蛋白酶

APPLIED AND ENVIRONMENTAL MICROBIOLOGY, Nov. 2010, p. 7500–7508 Vol. 76, No. 22 0099-2240/10/$12.00 doi:10.1128/AEM.01558-10 Copyright © 2010, American Society for Microbiology. All Rights Reserved. Novel Fluorescence-Assisted Whole-Cell Assay for Engineering and Characterization of Proteases and Their Substrates� George Kostallas and Patrik Samuelson* Division of Molecular Biotechnology, School of Biotechnology, AlbaNova University Center, Royal Institute of Technology (KTH), SE-106 91 Stockholm, Sweden Received 30 June 2010/Accepted 11 September 2010 We have developed a sensitive and highly efficient whole-cell methodology for quantitative analysis and screening of protease activity in vivo. The method is based on the ability of a genetically encoded protease to rescue a coexpressed short-lived fluorescent substrate reporter from cytoplasmic degradation and thereby confer increased whole-cell fluorescence in proportion to the protease’s apparent activity in the Escherichia coli cytoplasm. We demonstrated that this system can reveal differences in the efficiency with which tobacco etch virus (TEV) protease processes different substrate peptides. In addition, when analyzing E. coli cells expressing TEV protease variants that differed in terms of their in vivo solubility, cells containing the most-soluble protease variant exhibited the highest fluorescence intensity. Furthermore, flow cytometry screening allowed for enrichment and subsequent identification of an optimal substrate peptide and protease variant from a large excess of cells expressing suboptimal variants (1:100,000). Two rounds of cell sorting resulted in a 69,000-fold enrichment and a 22,000-fold enrichment of the superior substrate peptide and protease variant, respectively. Our approach presents a new promising path forward for high-throughput substrate profiling of proteases, engineering of novel protease variants with desired properties (e.g., altered substrate specificity and improved solubility and activity), and identification of protease inhibitors. Proteases constitute a group of enzymes that irreversibly catalyze the cleavage of peptide bonds and represent approx- imately 2% of all protein-encoding genes in living organisms (39). Besides acting as virulence factors for many pathogens (16), proteases are crucial for the regulation of numerous biological processes that influence the life and death of a cell (4). These enzymes also underlie several pathological con- ditions, such as cancer (13) and neurodegenerative (20) and cardiovascular (8) diseases. A key issue for increasing our knowledge about such complex biological processes, and thereby hopefully also providing possibilities for new therapeu- tic strategies, is to deduce the proteases’ substrate repertoires. Consequently, a lot of efforts around the world are dedicated to the characterization of proteases and their substrates (2, 31). In addition to their biological importance, proteases have attracted much interest in several biotechnological and in- dustrial applications, such as removal of “fusion tags” from recombinant target proteins (38), as supplements in dish- washing and laundry detergents, or for bating of hides and skin in the leather industry (41, 44). Sometimes, however, their use is hindered due to limitations inherent to a specific pro- tease: for example, low solubility, poor enzyme stability and specificity, or limited activity. It would therefore be of great aid to have powerful and straightforward methods available that facilitate the engineering of novel protease variants not suffer- ing from such limitations. Traditionally, protease substrate specificity has been studied by comparison and alignment of naturally occurring substrate peptide sequences (7) or through biochemical analysis of cleav- age products with synthetic peptides (47). More recent and powerful methods instead rely on the use of combinatorial substrate libraries, which can either be chemically or