to solubilize the inclusion bodies, BPTI is in a tight complex with a substance that affects its solubility and folding properties. This substance was found to be an acidic polymer, which unfortunately was not characterized further. Cardamone et al. 14 reported that the recovery yield of recombinant porcine growth hormone from inclusion bodies is lower than what can be achieved with the purified protein. They proposed that "morphopoietic factors" intrinsic to the inclusion bodies were responsible for preventing the protein from following the same folding pathway during renaturation, thus enhanc- ing aggregation. From these results it should be evident that the preparation of inclusion bodies suitable for protein recovery requires careful consideration of the expression conditions to minimize both the extent of nonspecific protein incorporation and the purification protocol. Because the formation of inclu- sion bodies is protein dependent, it is not possible to develop procedures that work for every case. Therefore, the remainder of this article is intended more as a set of recommendations that generally give good results in our experience. Protein Expression The bacterial strain, expression vector, and growth conditions have a pronounced effect on inclusion body formation. Expression from a T7 promoter 15 is accompanied by very high rates of protein synthesis, which enhance inclusion body formation. Genes placed downstream of a T7 pro- moter are transcribed by the T7 RNA polymerase. The gene for the latter is usually expressed from a relatively weak inducible promoter such as the lac promoter. Addition of the inducer IPTG turns on the synthesis of T7 RNA polymerase, which in turn transcribes the desired gene from the T7 promoter. For most applications, expression from a T7 promoter is the preferred way for inducing the formation of inclusion bodies. Ideally, the host strain should be a mutant defective in the gene responsible for the transport of the inducer into the cell. In such a host, induction with subsatu- rating concentrations of inducer can be employed to adjust the level of transcription per cell. 16 Another promoter that we have found to be favor- able for inclusion body formation is the PL of bacteriophage A. With the PL promoter, a very high rate of transcription is achieved by a temperature 14 M. Cardamone, N. K. Purl, and M. R. Brandon, Biochemistry 34, 5773 (1995). 15 W. F. Studier, A. H. Rosenberg, J. J. Dunn, and J. W. Dubendorff, Meth. Enzymol. 85, 61 (1990). 16 D. A. Siegele and J. C. Hu, Proc. Natl. Acad. Sci. U.S.A. 94, 8168 (1997).
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`[3] ISOLATING INCLUSION BODIES 53 upshift to 42 °, although cold temperature induction can also be employed. 17 At 42 ° the combination of a higher temperature, which generally favors aggregation over folding, and the high rate of protein synthesis favor inclu- sion body formation. However, exposure of the bacteria to a supraoptimal temperature is also accompanied by the increased synthesis of heat shock proteases, which may cause degradation of the cloned product. In E. coli, protein aggregation is enhanced by mutations in certain global regulatory pathways or in specific chaperone genes. The heat shock transcription factor 0 -32 is responsible for upregulating the expression of a number of E. coli chaperones under stressful conditions, including the DnaK-DnaJ-GrpE and GroEL-GroES systems. Mutations in the rpoH gene encoding 0-32 have been shown to induce the complete aggregation of a normally soluble recombinant protein. 18't9 However, massive aggregation of host proteins also occurs in rpoH mutants and, as a result, the insoluble fraction contains, in addition to the desired polypeptide, a number of con- taminating proteins. Mutations in the dnaK, dnaJ, grpE, groEL, or groES genes have also been found to favor the aggregation of recombinant proteins while having little effect on host protein solubility. 19 Hosts carrying muta- tions in dnaK, dnaJ, or grpE are particularly useful when it is desired to direct a protein into inclusion bodies. The DnaK-DnaJ-GrpE chaperone machinery binds newly synthesized polypeptides and mutations that inter- fere with its function are more likely to lead to rapid aggregation before proteolytic degradation can occur. Indeed, a number of unrelated overex- pressed proteins have been found to aggregate extensively in a grpE280 mutant, whereas the extensively studied groES30 or groEL140 alleles af- fected folding of only a limited number of proteins, often at the expense of reduced yields (J. Thomas, personal communication). 2°,21 Normally secreted heterologous proteins can be expressed either in the cytoplasm or with a prokaryotic leader peptide for targeting to the periplasmic space. Inclusion bodies can form in either cellular compartment. For secreted proteins containing two or more cysteine residues, the forma- tion of protein aggregates in the periplasm may be increased by the coex- pression of E. coli cysteine oxidoreductases (DsbC or DbA). 22 Periplasmic inclusion bodies are smaller and of irregular shape, most likely because 17 S. C. Macrides, Microbiol. Rev. 60, 512 (1996). 18 A. I. Gragerov, E. S. Martin, M. A. Krupenko, M. V. Kashlev, and V. G. Nikiforov, FEBS Lett. 291(2), 222 (1991). 19 A. Gragerov, E. Nudler, N. Komissarova, G. A. Gaitanaris, M. E. Gottesman, and V. Nikiforov, Proc. Natl. Acad. Sci. U.S.A. 89, 10341 (1992). 2o j. G. Thomas and F. Baneyx, Mol. Microbiol. 21, 1185 (1996). 21 j. G. Thomas and F. Baneyx, Prot. Express. Purif. 11, 289 (1997). 2z j. C. Joly, W. S. Leung, and J. Swartz, Proc. Natl. Acad. Sci. U.S.A. 95, 2773 (1998).
