Patent Application
20240018501
The instant application contains a Sequence Listing that has been submitted in XML format via Patent Centrer and is hereby incorporated by reference in its entirety. Said XML copy, created on Sep. 25, 2023, is named UIU01-015-PCT.ST25.txt, and is 156,356 bytes in size.
This invention pertains to post-translational processing enzymes and methods for preparing a polypeptide product from a polypeptide precursor containing a lanthipeptide protease cleavage site.
The use of proteases has led to advances in analytical chemistry, proteomics, medicine, and the food, detergent and leather industries. Sequence-specific proteases are of particular interest as precise cleavage of proteins or peptides has great utility in many settings. Although much effort has been spent on engineering of proteases with desired sequence specificity using both rational design and high throughput screening, few such efforts have reached the stage of commercial applications. Major challenges are the sacrifice of efficiency and stability when engineering new substrate specificity, or the loss of sequence specificity when focusing on improving protein robustness. Thus, nature is still the major source of proteases with novel recognition sequences. The biosynthetic machinery responsible for the production of natural products known as ribosomally synthesized and post-translationally modified peptides (RiPPs) is a promising area for discovering new sequence-specific proteases, as dedicated proteolytic enzymes are employed to remove leader peptides with highly diverse P and P′ positions.
Lanthipeptides are (methyl)lanthionine-containing peptides that belong to the growing class of RiPPs. Similar to most RiPPs, lanthipeptides are synthesized as a precursor peptide (LanA) composed of an N-terminal leader peptide and a C-terminal core region harboring the different post-translational modification sites. The (methyl)lanthionine residues in lanthipeptides are installed in a two-step biosynthetic process. First, a lanthipeptide dehydratase catalyzes the elimination of water from Ser and Thr residues in the core region to yield dehydroalanine (Dha) and dehydrobutyrine (Dhb), respectively. A lanthipeptide cyclase then catalyzes the Michael type addition of cysteinyl thiols onto the dehydroamino acids. Following the modifications of the C-terminal core peptide, the modified precursor peptide is usually processed by a lanthipeptide peptidase that removes the N-terminal leader peptide (
Lanthipeptides are classified into four classes (class I-IV) on the basis of differences in the biosynthetic machinery responsible for installing the (methyl)lanthionines (
Compared to the well-characterized lanthionine synthetases, the proteases responsible for leader peptide removal to generate mature lanthipeptides are much less studied. Three different types of proteases have been reported, including the subtilisin-like LanP proteases found in both class I and class II lanthipeptide biosynthesis (e.g. NisP for nisin, EpiP for epidermin, ElxP for epilancin 15X, CytA for cytolysin, LicP for lichenicidin, and CerP for cerecidins), the cysteine protease domain in bi-functional LanT transporter proteins encountered in class II lanthipeptide biosynthesis (e.g. LctT for lacticin 481 and NukT for nukacin), and a prolyloligopeptidase-type protease identified for the biosynthesis of the class III lanthipeptide flavipeptin (
Although most class II lanthipeptides employ LanT proteins for leader peptide removal, a few use LanP proteases. Most of these LanPs appear to remove short N-terminal oligopeptides after LanT proteins remove the majority of leader peptides at a double Gly-type cleavage site. For example, CylA is an extracellular serine protease required for biosynthesis of the enterococcal cytolysin. After installation of the thioether rings in the precursor peptides CylLL and CylLS, CylB (a LanT protein) removes the majority of their leader peptides to generate CylLL′ and CylLS′ (
CylA contains an N-terminal secretion signal peptide and was reported to lack the first 95 amino acids when purified from the producing strain. Similar observations were also reported for two class I LanPs, NisP and EpiP, which lacked the first 195 and 99 amino acids in their mature forms, respectively. The removal of such a pro-sequence may activate the proteases for their LanA substrates. However, no activity comparison has been performed between the mature, processed form of LanP and its full-length version to confirm such activation. For CylA, the mature form was reported to exhibit the desired activity against CylLL′ and CylLS′ when purified from the producing strain supernatant, but no studies have been performed to define its substrate specificity.
A homology model of NisP, the peptidase involved in nisin biosynthesis, suggested that the substrate specificity of NisP relies on electrostatic and hydrophobic interactions between the S1/S4 NisP pockets and the residues in the −1 (Arg) and −4 positions (Ala) of nisin's precursor peptide NisA (
In general, the understanding of the substrate specificity of LanP enzymes is still limited in part because of the lack of detailed in vitro mechanistic studies as a result of the intrinsic low expression and poor solubility associated with these enzymes. Having a greater insight to these activities will improve the efficiency of using these lanthipeptide enzymes and their associated substrates in biotechnological, medical, and diagnostic applications.
In a first aspect, an isolated nucleic acid that includes an open reading frame encoding a lanthipeptide protease polypeptide for scarless tag removal from a polypeptide is disclosed.
In a second aspect, an expression cassette that includes an open reading frame for a polypeptide, wherein the open reading frame encodes a substrate recognition sequence for a lanthipeptide protease polypeptide, is disclosed.
In a third aspect, a method of scarless tag removal from a polypeptide is disclosed. The method includes two steps. The first step includes providing the polypeptide, wherein the polypeptide includes the structure: T-R-P, wherein T comprises a tag motif, R comprises a lanthipeptide protease substrate recognition sequence and P comprises an open reading frame encoding a polypeptide without the tag motif and lanthipeptide protease substrate recognition sequence. The second step includes subjecting the polypeptide to a lanthipeptide protease having specificity for catalyzing proteolytic cleavage at the lanthipeptide protease substrate recognition sequence, thereby providing the polypeptide without a tag scar.
In a fourth aspect, a kit for expressing a polypeptide without a tag scar is disclosed. The kit includes an expression vector that includes an expression cassette, wherein the expression cassette encoding a polypeptide including the structure: T-R-P, wherein T includes a tag motif, R includes a lanthipeptide protease substrate recognition sequence and P includes an open reading frame encoding a polypeptide without the tag motif and lanthipeptide protease substrate recognition sequence. The kit also includes a lanthipeptide protease having specificity for catalyzing proteolytic cleavage at the lanthipeptide protease substrate recognition sequence, thereby providing the polypeptide without the tag scar.
In a fifth aspect, isolated polypeptide including the structure T-R-P is disclosed. The T includes a tag motif, the R includes a lanthipeptide protease substrate recognition sequence and the P includes an open reading frame encoding a polypeptide without the tag motif and lanthipeptide protease substrate recognition sequence.
FIG. A depicts biosynthesis of epilancin 15X, involving dehydration of Ser and Thr residues by ElxB to yield dehydroalanine (Dha, green), and dehydrobutyrine (Dhb, purple), formation of lanthionine (red) or methyllanthionine (blue) rings catalyzed by ExC, removal of the leader peptide (bold) by the peptidase ElxP.
Reagents, expression constructs and methods are disclosed herein for preparing a scarless tag polypeptide product from a tagged polypeptide precursor containing a lanthipeptide protease cleavage site. The reagents are directed to the use of novel lanthipeptide proteases for processing polypeptide precursors that include highly specific lanthipeptide protease substrate recognition sequence. Methods are provided that enable scarless tag removal from a cognate lanthipeptide, a non-cognate lanthipeptide or a heterologous polypeptide that includes extraneous amino acid sequences, such as leader peptides and tags.
To aid in understanding the invention, several terms are defined below.
Unless defined otherwise, all technical and scientific terms used herein have the same meaning as commonly understood by one of skill in the art. Although any methods and materials similar to or equivalent to those described herein can be used in the practice or testing of the claims, the exemplary methods and materials are described herein.
Moreover, reference to an element by the indefinite article “a” or “an” does not exclude the possibility that more than one element is present, unless the context clearly requires that there be one and only one element. The indefinite article “a” or “an” thus usually means “at least one.”
The term “about” means within a statistically meaningful range of a value or values such as a stated concentration, length, molecular weight, pH, time frame, temperature, pressure or volume. Such a value or range can be within an order of magnitude, typically within 20%, more typically within 10%, and even more typically within 5% of a given value or range. The allowable variation encompassed by “about” will depend upon the particular system under study.
The terms “comprising,” “having,” “including,” and “containing” are to be construed as open-ended terms (i.e., meaning “including, but not limited to”) unless otherwise noted.
Recitation of ranges of values herein are merely intended to serve as a shorthand method of referring individually to each separate value falling within the range, and includes the endpoint boundaries defining the range, unless otherwise indicated herein, and each separate value is incorporated into the specification as if it were individually recited herein.
The terms “nucleic acid” and “oligonucleotide,” as used herein, refer to polydeoxyribonucleotides (containing 2-deoxy-D-ribose), polyribonucleotides (containing D-ribose), and to any other type of polynucleotide that is an N glycoside of a purine or pyrimidine base. There is no intended distinction in length between the terms “nucleic acid”, “oligonucleotide” and “polynucleotide”, and these terms will be used interchangeably. These terms refer only to the primary structure of the molecule. Thus, these terms include double- and single-stranded DNA, as well as double- and single-stranded RNA. For use in the present invention, an oligonucleotide also can comprise nucleotide analogs in which the base, sugar or phosphate backbone is modified as well as non-purine or non-pyrimidine nucleotide analogs.
Oligonucleotides can be prepared by any suitable method, including direct chemical synthesis by a method such as the phosphotriester method of Narang et al., 1979, Meth. Enzymol. 68:90-99; the phosphodiester method of Brown et al., 1979, Meth. Enzymol. 68:109-151; the diethylphosphoramidite method of Beaucage et al., 1981, Tetrahedron Letters 22:1859-1862; and the solid support method of U.S. Pat. No. 4,458,066, each incorporated herein by reference. A review of synthesis methods of conjugates of oligonucleotides and modified nucleotides is provided in Goodchild, 1990, Bioconjugate Chemistry 1(3): 165-187, incorporated herein by reference.
The term “primer,” as used herein, refers to an oligonucleotide capable of acting as a point of initiation of DNA synthesis under suitable conditions. Such conditions include those in which synthesis of a primer extension product complementary to a nucleic acid strand is induced in the presence of four different nucleoside triphosphates and an agent for extension (for example, a DNA polymerase or reverse transcriptase) in an appropriate buffer and at a suitable temperature.