biologi- cally generated (6, 15). Although all of these methods have proven useful in determining protease function, many suffer from being laborious and of limited throughput capacity, hav- ing an insufficient dynamic range, and resulting in limited in- formation on the substrate profile. Moreover, only a small fraction of all proteases have been studied to date, and there is a need for novel approaches that allow for determination of protease specificity in a rapid, accurate, and quantitative manner. Concerning the engineering of enzymes toward novel de- sired properties, like altered substrate specificity and improved activity, solubility, and stability; researchers have relied on the use of rational design and/or directed-evolution methods in combination with appropriate screening and selection proce- dures (1, 10, 11, 22). For instance, various mutagenesis proce- dures and subsequent screening via assays that report on the successful folding of a protein of interest (9, 32, 45) have been used to engineer protein variants exhibiting improved solu- bility (35, 37, 46). Despite the obvious success of using such folding reporters in solubility/folding engineering projects, there is a risk that the engineered protein may lose its inherent activity since these screening procedures in general do not select for retained activity but only improved solubility/folding. Therefore, as in the case of a protease, it would be advanta- geous to establish a screening or selection system that has the ability to simultaneously address traits such as improved fold- ing/solubility without loss of proteolytic activity. However, di- rected evolution of desired catalytic properties has proven * Corresponding author. Mailing address: Division of Molecular Biotechnology, School of Biotechnology, AlbaNova University Center, Royal Institute of Technology (KTH), SE-106 91 Stockholm, Sweden. Phone: 46-8-5537 8335. Fax: 46-8-5537 8481. E-mail: patrik@biotech .kth.se. � Published ahead of print on 17 September 2010. 7500 quite a challenge. A popular strategy has been to use phage display technologies, often in combination with transition state analogues (18) or mechanism-based suicide inhibitors, for se- lection (30). Although successful, the enrichment conferred by these methods is generally based on binding rather than catal- ysis. Georgiou and coworkers circumvented this potential problem by developing an interesting system that actually enables function-based isolation of novel protease variants from large libraries (34, 42, 43). However, their methodol- ogy is dependent on the use of cell surface-displayed pro- teases, which is not applicable to all proteases and therefore may limit its usefulness. Herein, we present a novel, function-based, and highly effi- cient fluorescence-assisted whole-cell assay for characteriza- tion and engineering of proteases and their cognate substrate peptides. The method takes advantage of genetically encoded short-lived fluorescent substrates that upon coexpression of a substrate-specific protease result in a fluorescence signal, which can easily be monitored on a flow cytometer. Cells having a desired fluorescence profile can then be collected through sort- ing and sequenced to identify the protease-sensitive substrate peptide or protease capable of processing a particular peptide. Using this approach, we show that it is possible to analyze the efficiency with which the highly sequence-specific tobacco etch virus protease (TEVp) processes different substrate peptides and in model experiments also identify and enrich cells ex- pressing the most favorable substrate peptide or protease from a large background of cells harboring less-efficient variants. MATERIALS AND METHODS Bacterial strains and reagents. E. coli strain RR1�M15 [F� lacIq �(lacZ)M15 supE44 lacY1 lacZ ara-14 galK2 xyl-5 mtl-1 leuB6 proA2 �(mcrC-mrr) recA� rpsL20 thi-1 ��] (40) was used as the host during construction of plasmids. E. coli strain