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`54 In Vivo PROTEIN DEPOSITION [3] topological constraints limit the growth of the aggregate. 8 Inclusion bodies in the cytoplasm are larger and often highly regular (Fig. 1). For this reason, they can be separated more readily from cell debris and may contain lower amounts of extraneous polypeptides, phospholipids, and nucleic acids. 6 In some cases, high levels of expression of secreted proteins that are prone to aggregation are accompanied by the accumulation of the precursor form (the preprotein) with the leader peptide uncleaved. 6'23 The presence of the leader peptide renders preproteins highly insoluble and they are found either in the membrane fraction or are sequestered within cytoplasmic aggregates. These aggregates cofractionate with periplasmic inclusion bod- ies formed by the mature protein in the periplasm. The formation of a mixed aggregate population, consisting of preprotein and mature protein inclusion bodies, poses further complications during refolding and should be avoided. The growth conditions can also be optimized to enhance inclusion body formation. As was mentioned earlier, growth at 42 ° results in decreased yields of soluble protein. Aeration, i.e., the concentration of dissolved oxygen, has been shown to affect protein aggregation. 24 The effect of dis- solved oxygen on protein aggregation is complex and, in our experience at least, growth in a low dissolved oxygen environment can either enhance or reduce the formation of inclusion bodies, depending on the promoter and plasmid vector employed (unpublished data). The addition of ethanol to the growth media at concentrations around 3% (v/v) has been found to increase inclusion body formation with some proteins, such as human SPARC, but had the reverse effect with others. 21 Finally the culture pH, carbon source, and growth in minimal versus rich media also affect in vivo solubility in a protein-dependent manner. Inclusion Body Isolation Obtaining a homogeneous preparation of inclusion bodies requires a three-step process involving (i) cell lysis, (2) fractionation of the cell lysates to resolve the inclusion bodies from the cell debris by taking advantage of differences in size and density, and (3) removal of adsorbed contaminants. Although incubation of intact E. coli with denaturants has been used for the in situ solubilization of aggregated proteins, such as IGF-I, 25 in general it is necessary to first lyse the bacteria. Laboratory methods for 23 N, Sriubolmas, W. Panbangred, S. Sriurairatana, and V. Meevootisom, Appl. Microbiol. Biotechnol. 47, 373 (1997). 24 S. D. Betts, T. M. Hachigian, and E. Pichersky, Plant. Mol. Biol. 26, 117 (1994). 25 R. H. Hart, P. M. Lester, D. H. Reifsnyder, and J. R. Ogez, Bio/Technology 12,1113 (1994).
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`[3] ISOLATING INCLUSION BODIES 55 rupturing bacteria include repeated freeze-thaw cycles, lysozyme-EDTA treatment, sonication, and high-pressure homogenization using a French pressure cell. The method of lysis determines the size of cell debris that is generated. Because inclusion bodies are separated from the debris on the basis of size and density, it is an important parameter in recovery. High- pressure homogenization is the recommended method for obtaining a high degree of cell disruption (90-98% after one pass) 26 and smaller size debris. Multiple passes have been shown to further reduce the debris size down to a median diameter of around 0.3 /zm. 26 For comparison, the size of inclusion bodies ranges between 0.3 and 1.2 /zm. We have found that pretreatment with lysozyme-EDTA further improves inclusion body isola- tion. 6 In gram-negative bacteria, the outer membrane is linked covalently to the cell wall and therefore hydrolysis of the peptidoglycan by lysozyme may be necessary to facilitate the formation of smaller size outer membrane vesicles. As a rule, three passes through either a French press at 20,000 psi or equivalent conditions for industrial scale high-pressure homgenization equipment are adequate for good separation. 27'2s Inclusion bodies are recovered from the cell lysate by centrifugation, usually at 15,000-30,000g in the laboratory (up to 20,000g for industrial scale). In addition to the aggregated protein, the pelleted fraction contains extraneous polypeptides and phospholipids. An appreciable amount of nucleic acids is also found. 