A primer is preferably a single-stranded DNA. The appropriate length of a primer depends on the intended use of the primer but typically ranges from about 6 to about 225 nucleotides, including intermediate ranges, such as from 15 to 35 nucleotides, from 18 to 75 nucleotides and from 25 to 150 nucleotides. Short primer molecules generally require cooler temperatures to form sufficiently stable hybrid complexes with the template. A primer need not reflect the exact sequence of the template nucleic acid, but must be sufficiently complementary to hybridize with the template. The design of suitable primers for the amplification of a given target sequence is well known in the art and described in the literature cited herein.
Primers can incorporate additional features which allow for the detection or immobilization of the primer but do not alter the basic property of the primer, that of acting as a point of initiation of DNA synthesis. For example, primers may contain an additional nucleic acid sequence at the 5′ end which does not hybridize to the target nucleic acid, but which facilitates cloning or detection of the amplified product, or which enables transcription of RNA (for example, by inclusion of a promoter), termination of RNA transcription (for example, a ribozyme), or translation of protein. The region of the primer that is sufficiently complementary to the template to hybridize is referred to herein as the hybridizing region.
The term “promoter” refers to a cis-acting DNA sequence that directs RNA polymerase and other trans-acting transcription factors to initiate RNA transcription from the DNA template that includes the cis-acting DNA sequence.
The terms “target, “target sequence”, “target region”, and “target nucleic acid,” as used herein, are synonymous and refer to a region or sequence of a nucleic acid which is to be amplified, sequenced or detected.
The term “hybridization,” as used herein, refers to the formation of a duplex structure by two single-stranded nucleic acids due to complementary base pairing. Hybridization can occur between fully complementary nucleic acid strands or between “substantially complementary” nucleic acid strands that contain minor regions of mismatch. Conditions under which hybridization of fully complementary nucleic acid strands is strongly preferred are referred to as “stringent hybridization conditions” or “sequence-specific hybridization conditions”. Stable duplexes of substantially complementary sequences can be achieved under less stringent hybridization conditions; the degree of mismatch tolerated can be controlled by suitable adjustment of the hybridization conditions. Those having ordinary skill in the art of nucleic acid technology can determine duplex stability empirically considering a number of variables including, for example, the length and base pair composition of the oligonucleotides, ionic strength, and incidence of mismatched base pairs, following the guidance provided by the art (see, e.g., Sambrook et al., 1989, Molecular Cloning-A Laboratory Manual, Cold Spring Harbor Laboratory, Cold Spring Harbor, New York; Wetmur, 1991, Critical Review in Biochem. and Mol. Biol. 26(3/4):227-259; and Owczarzy et al., 2008, Biochemistry, 47: 5336-5353, which are incorporated herein by reference).
The term “amplification reaction” refers to any chemical reaction, including an enzymatic reaction, which results in increased copies of a template nucleic acid sequence or results in transcription of a template nucleic acid. Amplification reactions include reverse transcription, the polymerase chain reaction (PCR), including Real Time PCR (see U.S. Pat. Nos. 4,683,195 and 4,683,202; PCR Protocols: A Guide to Methods and Applications (Innis et al., eds, 1990)), and the ligase chain reaction (LCR) (see Barany et al., U.S. Pat. No. 5,494,810). Exemplary “amplification reactions conditions” or “amplification conditions” typically comprise either two or three step cycles. Two-step cycles have a high temperature denaturation step followed by a hybridization/elongation (or ligation) step. Three step cycles comprise a denaturation step followed by a hybridization step followed by a separate elongation step.
The term “natural polymer” refers to any polymer comprising natural monomers found in biology. For example, polypeptides are natural polymers made from natural amino acids, where the term “amino acid” includes organic compounds containing both a basic amino group and an acidic carboxyl group. Natural protein occurring amino acids, which make up natural polymers, include alanine, arginine, asparagine, aspartic acid, cysteine, glutamic acid, glutamine, glycine, histidine, isoleucine, leucine, lysine, methionine, phenylalanine, serine, threonine, tyrosine, tryptophan, proline, and valine.
The term “non-natural polymer” refers to any polymer comprising natural and non-natural monomers found in biology. For example, a ribosome can be designed to produce a non-naturally occurring biopolymer based on amino acids where naturally occurring and/or synthetic versions of naturally occurring components are used. For example, non-natural polymers could be made that comprise both natural and unnatural amino acids. These unnatural amino acids could comprise modified and unusual amino acids (e.g., D-amino acids and β-amino acids), as well as amino acids which are known to occur biologically in free or combined form but usually do not occur in proteins. Natural non-protein amino acids include arginosuccinic acid, citrulline, cysteine sulfinic acid, 3,4-dihydroxyphenylalanine, homocysteine, homoserine, ornithine, 3-monoiodotyrosine, 3,5-diiodotryosine, 3,5,5,-triiodothyronine, and 3,3′,5,5′-tetraiodothyronine. Modified or unusual amino acids include D-amino acids, hydroxylysine, 4-hydroxyproline, N-Cbz-protected amino acids, 2,4-diaminobutyric acid, homoarginine, norleucine, N-methylaminobutyric acid, naphthylalanine, phenylglycine, α-phenylproline, tert-leucine, 4-aminocyclohexylalanine, N-methyl-norleucine, 3,4-dehydroproline, N,N-dimethylaminoglycine, N-methylaminoglycine, 4-aminopiperidine-4-carboxylic acid, 6-aminocaproic acid, trans-4-(aminomethyl)-cyclohexanecarboxylic acid, 2-, 3-, and 4-(aminomethyl)-benzoic acid, 1-aminocyclopentanecarboxylic acid, 1-aminocyclopropanecarboxylic acid, and 2-benzyl-5-aminopentanoic acid.
As used herein, a “polymerase” refers to an enzyme that catalyzes the polymerization of nucleotides. “DNA polymerase” catalyzes the polymerization of deoxyribonucleotides. Known DNA polymerases include, for example, Pyrococcus furiosus (Pfu) DNA polymerase, E. coli DNA polymerase I, T7 DNA polymerase and Thermus aquaticus (Taq) DNA polymerase, among others. “RNA polymerase” catalyzes the polymerization of ribonucleotides. The foregoing examples of DNA polymerases are also known as DNA-dependent DNA polymerases. RNA-dependent DNA polymerases also fall within the scope of DNA polymerases. Reverse transcriptase, which includes viral polymerases encoded by retroviruses, is an example of an RNA-dependent DNA polymerase. Known examples of RNA polymerase (“RNAP”) include, for example, T3 RNA polymerase, T7 RNA polymerase, SP6 RNA polymerase and E. coli RNA polymerase, among others. The foregoing examples of RNA polymerases are also known as DNA-dependent RNA polymerase. The polymerase activity of any of the above enzymes can be determined by means well known in the art.
As used herein, a primer is “specific,” for a target sequence if, when used in an amplification reaction under sufficiently stringent conditions, the primer hybridizes primarily to the target nucleic acid. Typically, a primer is specific for a target sequence if the primer-target duplex stability is greater than the stability of a duplex formed between the primer and any other sequence found in the sample. One of skill in the art will recognize that various factors, such as salt conditions as well as base composition of the primer and the location of the mismatches, will affect the specificity of the primer, and that routine experimental confirmation of the primer specificity will be needed in many cases. Hybridization conditions can be chosen under which the primer can form stable duplexes only with a target sequence. Thus, the use of target-specific primers under suitably stringent amplification conditions enables the selective amplification of those target sequences that contain the target primer binding sites.
As used herein, “expression template” refers to a nucleic acid that serves as substrate for transcribing at least one RNA that can be translated into a polypeptide or protein. Expression templates include nucleic acids composed of DNA or RNA. Suitable sources of DNA for use a nucleic acid for an expression template include genomic DNA, cDNA and RNA that can be converted into cDNA. Genomic DNA, cDNA and RNA can be from any biological source, such as a tissue sample, a biopsy, a swab, sputum, a blood sample, a fecal sample, a urine sample, a scraping, among others. The genomic DNA, cDNA and RNA can be from host cell or virus origins and from any species, including extant and extinct organisms. As used herein, “expression template” and “transcription template” have the same meaning and are used interchangeably.
As used herein, “translation template” refers to an RNA product of transcription from an expression template that can be used by ribosomes to synthesize polypeptide or protein in vitro or in vivo.
As used herein, “cognate” as it modifies polypeptide with respect to protease substrates disclosed herein, refers to the natural polypeptide as expressed from an endogenous gene in the native cellular context. A protease that acts upon a cognate polypeptide is an endogenous protease that would usually act upon the polypeptide in the same native cellular context. By contrast, “non-cognate” as it modifies polypeptide, refers to a polypeptide substrate for a protease from a different native cellular context. Furthermore, a “heterologous” as it modifies polypeptide includes non-cognate polypeptides, fusion polypeptides and recombinant polypeptides that derive from a different native cellular context with respect to a given protease.
As used herein, “expression cassette” refers to a nucleic acid sequence that enables expression of an RNA having a defined nucleic acid sequence. The nucleic acid sequence can be either DNA or RNA, and the defined nucleic acid sequence can encode a polypeptide. The nucleic acid sequence can include sequences for initiating transcription (e.g., promoter and enhance elements) and terminating transcription; sequences for enhancing translation of the RNA to form polypeptides; and sequences that encode in-frame polypeptide leader and post-translational processing signals. For expression cassettes that produce polypeptides, the nucleic acid sequence can include sequences that encode for affinity tag motifs in-frame with the coding sequence for the polypeptide to enable affinity purification of the resultant polypeptide. For nucleic acid sequences composed of DNA, the expression cassette can include multiple cloning sites or polylinkers to enable cloning of polypeptide-coding genes in-frame with flanking sequences encoding for affinity tag motif(s), leader sequences and/or post-translational processing signals.
The term “tag” (or “tag motif”) refers to a sequence motif that does not normally form part of the native polypeptide to which the sequence motif is covalently linked. In this regard, a tag is a heterogeneous, non-cognate sequence motif with respect to the remainder of the polypeptide sequence. Where a polypeptide is initially synthesized as a precursor polypeptide that includes a leader peptide sequence, a tag also includes the leader peptide sequence with respect to the mature polypeptide. A tag may be covalently linked to the N-terminus, C-terminus or at an internal site (for example, a amino acid side chain) of a polypeptide. A tag can be used to detect, identify, select, enrich or purify the polypeptide to which the tag is covalently linked. A tag (or tag motif) can include a leader peptide sequence and/or an affinity tag.