DH5� [F� �80lacZ�M15 �(lacZYA-argF)U169 recA1 endA1 hsdR17 (rk� mk�) phoA supE44 �� thi-1 gyrA96 relA1] (Gibco) was used for flow cytometry analysis and cell sorting experiments. Culture media, chemicals, and DNA-modifying enzymes were purchased from Merck (Darmstadt, Germany), Sigma-Aldrich (St. Louis, MO), and New England Biolabs (Ipswich, MA), re- spectively. Oligonucleotides. The following oligonucleotides (Eurofins MWG Operon) were used (5�33�): GEKO14 (GCAAACGACGAAAACTACAACTACGCTT TAGCAGCTTAAGCATGCAAG), GEKO15 (CTTGCATGCTTAAGCTGCT AAAGCGTAGTTGTAGTTTTCGTCGTTTGC), GEKO20 (CTCATCGATG GGCGCAACATGATAATTATTCGC), GEKO21 (GCGAATAATTATCATC AAGCGCCCATCGATGAG), PEAKfor (GGGGTACCCATCATCATCATC ATCATCATGGAG), PEAKrev (GATGGGTACCCATAATCTATGGTCCTT GTTGGT), SAPA46 (CTCTCGAGCTCGAATTCTCTAGATTAAAGAGGA GAAAGGTACCCATGAGTAAAGGAGAAGAACTTTTC), SAPA47 (CTCT CAAGCTTGCATGCTTAAGCTGCTAAAGCGTAGTTTTCGTCGTTTGCT GCGTCGACTTTGTATGTTCATCCATGCCATG), SAPA60 (TCGATGAA GCCCTGAAAGACG), SAPA61 (GGCGATTAAGTTGGGTAACGC), SAPA62 (TCGACGAAAACCTGTACTTCCAGGGTG), SAPA63 (TCGACACCCTG GAAGTACAGGTTTTCG), SAPA66 (TCGACGAAAACCTGTACTTCCAG CCGG), SAPA67 (TCGACCGGCTGGAAGTACAGGTTTTCG), SAPA68 (T CGACGAAAACCTGTACTTCCAGGGTTAAG), SAPA69 (TCGACTTAAC CCTGGAAGTACAGGTTTTCG), SAPA72 (GTTGGTATACACTCAGCA TCG), and SAPA73 (CGATGCTGAGTGTATACCAAC). Construction of plasmids. Heterologous expression of the catalytic domain of the tobacco etch virus protease (TEVp) in E. coli results in low yields of active protease due to the presence of several rare arginine codons in the TEVp gene (24) and self-inactivation caused by autoproteolysis between residues 218 and 219 (36). However, these negative effects can be relieved by exchanging the uncommon arginine codons for synonymous ones that are used more frequently in E. coli (24) and by creating a TEVp mutant (S219V) that is as active as but far more stable than the wild-type protease (26). Therefore, we combined relevant regions from the TEVp-encoding vectors pRK693 (24) and pRK793 (26) through gene splicing by overlap extension (23). First, two separate gene fragments were PCR amplified from pRK693 and pRK793, using primer pairs SAPA60/SAPA73 and SAPA72/SAPA61, respectively. Then, the two products were purified, mixed, and used as the template in an additional PCR, now with primers SAPA60 and SAPA61, to generate the PCR-spliced full-length product. The final amplicon was digested with HindIII and BamHI and ligated into the HindIII/BamHI-digested backbone of pRK693, resulting in pMal-TEV1. Based on this plasmid, we then created pMal-TEV2 by transferring the HindIII/MluI fragment (which encodes maltose-binding protein (MBP) fused to the TEVp substrate peptide, ENLYFQG) from pRK793 into the HindIII/MluI-digested backbone of pMal-TEV1. Thus, when TEVp is expressed from pMal-TEV2, the protease will only be transiently fused to the solubility-enhancing MBP moiety as this domain is removed from the fusion protein (MBP-ENLYFQG-TEVp) in- tracellularly, through TEVp-mediated substrate processing. We also constructed plasmid pTEV, which encodes TEVp without any solubility-enhancing MBP fusion. Thus, when the protease is expressed from this plasmid, it should exhibit very poor in vivo solubility (27). pTEV was constructed from pMal-TEV2, which served as the template in a PCR with primers PEAKfor and PEAKrev. The PCR product was treated with DpnI, purified on a Qiaquick PCR cleanup column (Qiagen), digested with KpnI, and then purified again. Finally, the digested amplicon was circularized by ligation, which resulted in pTEV. An expression vector for the catalytically inactive TEV protease variant (D81N) (26) was also created. pMal-TEV2 was used as the template in a QuikChange site-directed mutagenesis reaction (Stratagene) with primers GEKO20 and GEKO21 to in- troduce the D81N mutation, yielding pMal-InTEV. pGFP-ssrA, which encodes ssrA-tagged green fluorescent protein (GFP-ssrA; where ssrA is AANDENYA LAA) that is destroyed by the cytoplasmic degradation