29 The insoluble fraction obtained after centrifu- gation contains two prominent bands of molecular mass approximately 34 and 36 kDa (their migration may vary somewhat depending on the way the samples are treated prior to loading on the gel). These correspond to the major outer membrane proteins OmpA and OmpC/F, respectively, indicating that outer membrane fragments represent a major source of contaminating material. Outer membrane proteins are found in both cyto- plasmic and periplasmic inclusion body preparations. Ribosomal proteins have also been found in inclusion bodies. 3° Cosedimenting ribosomes and outer membrane vesicles can be sepa- rated from inclusion bodies by density gradient centrifugation. 8,31 Inclusion bodies have a density comparable to that of proteins (1.3-1.4 g/ml), whereas the density of outer membrane vesicles is 1.22 g/ml and that of ribosomes 26 H. H. Wong, B. K. O'Neill, and A. P. J. Middleberg, Biotech. Bioeng. 55, 556 (1997). 27 E. A. Burks, and B. L.Iverson, Biotech. Progr. 11, 112 (1995). 28 M. E. Gustafson, K. D. Junger, B. A. Foy, J. A. Baez, B. F. Bishop, S. H. Rangwala, M.L. Michener, R. M. Leimgruber, K. A. Houseman, R. A. Mueller, B. K. Matthews, P. O. Olins, R. W. Grabner, and A. Hershman, Prot. Expres. Purif 6, 512 (1995). 29 K. E. Langley, T. F. Berg, and T. W. Strickland, Eur. J. Biochem. 163, 313 (1987). 30 U. Rinas and J. Bailey, Appl. Microbiol. Biotechnol. 37, 609 (1992). 31 L. A. Classen, B. Ahn, H.-S. Koo, and L. Grossman, J. BioL Chem. 266, 11380 (1991).
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`56 In Viyo PROTEIN DEPOSITION [3] around 1.5 g/ml. In a sucrose step gradient, inclusion bodies become focused into a relatively tight and visible band that can be distinguished clearly from membrane vesicles that are lighter and form a separate band. Good results have been obtained with a variety of inclusion body proteins using a 40-53-67% (w/w) sucrose step gradient, but slightly different conditions have also been used successfully. 3° The inclusion body band is found close to or at the interface of the 53 and 67% (w/w) sucrose layers and is collected with a Pasteur pipette. The inclusion bodies are then resuspended in buffer and precipitated by low-speed centrifugation several times to remove the sucrose. Analysis of the purified inclusion bodies by SDS-PAGE should show a clear reduction in the amount of outer membrane protein. The amount of lipid in the preparation is reduced by up to 8-fold, although in many cases a substantial amount of lipid still remains. 6 The nucleic acid content is also reduced significantly, typically by 10- to 20-fold over the amount in the inclusion body pellet prior to purification. If the density of the inclusion bodies is low, the respective band is not well resolved from the cell membrane material. In that case, further purification can be obtained using a second sucrose density gradient centrif- ugation step. 6 Either an identical step gradient or a flotation gradient can be used at this stage. 32 For the latter, material is applied to the bottom of the tube and floats up a 60-40% (w/w) gradient until it reaches its buoyant density. For large-scale protein recovery, cross-flow filtration has been used as an alternative to centrifugation. Cross-flow filtration is a separation tech- nique in which the suspension is flowed parallel to the membrane in order to avoid the accumulation of material on the filter surface and to improve the filtration rate. Particles above a certain size cutoff are retained by a membrane and are collected continuously. With cross-flow filtration, inclu- sion body concentration and washing can be performed continuously. Pro- tein recovery yields comparable to those obtained by centrifugation have been reported. 33 After separation from cell debris, a series of wash/extraction steps is used to further remove contaminating material that had cosedimented or was adsorbed on the inclusion bodies. TM Membrane material associated with the inclusion bodies is typically removed using mild detergents. An appropriate detergent should allow efficient extraction of impurities without solubilizing the inclusion bodies or irreversibly binding to the aggregated 32 I. Poquet, M. G. Kormacher, and A. P. Pugsley, Mol. Microbiol. 9, 1061 (1993). 33 M. M. Meagher, R. T. Barlett, V. R. Rai, and F. R. Khan, Biotech. Bioeng. 43, 969 (1994). 34 F. A. O. Marston, in "DNA Cloning: A Practical Approach" (D. M. Glover, ed.), Vol. 3, p. 59. IRL Press, Oxford, 1987.