The term “affinity tag” refers to a sequence that permits detection and/or selection of a polypeptide sequence. For the purposes of this disclosure, a recombinant gene that encodes a recombinant polypeptide may include an affinity tag. In particular, an affinity tag is positioned typically at either the N-terminus or C-terminus of the coding sequence for a polypeptide through the use of recombination technology. Exemplary affinity tags include polyhistine (for example, (His6)), maltose binding protein, glutathione-S-transferase (GST), HaloTag®, AviTag, Calmodulin-tag, polyglutamate tag, FLAG-tag, HA-tag, Myc-tag, S-tag, SBP-tag, Softag 3, V5 tag, Xpress tag, among others.
“Recombinant,” as used herein, refers to an amino acid sequence or a nucleotide sequence that has been intentionally modified by recombinant methods. By the term “recombinant nucleic acid” herein is meant a nucleic acid, originally formed in vitro, in general, by the manipulation of a nucleic acid by endonucleases, in a form not normally found in nature. Thus an isolated nucleic acid in a linear form, or an expression vector formed in vitro by ligating DNA molecules that are not normally joined, are both considered recombinant for the purposes of this invention. It is understood that once a recombinant nucleic acid is made and reintroduced into a host cell, it will replicate non-recombinantly, i.e., using the in vivo cellular machinery of the host cell rather than in vitro manipulations; however, such nucleic acids, once produced recombinantly, although subsequently replicated non-recombinantly, are still considered recombinant for the purposes of the invention. A “recombinant protein” is a protein made using recombinant techniques, i.e., through the expression of a recombinant nucleic acid as depicted above.
A nucleic acid is “operably linked” when it is placed into a functional relationship with another nucleic acid sequence. For example, a promoter or enhancer is operably linked to a coding sequence if it affects the transcription of the sequence; or a ribosome binding site is operably linked to a coding sequence if it is positioned so as to facilitate translation.
The term “vector” refers to a piece of DNA, typically double-stranded, which may have inserted into it a piece of foreign DNA. The vector may be, for example, of plasmid origin. Vectors contain “replicon” polynucleotide sequences that facilitate the autonomous replication of the vector in a host cell. Foreign DNA is defined as heterologous DNA, which is DNA not naturally found in the host cell, which, for example, replicates the vector molecule, encodes a selectable or screenable marker, or encodes a transgene. The vector is used to transport the foreign or heterologous DNA into a suitable host cell. Once in the host cell, the vector can replicate independently of or coincidental with the host chromosomal DNA, and several copies of the vector and its inserted DNA can be generated. In addition, the vector can also contain the necessary elements that permit transcription of the inserted DNA into an mRNA molecule or otherwise cause replication of the inserted DNA into multiple copies of RNA. Some expression vectors additionally contain sequence elements adjacent to the inserted DNA that increase the half-life of the expressed mRNA and/or allow translation of the mRNA into a protein molecule. Many molecules of mRNA and polypeptide encoded by the inserted DNA can thus be rapidly synthesized.
As used herein, “scar” refers to a remnant of polypeptide sequence attached to a mature polypeptide that does not form part of the natural amino acid sequence of the mature polypeptide. An example of a scar includes a portion of a leader sequence that is not proteolytically processed accurately from a natural precursor polypeptide to yield a natural mature polypeptide so that the portion of the leader sequence remains attached to the mature polypeptide. Another example of a scar includes a portion of a tag peptide of a precursor recombinant polypeptide that is not completely removed by a protease to generate the recombinant polypeptide without the tag.
As used herein “scarless tag removal” refers to the processing of a precursor polypeptide that contains a tag motif with a protease to yield a polypeptide product having no scar of the tag motif.
As used herein, “codon optimized” refers to a nucleic acid encoding an open reading frame of a polypeptide in which the codons have been selected to permit efficient expression of the polypeptide in a particular host organism or host cell. Exemplary host organisms and host cells (“expression hosts”) for expressing polypeptides (for example, recombinant proteins) include E. coli, S. cerevisiae, S. pombe, P. pastoris, insect cells (for Baculovirus expression), and various mammalian cell lines (for example, HeLa, Jurkat, 293, CHO and COS, among others). Model expression hosts for expressing heterologous polypeptides are known in the art and codon optimized heterologous gene sequences can be deduced from codon usage frequencies of highly expressed polypeptides in such organisms.
As used herein, “substantially identical” as the term modifies a biological composition, such as a nucleotide sequence or a polypeptide sequence, refers to a first primary sequence, including fragments thereof, having at least 75% identity of the intact primary sequence of the reference nucleotide sequence or a polypeptide sequence and/or a second primary sequence having at least 80% sequence homology of the primary sequence of the reference nucleotide sequence or a polypeptide sequence, wherein the first and second primary sequences have at least 70% of the functional activity of the reference nucleotide sequence or a polypeptide sequence.
As used herein, “biological composition” refers to a composition that includes a biological molecule, including for example, a nucleic acid or a polypeptide.
As used herein, “an equivalent thereof” refers to a biological composition, such as a nucleotide sequence or a polypeptide sequence, that encodes the identical or substantially identical structurally- and functionally-defined biological composition as the biological composition being referenced. Sequence homology among nucleotide sequences include nucleotide sequences having degenerate codons and codon-optimized sequences for expression in particular host organisms such that the nucleic acid sequences encode the same polypeptide. Sequence homology among polypeptide sequences include amino sequences having conservative structural changes in terms of hydrophobic, hydrophilic, and ionic side-chain properties such that the resultant polypeptides encode the same functional activity (e.g., identical substrate specificity).
As used herein, “a derivative thereof” refers to a biological composition having at least 10% of the activity of a reference biological composition. More preferably, “a derivative thereof” refers to a biological composition having greater than about 50% of the activity of a reference biological composition, such as about 60%, about 70%, about 90% and about 100% of the activity of a reference biological composition. An example of a LanP protease derivative includes a LanP polypeptide that lacks the signal sequence of the pro-form, nascent LanP polypeptide, such as a LanP polypeptide encoded within an organism genome. Additional examples of a LanP protease derivative includes a LanP polypeptide modified to include tag, such as an affinity tag. An example of a derivative of a recognition substrate for a LanP protease (“lanthipeptide protease substrate recognition sequence”) includes a polypeptide sequence having a P-X motif, wherein the P includes a polypeptide sequence recognized by the LanP protease and X includes an amino acid, wherein the LanP protease catalyzes cleavage of a recognition substrate at the bond between the P and X moieties of the P-X motif. With regard to a derivative of a recognition substrate for a LanP protease, activity refers to one of the rate of catalyzed reaction or the specificity of the cleavage. Thus, sequence variations in either P or X are included in a derivative of a recognition substrate for a LanP protease.
Class I and class II LanP proteases having sequence-specific protease activity are presented. Each of the proteases can serve as an efficient sequence-specific traceless protease for general traceless tag removal applications. Compositions and methods for preparing and using the novel proteases are disclosed herein.
To analyze the substrate specificity of ElxP, the sequences of several LanAs from class I lanthipeptides were aligned. The alignment clearly shows that the sequences near the proteolytic cleavage site group into two different types (
To test the hypothesis that the conserved motif in ElxA is important for ElxP activity, ElxA was expressed in Escherichia coli as an N-terminally hexahistidine tagged peptide and purified by metal affinity chromatography. ElxP [(SEQ ID NO: 4) (DNA); (SEQ ID NO: 5) polypeptide] was expressed in E. coli fused to the C-terminus of maltose binding protein (MBP-ElxP; SEQ ID NO: 6 (DNA); SEQ ID NO: 7 (polypeptide)). We then performed alanine scanning mutagenesis on the E/D-L/V-x-x-Q motif present in the ElxA leader peptide. Indeed, single alanine mutations at the Q-1, L-4, and D-5 positions in the leader peptide of ElxA significantly reduced the cleavage efficiency of MBP-ElxP as observed by matrix-assisted laser desorption ionization time-of-flight mass spectrometry (MALDI-TOF MS) (Table 1).
1Observed masses are shown with the calculated mass in parenthesis. MBP-ElxP (5 μM) was incubated with mutant or wild type His6-ElxA (50 μM in reaction buffer (50 mM Tris HCl pH 8.0) at room temperature for 2 h. Reactions were analyzed by MALDI-TOF MS. Results are based on the amount of substrate left after the assay based on normalized ion intensities.
To quantify and distinguish the contribution of each amino acid to the substrate specificity of ElxP, we next determined the kinetic parameters of MBP-ElxP by using the wild type peptide and three ElxA mutants Q-1A, P-2A and L-4A as substrates. Reversed phase high performance liquid chromatography (RP-HPLC) was used to monitor N-terminal leader peptide formation at different substrate concentrations in the assay. The analysis showed that MBP-ElxP cleaved the wild type ElxA with an efficiency of ˜240 M-'s 1 (
Although it is possible that proteolysis by wild type ElxP of the ElxA peptide possessing the thioether rings would be more efficient, the catalytic efficiency of MBP-ElxP observed in this study with linear peptides is sufficient for application of the enzyme as a sequence specific protease.
Many leader peptides of class I lanthipeptides share an F-D/N-L-N/D sequence motif (
Insertion of the ElxP Recognition Motif into the NisA Peptide.
Based on our data, the conserved E/D-L/V-x-x-Q-T1/S1 motif (SEQ ID NO: 3) present in the ElxA-group of LanAs likely serves as the main recognition element for their LanPs. This sequence could possibly be used as a tool to selectively remove tags from fusion proteins or leader peptides from other RiPPs. To determine this potential, we analyzed the ElxP activity on the non-cognate lanthipeptide precursor peptide NisA and NisA mutants (
Previous attempts to use the dehydratase ElxB, the cyclase ElxC, and the peptidase ElxP to generate dehydroepilancin 15X, the substrate of the dehydrogenase ElxO, were unsuccessful. However, we showed that His6-ElxO catalyzes the conversion of the synthetic peptide Pyr-AAIVK, resembling the N-terminal region of dehydroepilancin 15X (
Attempts to determine the steady state kinetic parameters kcat and Km using a subset of peptides were not successful, since it was not possible to saturate the enzyme with the substrates before reaching the peptide solubility limits. Therefore, the kinetic constants kcat/Km were determined by measuring the reaction rates at various peptide concentrations (Table 2).
For all tested substrates, the values of kcat/Km were relatively small, presumably because the peptides are lacking structural features compared to the expected physiological substrate, such as the thioether rings or additional amino acids. The smaller peptides Pyr-AAIV and Pyr-AAI (Table 2, entries 2 and 3), but not Pyr-AA and Pyr-A (Table 2, entries 4 and 5), were reduced by His6-ElxO in the presence of NADPH based on LC-MS analysis, although with considerably lower reaction rates compared with Pyr-AAIVK. In contrast, the peptide Pyr-AAIVKBBIKAAKK, where B stands for L-2-aminobutyric acid, was converted at a higher rate (Table 2, entry 6), suggesting that the length of the peptide is important for substrate recognition. The Ala scanning analysis performed along the sequence of Pyr-AAIVK (Table 2, entries 7-9) indicated that the enzyme was able to reduce all the peptides tested albeit with a lower kcat/KM for Pyr-AAAVK (Table 2, entry 9).