complex, ClpXP (14, 17), was used as a parental vector for subsequent construction of various TEVp substrate reporter plasmids. For construction of pGFP-ssrA, GFPmut3 (12) was PCR amplified using primers SAPA46 and SAPA47. The amplicon was then digested with SacI and HindIII and ligated into the SacI/HindIII-digested back- bone of pBAD33 (19), yielding pGFP-ssrA. A very similar plasmid, pGFP- ssrANY, that instead encodes a modified ssrA tag containing an additional pair of asparagine and tyrosine residues (in boldface) (AANDENYNYALAA), which improves ClpXP-mediated degradation efficiency (21), was also constructed. This was done by a QuikChange site-directed mutagenesis reaction (Stratagene) with primers GEKO14 and GEKO15 on the template pGFP-ssrA. Two TEVp substrate linkers, encoding either glycine (G) or proline (P) in the P1� position of the wild-type substrate peptide (resulting in ENLYFQG/P), were created from primer pairs SAPA62/SAPA63 and SAPA66/SAPA67, respectively. In addition, a third linker, encoding the wild-type substrate peptide directly followed by a stop codon, was made of primers SAPA68 and SAPA69. The linkers were inserted between the GFP and the ssrA coding regions of SalI-digested pGFP- ssrANY (or pGFP-ssrA) to create pGFP-subG-ssrANY (or pGFP-subG-ssrA), pGFP-subP-ssrANY, and pGFP-subG, respectively (subG is ENLYFQG and subP is ENLYFQP). All plasmid constructs were verified by standard DNA sequencing. Vectors encoding the substrate reporters all contained a p15A origin of replication, a chloramphenicol acetyltransferase gene, and a PBAD promoter controlling the reporter expression. The plasmids constructed for TEVp expres- sion all harbored the Ptac promoter (regulating protease expression), a ColE1 origin of replication, and an ampicillin resistance marker (bla gene, coding for �-lactamase). Flow cytometry analysis and cell sorting. A 0.1 mM concentration of isopro- pyl-�-D-1-thiogalactopyranoside (IPTG) and 0.2% arabinose (final concentra- tions) were used for induction of protein expression in all cases, unless otherwise stated. For clone analysis and cell sorting, overnight cultures of DH5� cells harboring a TEVp expression vector (pMal-TEV2, pMal-InTEV, or pTEV) and a relevant TEVp substrate reporter plasmid (either pGFP-subG-ssrA, pGFP- ssrA, pGFP-subG, pGFP-subG-ssrANY, pGFP-subP-ssrANY, or pGFP-ssrANY) were subcultured by dilution (1:75 for clone analysis and 1:150 for the sorting experiments) into fresh LB broth, containing 100 g/ml ampicillin and 20 g/ml chloramphenicol, and incubated at 37°C in a rotary shaker set at 150 rpm. When the cultures reached a cell density (optical density at 600 nm [OD600]) of 0.5, IPTG was added to initiate TEVp expression and the cultures were transferred to a rotary shaker set at 30°C, 150 rpm. Thirty minutes later, L(�)-arabinose was added to induce the reporter expression. After 2 h, 1 ml of each culture was placed on ice and 5 to 10 l from each sample was diluted with 1 ml ice-cold 1� PBS (11.68 g NaCl, 9.44 g Na2HPO4, 5.28 g NaH2PO4 � 2H2O, 1,000 ml MilliQ, pH 7.2) and kept on ice until analyzed on a FACSVantage SE flow cytometer (Becton Dickinson). The throughput rate for the flow cytometry analysis was 300 events/s, with an excitation wavelength of 488 nm (argon ion laser) and emission detection between 510 and 530 nm, and 10,000 events were recorded for each sample. For the experiments that aimed at finding induction conditions resulting VOL. 76, 2010 IN VIVO PROTEASE PROFILING SYSTEM 7501 in improved resolution between positive and negative cells with respect to the whole-cell fluorescence intensity, several different IPTG and arabinose concen- trations were tested. In these instances, the final concentrations of IPTG and arabinose ranged from 0.1 mM to 0.00016 mM and 0.2% to 0.00032%, respec- tively, in 5-fold dilution steps. We also conducted several cell-sorting experi- ments. In one series, we tried to enrich cells expressing GFP-subG-ssrANY (DH5�/pMal-TEV2/pGFP-subG-ssrANY) from a large excess of cells expressing GFP-subP-ssrANY (DH5�/pMal-TEV2/pGFP-subP-ssrANY). In another set of experiments, the aim instead was to enrich cells expressing TEVp in a soluble format (DH5�/pMal-TEV2/pGFP-subG-ssrANY) from a large background of cells expressing “insoluble” TEVp (DH5�/pTEV/pGFP-subG-ssrANY). The samples were prepared for fluorescence-activated cell sorting in essentially the same way as for flow cytometry analysis of individual clones. However, right before the first round of sorting, the different cell types were mixed to achieve a 100,000-fold excess of “background” cells. The populations were then analyzed on the flow cytometer with a throughput rate of 300 to 500 events/s. The cells were sorted, according to desired fluorescence intensity criteria, directly into LB broth. Collected cells were then either plated on solid LB agar containing ampicillin and chloramphenicol or regrown overnight for additional rounds of analysis and cell sorting. Western blot analysis. Overnight cultures of DH5� cells containing pMal- TEV2 and a TEVp substrate reporter plasmid (either pGFP-subG, pGFP-subG- ssrANY, or pGFP-ssrANY) were diluted 1:75 into fresh LB broth supplemented with 100 g/ml ampicillin and 20 g/ml chloramphenicol. The cultures grew at 37°C in a rotary shaker set at 150 rpm until they reached a cell density (OD600) of 0.5. At this point, TEVp expression was initiated by adding IPTG to a final concentration of 0.1 mM, and then the cultures were incubated at 30°C, 150 rpm. Reporter expression commenced 30 min later by addition of L-arabinose to a final concentration of 0.2%. Two hours later, a volume corresponding to 1.75 OD600 equivalents was withdrawn from each culture, and the cells were har- vested by centrifugation at 10,000� g. The cell pellets were then treated with 400 l BugBuster (Novagen) in order to release the protein content. After addition of 20 l 3� SDS denaturing buffer (150 mM Tris-Base, 300 mM dithiothreitol [DTT], 6% SDS, 0.3% bromophenol blue, 30% glycerol) to 40 l of each whole-cell lysate, the samples were heat denatured (96°C, 8 min) and run on an SDS-PAGE gel (Novex 4 to 12% Tris-glycine gradient gel; Invitrogen). The separated proteins were then transferred to a 0.45- m-pore-size polyvinylidene difluoride (PVDF) membrane (Invitrogen) and treated overnight with blocking buffer (5% skim milk, 0.5% Tween 20) at 8°C. The blocked membrane was incubated with a rabbit anti-GFP polyclonal antibody (1:2,000 dilution; Dianova) at room temperature for 1 h. After extensive washing with TBST (150 mM NaCl, 10 mM Tris base, 0.05% Tween 20) the membrane was incubated with horse- radish peroxidase (HRP)-conjugated goat anti-rabbit IgG (1:1,000 dilution; Pierce) for 1 h at room temperature. Finally, the protein bands were visualized with a chromogenic HRP substrate (Moss). RESULTS Development of a fluorescence-assisted whole-cell assay for protease activity. We sought to develop a label-free and sen- sitive high-throughput assay for in vivo monitoring of protease activity in E. coli by creating a genetically encoded protease- specific reporter system suitable for flow cytometry analysis and cell sorting. The reporter system was based on previous findings showing that a C-terminal fusion of the ssrA peptide (AANDENYALAA) to green fluorescent protein (GFP) ren- ders the whole fusion protein susceptible to intracellular deg- radation by the cytoplasmic protease ClpXP, which effectively eliminates the GFP-mediated whole-cell fluorescence (14, 28). By including a protease substrate peptide between GFP and the ssrA moiety in the reporter construct, it should be possible to remove the ssrA degradation tag and thereby rescue GFP from destruction, given that the fusion protein is coexpressed with a substrate-specific protease. Consequently, there should be an increase in the whole-cell fluorescence intensity, reflect- ing the protease’s apparent catalytic efficiency, whi