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`[3] ISOLATING INCLUSION BODIES 57 polypeptide chains. Price can also be a consideration as some detergents are very costly. Several detergents have been used in the literature, including Triton X-100, deoxycholate, Berol 185, and octylglucoside. 7'35 Nonspecifi- cally adsorbed material can also be removed using low concentrations of chaotropic agents such as urea or guanidine hydrochloride. Because these agents are generally used for the solubilization of aggregated proteins, their concentration must be carefully optimized in order to efficiently remove contaminating material without significantly solubilizing the inclusion bod- ies. Alternatively, inclusion bodies may be treated with a combination of detergents and low concentrations of chaotropic agents. For example, hu- man cathepsin B inclusion bodies were purified by extraction in 0.1% Triton X-100 once and 2 M urea twice, 36 whereas Belew et aL 35 used 0.1% Berol 185 and 0.5 M urea to remove impurities from recombinant human granulo- cyte-macrophage colony-stimulating factor. The remainder of this article describes in detail a general procedure we have found useful for inclusion body isolation. Cell L ysis The conditions for E. coli growth and protein synthesis have to be optimized first, depending on the expression system. At the appropriate time after induction, the cells are harvested by centrifugation at 8000g for 10 min at 4 ° and washed once in buffer. The cell pellet is resuspended in 10 mM Tris-HC1, pH 7.5, containing 0.75 M sucrose and 0.2 mg/ml lysozyme. It is recommended to resuspend the cell pellet from a 50-ml culture of 1.0 OD600 into 1 ml of buffer. After a 10-rain incubation at room temperature, a 3 mM EDTA solution is added at a 2:1 (v/v) ratio and transferred to ice for approximately 5 min. Subsequently, the cells are lysed by passing through a French press three times at 20,000 psi. Inclusion Body Isolation Following cell disruption, the lysate is centrifuged at 12,000g for 30 min at 4 ° and the pellet, which contains the inclusion bodies, is resuspended in 10 mM Tris-HC1 buffer, pH 8.0, containing 0.25 M sucrose, 1 mM EDTA, and 0.1% sodium azide (1.25 ml of buffer per 50 ml of 1.00D600). A tissue homogenizer may be used to resuspend the pellet thoroughly. The resuspended pellet is layered on the top of a sucrose step gradient [40, 53, and 67% (w/w)] in 1 mM Tris-HC1 buffer, pH 8.0, containing 0.1% sodium azide and 1 mM EDTA. The sucrose gradient is prepared by carefully 35 M. Belew, Y. Zhou, W. Wang, L.-E. Nystr6m, and J.-C. Janson, Chromatogr. A 679, 67 (1994). 36 R. Kuhelj, M. Dolinar, M. Pugercar, and V. Turk, Eur. J. Biochem. 229, 533 (1995).
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`58 In Vil;o PROTEIN DEPOSITION 141 layering the sucrose solutions, with the more dense one at the bottom of the tube. Centrifugation is performed at 108,000g for 90 min at 40. 8 The inclusion bodies become focused in a band at the interface between the 53 and 67% sucrose layers and are recovered and resuspended in either water or a suitable buffer, as required for further refolding steps. Several washes, followed by reprecipitation of the inclusion bodies by centrifugation at 12,000g for 30 min, are required to remove the sucrose. Alternatively, the sucrose can be removed by dialysis. At this point the aggregated protein should be examined by SDS-PAGE to evaluate the degree of purity. If the desired polypeptide constitutes less than 70% of the insoluble protein, then additional purification is recommended: The inclusion bodies are resus- pended in 50 mM KH2PO4, pH 7.0, containing 50 mM octylglucoside and incubated for 15 min at room temperature. Following centrifugation at 12,000g for 20 min, the pellet is resuspended in 0.25 M sucrose solution and applied to a second sucrose gradient, as described previously. [4] Isolation of Amyloid Deposits from Brain By ALEX E. ROHER and Yu-MIN Kuo Introduction The profuse deposition of insoluble amyloid-/3 (A/3) fibrils in the paren- chymal and vascular extracellular spaces of the cerebral cortex and lepto- meningeal vessels is one of the main histopathological lesions of Alzhei- mer's disease (AD). Fibrillar deposits of amyloid concentrate at the center of the senile plaques, usually surrounded by dystrophic neurites, or accumu- late around cerebral blood vessels, leading to the death of vascular myo- cytes. 1 Nonfibrillar amyloid also deposits in the cerebral cortex in the form of diffuse plaques, which are apparently devoid of neuritic pathology. 2 The 40 to 42 amino acid A/3 peptides result from the proteolytic degradation of the transmembrane/3-amyloid precursor protein? Soluble monomeric and oligomeric forms of these peptides are normally present in the human 1 H. M. Wisniewski, C. Bancher, M. Barcikowska, G. Y. Wen, and J. Currie, Acta Neuropathol. 78, 337 (1989). 2 H. Yamaguchi, Y. Nakazato, S. Hirai, M. Shoji, and Y. Harigaya, Am. J. Pathol. 135, 593 (1989). 3 D. J. Selkoe, Annu. Rev. Neurosci. 17, 489 (1994). Copyright © 1999 by Academic Press All rights of reproduction in any form reserved. METHODS IN ENZYMOLOGY. VOL. 309 0076-6879/99 $30.00
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