Next, we evaluated peptides containing amino acids with nonpolar (Pro, Met, Gly, Ile, Val, Phe; entries 14, 15, and 19), polar (Asn, Thr, Tyr; entries 13 and 20), acidic (Asp; entry 12), or basic (Arg, Lys, His; Table 2, entries 10, 11, and 21) residues at position 2, and found that they were all transformed to the reduced products. These results suggest that no substrate residue is absolutely required for enzymatic activity and that the minimal length of the peptide to be accepted as substrate is four residues. Interestingly, Pyr-RAIVK, Pyr-KAIVK, Pyr-NAIVK, and Pyr-MAIVK were better substrates for ElxO than Pyr-AAIVK (Table 2, entries 10, 11, 13, 15), whereas Pyr-DAIVK and Pyr-PAIVK were converted considerably less efficiently (Table 2, entries 12 and 14), suggesting that negatively charged residues and Pro in position 2 are not well tolerated.
OBu-AAIVK and OBu-RAIVK were also accepted as substrates leading to the formation of an N-terminal 2-hydroxybutyryl group (Hob), although at lower rates (Table 2, entries 16 and 17). Similarly, the peptides OBu-AAAVK and OBu-AAIAK were substrates for the enzyme. However, when Pyr was substituted by a glyoxyl (Glx) group, such as in the peptide Glx-AAIVK (Table 2, entry 18), no significant formation of the reduced peptide was observed.
In addition to epilancin 15X, two other lantibiotics, epilancin K7 and epicidin 280, contain an N-terminal Lac moiety. To explore the potential of using His6-ElxO for the synthesis of other lantibiotics, peptides mimicking the N-terminal portion of dehydroepilancin K7 (Pyr-AAVLK) and dehydroepicidin 280 (Pyr-LGPAIK) were synthesized and tested as substrates. His6-ElxO reduced both peptides, even though their sequences are quite different from the N-terminus of epilancin 15X. Similarly, peptides resembling the N-terminus of lactocin S (Pyr-APVLA and Pyr-BPVLAAVAVAKKK) and Pep5 (OBu-AGPAIR) were incubated with His6-ElxO in the presence of NADPH. All the peptides were reduced as confirmed by LC-MS analysis.
Encouraged by these results with short peptides we next turned to lactocin S, a 37-residue lantibiotic (
Although all LanPs characterized to date belong to the subtilisin-like serine endopeptidases superfamily, their cleavage sequences are more diverse than those of class II lanthipeptide peptidases. Our current work provides evidence that the LanA and LanP proteins likely co-evolved and that LanP sequence specificity is mainly determined by the amino acids near the proteolytic site. Specifically, the conserved E/D-L/V-x-x-Q-T1/S1 sequence motif (SEQ ID NO: 3) present in the ElxA-group of LanAs provides the full recognition elements for ElxP. This recognition sequence may find use in applications of ElxP for cleaving off fusion tags or removing leader peptides from RiPPs.
CylA shows considerable homology to class I LanPs but is located in a separate clade in a Markov chain Monte Carlo phylogenetic tree (
Based on its sequence, CylA is a subtilisin-like serine protease with a conserved catalytic triad consisting of aspartate, histidine and serine. Given that CylA cleaves its substrates CylLL and CylLS after a Glu, we wondered whether the observed cleavage of CylA could be autocatalytic. If so, the proteolysis should be abolished by disrupting the catalytic triad. We therefore constructed another CylA mutant with the catalytic Ser359 changed to Ala. The mutant protein was expressed and purified using the same procedure as that for wild type CylA. Indeed, only one band was observed by SDS-PAGE for purified His6-CylA-27-412-S359A, corresponding to the full length protein. The expression of full length His6-CylA-27-412-S359A was further confirmed by MALDI-TOF MS (
With purified CylA in hand, we set out to test its proteolytic activity with the peptides CylLL and CylLS. A previous study reported that CylA catalyzed the removal of the 6-residue sequence GDVQAE from the N-terminus of both modified core peptides subsequent to leader peptide removal at the GS motif by CylB in the producing strain Enterococcus faecalis. In this work, dehydrated and cyclized CylLL and CylLS peptides were obtained by co-expression of CylLL and CylLS peptides with their lanthionine synthetase CylM in E. coli. Instead of using the membrane bound protein CylB, we employed the commercial protease AspN, which specifically cleaved N-terminal to Asp-5, leaving 5 amino acids (DVQAE) on the core peptides. These peptides were then incubated with CylA and the 5 amino acids were successfully removed (
Based on these results, we hypothesized that CylA specifically recognizes the GDVQAE sequence (SEQ ID NO: 46). To test this hypothesis, the GDVQAE sequence was engineered into HalA2, the precursor peptide for the lantibiotic haloduracin β (Halβ). Halβ and haloduracin α (Halα) constitute a two-component lantibiotic. The putative recognition sequence was installed between the HalA2 leader and core peptides by substituting the residues at positions −6 to −1. This HalA2-GDVQAE peptide was co-expressed with the cognate lanthionine synthetase HalM2 in E. coli, resulting in the desired 7 dehydrations (
To further evaluate the substrate scope of CylA, we explored its activity against a variant of the precursor peptide for Halα. Again, the residues at positions −6 to −1 were replaced with GDVQAE by site-directed mutagenesis to produce HalA1-GDVQAE. Halα does not show any sequence homology with the two cytolysin peptides or Halβ (
To test whether CylA accepts linear peptides containing the GDVQAE sequence other than CylLL and CylLS, we engineered the recognition sequence into two more peptides—ProcA1.7 and NisA. Upon incubation with CylA, the leader peptides of both His6-ProcA1.7-GDVQAE and His6-NisA-GDVQAE were successfully removed, indicating the broad substrate scope of CylA (
[a]These peptides were substrates with the linear sequences shown and also after the Cys in the 5th position formed a methyllanthionine with the Thr at position 1.
[b]Peptide was modified by HalM2 with a MeLan residue at the P1′ position.
[c]Peptide was modified by HalM1 with either a Cys or a disulfide linkage at the P1′ position.
[d]GDVQAE sequence inserted between the plasmid encoded His6-tag sequence and ProcA1.7.
The enzyme tolerates hydrophobic, hydrophilic, branched and aromatic amino acids in the P1′ position. Its substrate scope is not limitless, however, as it did not accept Glu or Lys in the P1′ position.
We next returned to the importance of the autocatalytic processing step for activity. Incubation of modified CylLS with His6-CylA-27-412-E95A resulted in cleavage, suggesting that the processing event is not absolutely required for proteolytic activity. To assess the effect of self-cleavage on the rate of cleavage, CylA-96-412 and His6-CylA-27-412-E95A were incubated with modified CylLS, and the formation of CylLS″ was monitored by liquid chromatography MS (LC/MS). CylA-96-412 catalyzed the proteolysis of 20 substrate peptides per minute under the condition we tested, whereas His6-CylA-27-412-E95A exhibited an approximate 10-fold lower rate of producing CylLS″ (
In vitro characterization of LicP, a class II LanP protease, involved in the biosynthesis of the lantibiotic lichenicidin, revealed a self-cleavage step that removes 100 amino acids from the N-terminus. Investigation of its substrate specificity demonstrated that LicP can serve as an efficient sequence-specific traceless protease. Encouraged by these findings for LicP, we identified 12 other class II LanPs, nine of which were previously unknown, and suggest that these proteins may serve as a pool of proteases with diverse recognition sequences for general traceless tag removal applications, expanding the current toolbox of proteases.
The licP gene was amplified from genomic DNA of B. licheniformis ATCC 14580 and cloned into an expression vector. A hexa-histidine tag was installed at its N terminus, and the first 24 amino acids of LicP, which correspond to a secretion signal peptide, were omitted. Upon expression in Escherichia coli BL21 (DE3) and purification using immobilized metal affinity chromatography, two bands were observed by gel electrophoresis (
Such proteolytic processing has been reported for several extracellular class I LanP proteases and was suggested to be autocatalytic like other subtilisins. To test whether this mechanism applied to the class II enzyme LicP, we substituted the predicted catalytic Ser376 with Ala. Unfortunately, His6-LicP-25-433-S376A was expressed almost exclusively in the insoluble fraction. We also mutated His186 predicted to be involved in the catalytic triad, but His6-LicP-25-433-H186A was also expressed insolubly. We eventually were able to obtain a very small amount of soluble His6-LicP-25-433-S376A, demonstrating that indeed the proteolytic cleavage after Glu100 was abolished (
We next tested LicP activity against the substrate peptide. It has been suggested that LicP trims off the 6-residue oligopeptide NDVNPE from LicA2′ to generate mature LicP. In this work, dehydrated and cyclized LicA2 was obtained by co-expressing LicA2 with its cognate lanthionine synthetase LicM2 in E. coli (
We further investigated whether the enzyme displays a preference for modified or linear LicA2. Liquid chromatography-based kinetic analysis of the time and concentration dependence of the cleavage reactions was hampered by the poor solubility of the LicA2 and Licβ peptides. Instead, we employed a competitive MALDI-TOF MS assay for a semi-quantitative time-dependent analysis at one substrate concentration, in which LicP was supplied to an equimolar mixture of modified and linear LicA2 and the production of leader peptides was monitored over time. In order to differentiate the otherwise identical leader peptides after proteolysis, we introduced a Pro to Gly mutation between the hexa-histidine tag and the precursor peptide in linear LicA2 (G-LicA2). The leader peptides obtained by complete proteolysis of equimolar amounts of modified and linear LicA2 exhibited comparable signal intensities when monitored by MALDI-TOF MS, confirming that the Pro to Gly mutation does not alter the ionization efficiency significantly (
The observation that LicP removes the oligopeptide NDVNPE and the entire leader peptide from modified or linear LicA2 suggests that it specifically recognizes the NDVNPE sequence but is rather tolerant of other regions of the peptides. We decided to test this hypothesis in the lanthipeptide family, as site-specific removal of leader peptides is critical for producing lanthipeptides in vitro and this step is often challenging as only a limited choice of proteases is available. ProcA1.7 and NisA, the precursor peptides of the lanthipeptides prochlorosin 1.7 and nisin, were mutated to substitute the last six residues of their leader peptides with the NDVNPE sequence (
The experiments with linear and modified LicA2 as well as ProcA1.7-NDVNPE, NisA-NDVNPE and MBP-BamL demonstrated that LicP tolerates Dhb, Thr, Ser and Ile in the P1′ position. To further evaluate its tolerance, we altered the P1′ position in NisA-NDVNPE from Ile to eight other amino acids (Gly, Cys, Thr, Leu, Phe, Trp, Glu and Lys;
ahe vertical bar (|) separating the P and P′ positions denotes the cleavage site for the specified substrate.
bRT: room temperature.
cPerformed at 4º C.
The substrate specificity of LicP was further evaluated by a gel-based assay monitoring the time-dependent cleavage of mutants of linear LicA2. The presence of a Glu at the P1 position was critical for LicP activity as LicA2-E-1A was not a substrate under the assay conditions and even substitution with structurally related amino acids Asp and Gln was not tolerated (
LanP genes are not often found in class II lanthipeptide gene clusters. Only four class II LanPs have been reported to date—LicP, CylA, CerP and CmP, which have been suggested to remove six-residue sequences at the N terminus of Licβ, cytolysin, cerecidins and carnolysin, respectively. Among them, the proteolytic activity, and hence the identity of the cleavage sites, has been confirmed for CylA, CerP and LicP. To identify additional class II LanP proteins and potentially identify additional recognition sequences that might be useful, we performed a search of the UniProtKB database with the LicP protease domain (LicP-101-433) as a query and the non-redundant protein sequence database with LicA2 as a query using the default Blast parameters for proteins provided by the National Center for Biotechnology Information (NCBI) website. The first 250 hits were subjected to further analysis and several were correlated to class II lanthipeptide biosynthesis by the observation of nearby genes encoding LanM proteins. Nine representative class II lanthipeptide gene clusters with LanP genes are shown in
B. licheniformis ATCC 14580
Enterococcus faecalis
B. licheniformis 9945A
Bacillus cereus Q1
Bacillus cereus FRI-35
Kyrpidia tusciae DSM 2912
Enterococcus caccae ATCC
Bacillus cereus VPC1401
Carnobacterium maltaromaticum
Bacillus bombysepticus
Bacillus thuringiensis DB27
Planomicrobium glaciei CHR43
Bacillus cereus VD045
Bacillus cereus VD156
Thus, the first heterologous expression of LanP proteins responsible for class II lanthipeptide biosynthesis is provided herein. We successfully reconstituted CylA's activity in vitro. In addition to its physiological role of removing the six residues at the N terminus of CylLL′ and CylLS′, CylA was capable of removing the entire leader peptides of modified CylLL and CylLS. A turnover rate of 20 min−1 was observed, indicating CylA is an efficient protease. In contrast to NisP and FlaP that have been reported to exhibit a preference for modified NisA and FlaA over the unmodified peptides, CylA also removed the leader peptides of linear CylLL and CylLS. Although multiple groups have reported that mature LanPs purified from their producing strains lacked 95-195 residues from the N terminus, our results serve as the first evidence with that a LanP protease employs an autocatalytic activation mechanism to cleave its lanthipeptide substrate. Our observations for CylA and LicP combined with the reported results for NisP and EpiP strongly suggest that all secreted LanPs may undergo self-cleavage and employ the self-cleaving-activation mechanism.
CylA was also active against unrelated peptide substrates as long as the recognition sequence was installed—it accepted a range of residues in the P1′ site such as glycine, aromatic residues (phenylalanine, tryptophan), branched residues (isoleucine), or modified residues (MeLan, disulfide linked cysteine), strongly suggesting its potential as a general traceless tag removal protease. CylA protein was stable at −20° C. and no obvious decrease of activity was observed after multiple rounds of freeze-thaw. The identification of new LanP-containing gene clusters for class II lanthipeptide biosynthesis indicates that class II LanPs occur more widely than previously believed. Although the recognition sequences of these currently identified class II LanPs show a certain level of homology, they also exhibit considerable diversity. As a result, these LanPs may serve as a basis to construct a protease pool for general traceless tag removal purposes.
Although LicP favors modified LicA2 over linear LicA2, which indicates that post-translational modifications in the core peptide contribute to LicP's substrate recognition in addition to the NDVNPE sequence, our observations with substrate analogs demonstrate its application as a sequence-specific protease for traceless removal of leader peptides and an expression tag. The substrate specificity of LicP was identified using both structural information and biochemical characterizations. The P5, P4 and P1 residues of LicA2 were found to be important for LicP recognition. These three sites were also suggested as the origin of specificity for a class I LanP, ElxP, as determined by kinetic analysis based on LC quantification. The similarity in the important positions suggests a general substrate recognition mechanism by the entire subtilisin-like LanP family.
The thermostability of subtilisin BPN′ and related proteases is enhanced significantly in the presence of calcium ions, which are necessary for maturation, and subsequent stabilization of a large loop in the catalytic domain. The calcium dependence constitutes a drawback for industrial utility of subtilisin BPN′. Much effort has been spent on engineering thermostable mutants of subtilisin that function in a calcium-independent manner. The structural and biochemical analysis of LicP (data not shown) reveals an elegant solution to this limitation, as maturation and subsequent stabilization of the enzyme is facilitated not by metal ions, but rather by the insertion of Trp111, liberated by cleavage of the linker between the prodomain and the catalytic domain, into a hydrophobic pocket located in the same vicinity as the calcium-binding site in subtilisin BPN′. A recent structure of the class I lanthipeptide protease NisP also demonstrated loss of a calcium binding site, although unlike the structure of LicP, the prodomain was not present in the NisP structure and its substrate specificity was not investigated.
Over the past several decades, the toolbox of useful proteases has been significantly enlarged. Several proteases with strict recognition sequences have been commercialized for biochemical or industrial applications, including factor Xa, enterokinase, and TEV protease. Factor Xa and enterokinase exhibit trypsin-like activity and cleave after an Arg or Lys. TEV protease recognizes a larger motif and exhibits better reliability in terms of specificity, but TEV protease requires either a Gly or Ser at the P1′ position for efficient cleavage. LicP is complementary in that it specifically cleaves after a Glu in the NDVNPE sequence, and is quite tolerant of various residues in the P1′ position. LicP accepted a range of residues at the P1′ site (Table 4) such as glycine, small polar residues (Ser, Thr, Cys) and large aliphatic residues (Ile). LicP also processed peptides with Leu, aromatic (Phe, Trp) and charged (Lys, Glu) residues at the P1′ position albeit with reduced efficiency. Additional favorable properties include its stability demonstrated by the persistent activity of LicP after 12 weeks at 4° C., and no obvious decrease in activity after multiple rounds of freeze-thaw procedures.
This disclosure identifies ten new class II lanthipeptide gene clusters containing lanP genes, suggesting they are more widely distributed than previously expected. Although the putative recognition sequences of these newly identified LanPs show a certain level of homology, they also exhibit considerable diversity. Similar to other proteases, most of these LanPs are predicted to cleave after charged residues such as arginine, glutamate or aspartate, but a few appear to cleave after unusual P1 residues such as histidine or alanine that are rarely the site of cleavage for other proteases. We confirmed the predicted sites for two examples that have a His and an Arg in the P1 position. Hence, this naturally occurring protease family may serve as a basis to construct a general protease pool for traceless tag removal purposes.
The discovery of novel lanthipeptide protease polypeptides and their substrate recognition rules, including the robust portability of their substrate recognition sites into heterologous polypeptide contexts, provides fundamentally new and non-obvious approaches to generating mature polypeptides or recombinant polypeptides completely devoid of extraneous N-terminal sequences, such as leader sequences or tag sequences (that is, scarless or traceless of leader or tag sequences). The present disclosure provides several aspects having broad utility in biotechnology, medical and diagnostic applications of lanthipeptide proteases for processing cognate lanthipeptides, noncognate lanthipeptide and heterologous peptides that can be suitably processed by lanthipeptide proteases to yield scarless tag polypeptide products.
In one aspect, an isolated nucleic acid comprises an open reading frame encoding a lanthipeptide protease polypeptide for scarless tag removal from a polypeptide is provided. In this regard, the lanthipeptide protease is codon optimized for expression in an expression host. The expression host can be selected from E. coli, S. cerevisiae, S. pombe, P. pastoris, an insect cell, a HeLa cell, a Jurkat cell, a 293 cell, a CHO cell and a COS cell. In these aspects, the lanthipeptide protease polypeptide can be selected from SEQ ID NOS: 5, 7, 9-25, 29 and 30, including equivalents thereof and derivatives thereof. Furthermore, in this respect, the lanthipeptide protease polypeptide can recognize a substrate recognition sequence selected from SEQ ID NOS: 1-3, 27, 31-46 and sequences of Table 3, including equivalents thereof and derivatives thereof. Additionally, the polypeptide can be a cognate lanthipeptide, a non-cognate lanthipeptide or a heterologous polypeptide.
The isolated nucleic acid can further include a vector having a transcription controlling signal, wherein the isolated nucleic acid can be operably linked to the transcriptional controlling signal to enable expression of the lanthipeptide protease polypeptide. In this regard, the transcriptional controlling signal of the nucleic acid can include a transcriptional initiation element. In this regard, the transcriptional controlling signals can further include a transcriptional termination element. The isolated nucleic acid can further include a translational controlling signal. Exemplary translational controlling signals include at least one selected from a translational enhancer and a post-translational processing element.
Recombinant DNA and molecular biology and biochemical methods for carrying out the preparations of these reagents are well understood in the art.
In another aspect, an expression cassette including an open reading frame for a polypeptide, wherein the open reading frame encodes a substrate recognition sequence for a lanthipeptide protease polypeptide, is provided. In this aspect, the substrate recognition sequence is selected from SEQ ID NOS: 1-3, 27, 31-46 and sequences of Table 3, including equivalents thereof and derivatives thereof. In this aspect, the lanthipeptide protease polypeptide is selected from SEQ ID NOS: 5, 7, 9-25, 29 and 30, including equivalents thereof and derivatives thereof. In this aspect, the polypeptide is selected from a cognate lanthipeptide, a non-cognate lanthipeptide or a heterologous polypeptide.
In another respect, this aspect further includes a transcription controlling signal, wherein the isolated nucleic acid is operably linked to the transcriptional controlling signal to enable expression of the polypeptide. In this regard, the transcriptional controlling signal includes a transcriptional initiation element. In one respect, the transcriptional controlling signals can further include a transcriptional termination element. In these latter respects, a translational controlling signal can be included. For example, the translational controlling signal can include at least one selected from a translational enhancer and a post-translational processing element.
Recombinant DNA and molecular biology and biochemical methods for carrying out the preparations of these reagents are well understood in the art.
In another aspect, an isolated polypeptide comprising the structure: T-R-P is provided, wherein T comprises a tag motif, R comprises a lanthipeptide protease substrate recognition sequence and P comprises an open reading frame encoding a polypeptide without the tag motif and lanthipeptide protease substrate recognition sequence.
In this respect, the isolated polypeptide can be codon optimized for expression in an expression host. Preferred expression hosts in this aspect include those selected from E. coli, S. cerevisiae, S. pombe, P. pastoris, an insect cell, a HeLa cell, a Jurkat cell, a 293 cell, a CHO cell and a COS cell. Protein expression in these systems is well known in the art. Moreover, the lanthipeptide protease substrate recognition sequence in this aspect can be selected from SEQ ID NOS: 1-3, 27, 31-46 and sequences of Table 3, including equivalents thereof and derivatives thereof.
With respect to the T comprising a tag motif, the tag motif can include an affinity tag. In this regard, the affinity tag can be preferably selected from polyhistine, maltose binding protein, glutathione-S-transferase, HaloTag®, AviTag, Calmodulin-tag, polyglutamate tag, FLAG-tag, HA-tag, Myc-tag, S-tag, SBP-tag, Softag 3, V5 tag and Xpress tag. In the foregoing aspects, the polypeptide is a cognate lanthipeptide, a non-cognate lanthipeptide or a heterologous polypeptide. These tag systems are well known in the art.
In another aspect, a method of scarless tag removal from a polypeptide is provided. The method includes two steps. The first step includes providing the polypeptide, wherein the polypeptide includes the structure: T-R-P. T comprises a tag motif, R comprises a lanthipeptide protease substrate recognition sequence and P comprises an open reading frame encoding a polypeptide without the tag motif and lanthipeptide protease substrate recognition sequence. The second step includes subjecting the polypeptide to a lanthipeptide protease having specificity for catalyzing proteolytic cleavage at the lanthipeptide protease substrate recognition sequence, thereby providing the polypeptide without a tag scar. In another aspect, the method further includes a step of purifying the polypeptide without a tag scar.
In these aspects, the lanthipeptide protease can be codon optimized for expression in an expression host. Suitable expression hosts in this regard include any of those selected from E. coli, S. cerevisiae, S. pombe, P. pastoris, an insect cell, a HeLa cell, a Jurkat cell, a 293 cell, a CHO cell and a COS cell. Protein expression in these systems is well known in the art. With respect to all of these aspects, the lanthipeptide protease polypeptide can be selected from SEQ ID NOS: 5, 7, 9-25, 29 and 30, including equivalents thereof and derivatives thereof. Likewise with respect to these aspects, lanthipeptide protease substrate recognition sequence is selected from SEQ ID NOS: 1-3, 27, 31-46 and sequences of Table 3, including equivalents thereof and derivatives thereof.
Similarly, the tag motif can include an affinity tag. In some aspects, the affinity tag can be selected from polyhistine, maltose binding protein, glutathione-S-transferase, HaloTag®, AviTag, Calmodulin-tag, polyglutamate tag, FLAG-tag, HA-tag, Myc-tag, S-tag, SBP-tag, Softag 3, V5 tag and Xpress tag. In some aspects, the polypeptide can be selected from a cognate lanthipeptide, a non-cognate lanthipeptide or a heterologous polypeptide. These tag systems are well known in the art.
In all of these aspects, the polypeptide is expressed in vivo or in vitro. In one respect, the polypeptide is expressed in vivo from an expression cassette in an expression host. In this regard, a suitable expression host can be selected from E. coli, S. cerevisiae, S. pombe, P. pastoris, an insect cell, a HeLa cell, a Jurkat cell, a 293 cell, a CHO cell and a COS cell. In another aspect, polypeptide is expressed in vitro from an expression cassette in a coupled transcription-translation system or from a translation template in a translation system. These systems and methods of expression are well known in the art.
In another aspect, a kit for expressing a polypeptide without a tag scar is provided. The kit includes two components. The first component includes an expression vector that includes an expression cassette, wherein the expression cassette can encodes a polypeptide that includes the structure: T-R-P. T comprises a tag motif, R comprises a lanthipeptide protease substrate recognition sequence and P comprises an open reading frame encoding a polypeptide without the tag motif and lanthipeptide protease substrate recognition sequence. The second component includes a lanthipeptide protease having specificity for catalyzing proteolytic cleavage at the lanthipeptide protease substrate recognition sequence, thereby providing the polypeptide without the tag scar. In a further refinement of this aspect, the kit includes a reagent to purify the polypeptide without the tag scar.
With respect to both of these aspects, an additional refinement includes an expression host. In this respect, the expression host can be selected from E. coli, S. cerevisiae, S. pombe, P. pastoris, an insect cell, a HeLa cell, a Jurkat cell, a 293 cell, a CHO cell and a COS cell.
The basic kit can include an lanthipeptide protease that is codon optimized for expression in the expression host. In this regard, the expression host can be selected from E. coli, S. cerevisiae, S. pombe, P. pastoris, an insect cell, a HeLa cell, a Jurkat cell, a 293 cell, a CHO cell and a COS cell. With respect to any of the foregoing aspects, the lanthipeptide protease polypeptide can be selected from SEQ ID NOS: 5, 7, 9-25, 29 and 30, including equivalents thereof and derivatives thereof. With respect to any of the foregoing aspects, the lanthipeptide protease substrate recognition sequence can be selected from SEQ ID NOS: 1-3, 27, 31-46 and sequences of Table 3, including equivalents thereof and derivatives thereof.
With respect to any of the foregoing aspects, the tag motif comprises an affinity tag. The affinity tag can be selected from polyhistine, maltose binding protein, glutathione-S-transferase, HaloTag®, AviTag, Calmodulin-tag, polyglutamate tag, FLAG-tag, HA-tag, Myc-tag, S-tag, SBP-tag, Softag 3, V5 tag and Xpress tag. With respect to any of the foregoing aspects, the polypeptide is a cognate lanthipeptide, a non-cognate lanthipeptide or a heterologous polypeptide.
Examples 1-9 are directed to ElxP protease activity, substrate recognition rules and demonstration of ElxP-mediated cleavage of heterologous fusion polypeptides. Examples 10-24 are directed to CylA and LicP protease activities and substrate recognition rules.
All oligonucleotides used in this study were purchased from Integrated DNA Technologies and are presented in Table 6 For microorganisms used, see Table 7
Lactobacillus sake L45
Pediococcus acidilactici Pac1.0
1Sources/References: DH5α (Grant, S. G. et al. Proc. Natl. Acad. Sci. U.S.A. 87, 4645-4649 (1990); BL21 DE3 and Rosetta2 (Novagen [EMD Millipore]); Lactobacillus sake L45 and Pediococcus acidilactici Pac 1.0 (Mortvedt, C. I. et al. Appl. Environ. Microbiol. 57, 1829-1834 (1991)).
Reagents used for molecular biology were purchased from New England BioLabs, Thermo Fisher Scientific, or Gold Biotechnology. Plasmid sequencing was performed by ACGT Inc. unless otherwise noticed. Escherichia coli DH5C and BL21 (DE3) were used for plasmid maintenance and protein or peptide overexpression, respectively. The strains L sake L45 and P. acidilactici Pac 1.0 were grown in de Man-Rogosa-Sharpe (MRS) solid agar or broth. MALDI-TOF measurements were performed using a Bruker UltrafleXtreme MALDI-TOF-TOF instrument using a positive reflective mode and sinapinic acid as a matrix unless otherwise noted.
The LanP sequences were aligned in ClustalX using default parameters with iteration at each alignment step, and the alignments were manually fine-tuned afterwards. Bayesian inference was used to calculate posterior probability of clades utilizing the program MrBayes (version 3.2). Final analyses consisted of two sets of eight chains each (one cold and seven heated), run to reach a convergence with standard deviation of split frequencies <0.005. Posterior probabilities were averaged over the final 75% of trees (25% burn in). The analysis utilized a mixed amino acid model with a proportion of sites designated invariant, and rate variation among sites modeled after a gamma distribution divided into eight categories, with all variable parameters estimated by the program based on BioNJ starting trees. Accession numbers are listed in Table 8.
Enterococcus faecalis
Bacillus licheniformis DSM 13
Lactococcus lactis subsp. lactis
Lactococcus lactis
Streptococcus salivarius
Staphylococcus epidermidis
Staphylococcus gallinarum
Staphylococcus aureus subsp.
aureus MW2
Staphylococcus aureus subsp ED133
Streptococcus uberis
Staphylococcus epidermidis
Staphylococcus epidermidis
Staphylococcus epidermidis
Bacillus subtilis BSn5
Cloning, Expression, and Purification of MBP-EIxP
Primers used for the construction of mutant substrates are listed in Table 6. The cloning of the gene encoding ElxP is described in Velasquez, J. E. et al. Chem. Biol. 18, 857-867 (2011). An aliquot of 50 ng of pelB-mbp-elxP-pET28b was used to transform 50 L of electrocompetent E. coli BL21 (DE3) Rosetta 2 cells following standard procedures. After the incubation period, cells were plated on LB agar (LBA) plates supplemented with kanamycin (kn, 25 μg mL−1) and chloramphenicol (cm, 12.5 μg mL−1) and grown at 37° C. overnight (O/N). A single colony was used to inoculate 1 mL of fresh LB supplemented with kn and cm and 1 μL was plated for a second time on LBA plates supplemented with kn and cm. Plates were kept at 37° C. O/N. For overexpression, 1 mL of LB supplemented with kn and cm was used to scrape the cells out of the O/N LBA plates to be used as initial inoculum. Overexpression was performed in 6 L of LB supplemented with kn and cm with a starter OD600 of 0.025. Cultures were incubated at 37° C., 250 rpm until the OD600 reached ˜1.0. At this OD, cultures were chilled on ice for 15 min and protein expression was induced with 0.1 mM isopropyl-β-D-1-thiogalactopyranoside (IPTG). Protein overexpression was carried out at 18° C., 250 rpm for 16 h. The cells were harvested (6976×g, 20 min, 4° C.), resuspended in 50 mL of lysis buffer (20 mM Tris-HCl pH 8.0, 500 mM NaCl, 10% (v/v) glycerol, 1 mM EDTA) and lysed using an EmulsiFlex-C3 homogenizer with a pressure of less than 500 bar. To remove cell debris, the lysed fraction was centrifuged (22,789×g, 4° C., 30 min) and the supernatant was cleared through a 0.45 μm syringe-tip filter (Millipore). The protein was purified by affinity chromatography using an ÄKTA purifier (GE healthcare) equipped with an MBPTrap HP 5 mL column pre-packed with Dextrin Sepharose™ (GE healthcare) according to the manufacturer's protocol. After loading the supernatant into the column pre-equilibrated with lysis buffer, the column was washed with lysis buffer until a stable baseline based on the absorbance at 280 nm was reached. Protein was eluted with a linear gradient from 0-100% (v/v) of elution buffer (20 mM Tris-HCl pH 8.0, 500 mM NaCl, 1 mM EDTA, 10 mM maltose) in lysis buffer over 30 min. Collected fractions were analyzed by SDS-PAGE using a 4-20% TGX mini-protean gel (BioRad) and visualized by Coomassie staining. Fractions containing the desired MBP-ElxP were collected and desalted using gel filtration chromatography on a HiLoad 16/60 column packed with Superdex 200 PG eluting with 1 mL min−1 of gel filtration buffer (300 mM NaCl, 20 mM Tris-HCl pH 8.0, 10% (v/v) glycerol). Protein was concentrated using an Amicon Ultra centrifuge tube of 30 kDa molecular weight cut off (MWCO) (3,488×g, 30 min, 4° C.) (Millipore). The concentration of the purified protein was determined spectrophotometrically using a calculated molar extinction coefficient of 113,485 M−1 cm−1 and a molecular weight of 80.504 kDa. Purity was assessed by SDS-PAGE analysis. Aliquots were frozen in liquid nitrogen for future use and stored at −80° C.
Cloning, Expression, and Purification of His6-ElxA and Mutant Variants
The cloning of the gene encoding ElxA into pET28b is described in Velasquez, J. E. et al. Chem. Biol. 18, 857-867 (2011). The ElxA leader peptide variants were constructed using QuikChange mutagenesis following the method of Liu, H., and Naismith, J. H. BMC biotechnology 8, 91 (2008). The his6-elxA-pET28b was used as a template to introduce the different mutations by PCR using the corresponding primer pair listed in the Table 6. A typical PCR reaction consisted of 1× HF Buffer, 0.2 mM DNTP, 1 μM forward and reverse primers, 10 ng template DNA, 3% (v/v) DMSO and 0.02 U μL−1 Phusion polymerase (NEB). After PCR, reactions were incubated at 37° C. for 2 h with DpnI (5 U). After treatment with DpnI, samples were purified using the QIAquick PCR purification kit (Qiagen). An aliquot of 10 μL was used to transform E. coli DH5ca cells using the heat shock method and cells were plated on LBA plates supplemented with kn (50 μg mL−1). Plates were incubated at 37° C. O/N. A single colony was inoculated in LB supplemented with kan, grown at 37° C. O/N, and plasmid was extracted using the QIAprep spin mini prep kit (Qiagen). The desired mutations were verified by DNA sequencing.
Overexpression and purification of wild type ElxA and mutant variants was carried out using a previously described method with minor modifications (Velasquez, J. E. et al. Chem. Biol. 18, 857-867 (2011)). An aliquot of 50 ng of recombinant DNA from wild type ElxA and each of the mutants described above was used to transform 50 μL of electrocompetent E. coli BL21 (DE3) cells and plated on LBA supplemented with kan (50 μg mL−1) at 37° C. O/N. A single colony was used to inoculate a starter inoculum of LB supplemented with kn and incubated at 37° C. for 12 h. After the incubation period, 6 L of Terrific Broth (TB) supplemented with kn, and glycerol (4 mL L−1), were inoculated with the starter culture to obtain an initial OD600 of 0.025. Flasks were incubated at 37° C., 250 rpm, and peptide expression was induced at 18° C., 250 rpm for 18 h with 1.0 mM IPTG when the OD600 reached 1.0. Cells were harvested by centrifugation (6,976×g, 20 min, 4° C.), resuspended in 30 mL of LanA Buffer 1 (6 M guanidine hydrochloride, 20 mM NaH2PO4 pH 7.5, 500 mM NaCl and 0.5 mM imidazole) and lysed by sonication (35% amplitude, 4.0 s pulse, 9.9 s pause, 15 min). Cell debris was removed by centrifugation (22789×g, 30 min, 4° C.) and supernatant was filtered through a 0.45 μm syringe filter unit. Purification was carried out using IMAC with a 5 mL HisTrap column pre-packed with Ni Sepharose™. After loading the supernatant into the column, the column was washed with 10 column volumes (CV) of LanA Buffer 1 followed by 10 CV of LanA Buffer 2 (4 M guanidine hydrochloride, 20 mM NaH2PO4 pH 7.5, 300 mM NaCl and 30 mM imidazole) to remove any non-specifically bound proteins, followed by peptide elution in 3 CV of LanA Elution Buffer (4 M guanidine hydrochloride, 20 mM Tris HCl pH 7.5, 100 mM NaCl, 1 M imidazole). To remove excess salts, the peptide was purified by reversed phase high performance liquid chromatography (RP-HPLC) using a C4 Waters Delta Pak cartridge column with a linear gradient of 2% (v/v) of solvent A [80% (v/v) MeCN, 20% (v/v) H2O, 0.086% (v/v) trifluoroacetic acid (TFA)] in solvent B [0.1% (v/v) TFA in H2O] to 75% (v/v) solvent A over 30 min. Fractions were analyzed for the desired peptide by MALDI-TOF MS. Fractions containing peptide were freeze dried using a lyophilizer (Labconco) and stored at −20° C. The purity of each peptide was assessed by analytical HPLC.
Cloning, Expression, and Purification of His6-NisA and Mutant Variants
The cloning of the gene encoding NisA into pRSF Duet-I is described in Garg, N. et al. Proc. Natl. Acad. Sci. U.S.A. 110, 7258-7263 (2013). The NisA leader peptide variants were constructed using QuikChange mutagenesis. The his6-nisA-pRSF Duet-I plasmid was used as a template to introduce the different mutations by PCR using the corresponding primer pair listed in Table 6. PCR conditions, overexpression, and purification of peptides were performed as described above.
A typical activity assay consisted of 50 mM Tris HCl pH 8.0, 50 μM peptide, 5 μM MBP-ElxP in a final volume of 100 μL. The sample was incubated for 2 h at room temperature. To monitor cleavage activity, samples were desalted using a zip tip concentrator (Millipore), mixed in a 1:1 ratio with sinapinic acid, and spotted on a MALDI-TOF Bruker plate. Ion intensities for the resulting precursor peptide, leader peptide and core peptide were normalized and the proteolytic efficiency was measure as the amount of substrate left after cleavage reaction (Table 1). Reactions were performed with tagged enzyme and substrates unless otherwise noticed.
The kinetic parameters of MBP-ElxP were determined using an HPLC based assay following the method of Ishii, S. et al. J. Biol. Chem. 281, 4726-4731 (2006). The peptidase activity of tagged ElxP was assayed in a 100 μL reaction mixture containing 50 mM Tris pH 8.0, and various concentrations of wild type His6-ElxA or mutant variants. The reaction was started by adding MBP-ElxP to a final concentration such as to consume less than 10% of the initial substrate concentration in the time frame of the assay. The enzyme concentration ranged from 0.25 μM to 1 μM. The reaction was incubated at room temperature and quenched in 0.1% (v/v) TFA and 5 mM TCEP at different time points. The reactions were loaded on a Hypersil Gold C4(250×4.6 mm, 5μ analytical column (Thermo Fisher Scientific)) connected to an Agilent 1260 Liquid Chromatography (HPLC) system (Agilent Technologies). Product formation was detected by monitoring the increase in the peak area of the leader peptide at 220 nm. The leader peptide was separated from the unmodified core and precursor peptide using a linear gradient from 2% to 75% (v/v) of solvent A [80% (v/v) MeCN, 20% (v/v) H2O, 0.086% (v/v) TFA] in solvent B [0.1% (v/v) TFA in H2O] over 30 min at a flow rate of 1 mL min−1 at room temperature. The concentration of the leader peptide was calculated by converting the area under the leader peptide peak to leader peptide concentration using a calibration curve made from purified leader peptide. Rates of leader peptide formation were then plotted against substrate concentration and the resulting graph was fit to the Michaelis-Menten equation. Values were plotted as the average and standard error of two independent experiments. Reactions were performed with tagged enzymes and substrates unless otherwise noticed.
Peptides were synthesized by standard Fmoc-based solid phase peptide synthesis.
To generate pHis6-ElxO(S139A), pHis6-ElxO(Y152F), pHis6-ElxO(K156A), and pHis6-ElxO(K156M), the entire pHis6-ElxO reported previously was amplified by PCR using Pfu Turbo hot-start DNA polymerase (Stratagene) or iProof high-fidelity polymerase (BioRad) with the appropriate mutagenesis primers ElxO.S139A.FP and ElxO.S139.RP, ElxO.Y152F.FP and ElxO.Y152F.RP, ElxO.K156A.FP and ElxO.K156A.RP, or ElxO.K156M.FP and ElxO.K156M.RP, followed by treatment with DpnI (New England Biolabs) and transformation of Escherichia coli DH5a cells. The correct sequence of the insert was confirmed by sequencing at the W. M. Keck Center for Comparative and Functional Genomics at the University of Illinois at Urbana-Champaign. The proteins were expressed and purified using a HisTrap HP column (GE Healthcare) as described elsewhere for His6-ElxO, followed by further purification by size exclusion chromatography using an ÄKTA purifier equipped with a HiLoad 16/60 Superdex 200 column (GE Healthcare) and a flow of 1.5 mL min−1 of running buffer (50 mM HEPES, 300 mM NaCl, 10% (v/v) glycerol, pH 7.4).
Wild type or mutant His6-ElxO (2 or 10 μM) and purified peptide (0.1 to 5 mM) were incubated with NADPH (2.5 mM) in assay buffer (100 mM HEPES, 500 mM NaCl, pH 7.5) at 25° C. Reaction progress was monitored by UV spectrophotometry to measure initial rates, measuring the disappearance of NADPH absorbance at 340 nm. Formation of reduced peptides was confirmed by LC-MS using an Agilent 1200 instrument equipped with a single quadruple multimode ESI/APCI ion source mass spectrometry detector and a Synergi Fusion-RP column (4.6 mm×150 mm, Phenomenex). The mobile phase was 0.1% (v/v) formic acid in water (A) and methanol (B). A gradient of 0-70% (v/v) B in A over 30 minutes and a flow rate of 0.5 mL min−1 were used.
Synthetic lactocin S (50 μM), obtained from Prof. J. Vederas (University of Alberta), was incubated with His6-ElxO (50 μM) and NADPH (10 mM) in assay buffer (100 mM HEPES, 500 mM NaCl, pH 7.5) at room temperature for 12 h. The formation of reduced peptide was confirmed by LC-MS using a Waters SYNAPT™ mass spectrometry system equipped with a ACQUITY UPLC®, an ESI ion source, a quadrupole time-of-flight detector, and a ACQUITY Bridged Ethyl Hybrid (BEH) C8 column (2.1 mm×50 mm, 1.7 μm, Waters). A gradient of 3-97% (v/v) B (0.10% (v/v) formic acid in methanol) in A (0.1% (v/v) formic acid in water) over 12 min was used.
Agar diffusion bioactivity assays were performed using de Man-Rogosa-Sharpe (MRS) agar media. For each assay, aliquots of agar medium inoculated with overnight cultures of indicator strain (1/100 dilution) were poured into sterile plates. Aliquots of 20 μL of sample were placed into wells made on the solidified agar and the plates were incubated at 37° C. overnight. For determination of critical concentration, the diameter of the inhibition zones were determined and fitted to the equation D=a+b×(C), where D is the diameter of the inhibition zone, C is the concentration of bacteriocin, and a and b are constant parameters. For MIC determinations, serial dilutions of peptides were prepared in MRS broth and aliquots of 50 μL were dissolved in 150 μL of a 1 to 50 dilution of an overnight culture of indicator strain in fresh MRS broth on 96-well plates. The cultures were incubated at 37° C. overnight and the wells with no bacterial growth (OD600<0.3) were determined.
The gene encoding CylA was synthesized by GeneArt (Invitrogen) with codon usage optimized for E. coli expression. The DNA sequences for cylM, cylA, cylLL and cylLS are listed in Table 9. All other oligonucleotides were synthesized by Integrated DNA Technologies and used as received. Restriction endonucleases, DNA polymerases, and T4 DNA ligase were obtained from New England Biolabs. Media components were purchased from Difco Laboratories. Trypsin was purchased from Worthington Biochemical Corporation; Factor Xa was obtained from New England Biolabs and other endoproteinases were ordered from Roche Biosciences. Defibrinated rabbit blood was purchased from Hemostat Laboratories and used within 10 days of receipt. Chemicals were ordered from Sigma Aldrich or Fisher Scientific unless specified otherwise. Miniprep, gel extraction and PCR purification kits were purchased from Qiagen.
All polymerase chain reactions (PCRs) were carried out on a C1000™ thermal cycler (Bio-Rad). DNA sequencing was performed by ACGT, Inc. Preparative HPLC was performed using a Waters Delta 600 instrument equipped with appropriate columns. Solid phase extraction was performed with a Strata X-L polymeric reverse phase column (Phenomenex). MALDI-TOF MS was carried out on a Bruker Daltonics UltrafleXtreme MALDI TOF/TOF instrument (Bruker) or a Voyager-DE-STR instrument (Applied Biosystems). LC-ESI-Q/TOF MS analyses were conducted using a Micromass Q-Tof Ultima instrument (Waters) equipped with a Vydac C18 column (5 μm; 100 Å; 250×1.0 mm). Absorbance of rabbit hemoglobin solution was measured in 96-well plates with a Synergy™ H4 Microplate Reader (BioTek). Negative numbers are used for amino acids in the leader peptide counting backwards from the leader peptide cleavage site.
The indicator strain, Lactococcus lactis HP and the lichenicidin producing strain, Bacillus licheniformis DSM 13=ATCC 14580 were both obtained from American Type Culture Collection. E. coli DH5a and E. coli BL21 (DE3) cells were used as host for cloning and plasmid propagation, and host for protein expression, respectively. The co-expression vector pRSFDuet-1 was obtained from Novagen.
The cylA, cylLL and cylLS genes were synthesized with codon usage optimized for E. coli expression, amplified using appropriate primers and cloned into the MCS1 of a pRSFDuet-1 vector using restriction sites EcoRI and NotI to generate the plasmids pRSFDuet-1/CylA-27-412, pRSFDuet-1/CylLL and pRSFDuet-1/CylLS. Primer sequences are listed in Table 10.
Genes encoding the mutant peptides were amplified by multi-step overlap extension PCR. First, the amplification of the 5′ leader part was carried out by 30 cycles of denaturing (95° C. for 10 s), annealing (55° C. for 30 s), and extending (72° C. for 15 s) using forward primers for halA2 and halA1 and appropriate reverse primers (Table 10) to generate a forward megaprimer (FMP). In parallel, PCR reactions using appropriate forward primers and reverse primers for halA2 and halA1 (Table 10) were performed to produce 3′ fragments (termed reverse megaprimer, RMP). The 5′ FMP fragment and the 3′ RMP fragment were purified by 2% agarose gel followed by use of a Qiagen gel extraction kit. The 2 fragments were combined in equimolar amounts (approximately 20 ng each for a 50 μL PCR) and amplified using the same PCR conditions as above with halA2 and halA1 primers. The resulting PCR products were purified, digested and then cloned into the MCS1 of pRSFDuet-1/HalM2-2 and pRSFDuet-1/HalM1-2, respectively, to generate pRSFDuet-1/HalA2-GDVQAE/HalM2-2, pRSFDuet-1/HalA2-GDVQAE-T2A/HalM2-2 and pRSFDuet-1/HalA1-GDVQAE/HalM1-2 vectors.
Mutant peptide genes were generated by a similar multi-step overlap extension PCR procedure as described above and cloned in the MCS1 of a pRSFDuet vector to generate pRSFDuet-1/ProcA1.7-GDVQAE and pRSFDuet-1/NisA-GDVQAE vectors. Primer sequences are listed in Table 10.
The expression vectors pRSFDuet-1/ProcA1.7-GDVQAE-T1G, pRSFDuet-1/ProcA1.7-GDVQAE-T1F, pRSFDuet-1/ProcA1.7-GDVQAE-T1W, pRSFDuet-1/CylA-27-412-E95A and pRSFDuet-1/CylA-27-412-S359A were generated using quick change methodology based on the pRSFDuet-1/ProcA1.7-GDVQAE and pRSFDuet-1/CylA-27-412 vectors, respectively. Primer sequences are listed in Table 10.
Bacillus licheniformis DSM 13=ATCC 14580 was grown in LB at 37° C. for 12 h with vigorously shaking and plasmid was extracted using a Qiagen miniprep kit. LicA2 and LicP genes were amplified from the plasmid using appropriate primers and cloned into the MCS1 of a pRSFDuet-1 vector using restriction sites BamHI and NotI to generate pRSFDuet-1/LicA2 and pRSFDuet-1/LicP-25-433, respectively. Primer sequences are listed in Table 10.
E. coli BL21 (DE3) cells were transformed with pRSFDuet-1/CylA-27-412, pRSFDuet-1/CylA-27-412-E95A, pRSFDuet-1/CylA-27-412-S359A or pRSFDuet-1/LicP-25-433 vectors and plated on an LB plate containing 50 mg/L kanamycin. A single colony was picked and grown in 20 mL of LB with kanamycin at 37° C. for 12 h and the resulting culture was inoculated into 2 L of LB. Cells were cultured at 37° C. until the OD at 600 nm reached 0.5, cooled and IPTG was added to a final concentration of 0.1 mM. The cells were cultured at 18° C. for another 10 h before harvesting. The cell pellet was resuspended on ice in LanP start buffer (20 mM HEPES, 1 M NaCl, pH 7.5 at 25° C.) and lysed by homogenization. The lysed sample was centrifuged at 23,700×g for 30 min and the pellet was discarded. The supernatant was passed through 0.45-μm syringe filters and the protein was purified by immobilized metal affinity chromatography (IMAC) loaded with nickel as described in B. Li et al. Methods Enzymol. 2009, 458, 533. The proteins were generally eluted from the column at an imidazole concentration between 150 mM and 300 mM and the buffer was exchanged using a GE PD-10 desalting column pre-equilibrated with LanP start buffer. Protein concentration was quantified by its absorbance at 280 nm. The extinction coefficient for His6-CylA-27-412, His6-CylA-27-412-E95A and His6-CylA-27-412-S359A was calculated as 30,830 M−1 cm−1. The extinction coefficient for His6-LicP-25-433 was calculated as 46,300 M−1 cm−1. Aliquoted protein solutions were flash-frozen and kept at −80° C. until further usage.
Modified peptides were obtained using a similar procedure described previously using the corresponding co-expression vectors (Y. Shi, et al. J. Am. Chem. Soc. 2011, 133, 2338; W. Tang and W. A. van der Donk, Nat. Chem. Biol. 2013, 9, 157).
E. coli BL21 (DE3) cells were transformed with pRSFDuet-1/CylLL, pRSFDuet-1/CylLS, pRSFDuet-1/ProcA1.7-GDVQAE, pRSFDuet-1/NisA-GDVQAE, pRSFDuet-1/ProcA1.7-GDVQAE-T1G, pRSFDuet-1/ProcA1.7-GDVQAE-T1F, pRSFDuet-1/ProcA1.7-GDVQAE-T1W or pRSFDuet-1/LicA2 plasmids and plated on an LB plate containing 50 mg/L kanamycin. A single colony was picked and grown in 10 mL of LB with kanamycin at 37° C. for 12 h and the resulting culture was inoculated into 1 L of LB. Cells were cultured at 37° C. until the OD at 600 nm reached 0.5 and IPTG was added to a final concentration of 0.2 mM. The cells continued to be cultured at 37° C. for another 3 h before harvesting. The cell pellet was resuspended at room temperature in LanA start buffer (20 mM NaH2PO4, pH 7.5 at 25° C., 500 mM NaCl, 0.5 mM imidazole, 20% glycerol) and lysed by sonication. The sample was centrifuged at 23,700×g for 30 min and the supernatant was discarded. The pellet was then resuspended in LanA buffer 1 (6 M guanidine hydrochloride, 20 mM NaH2PO4, pH 7.5 at 25° C., 500 mM NaCl, 0.5 mM imidazole) and sonicated again. The insoluble portion was removed by centrifugation at 23,700×g for 30 min and the soluble portion was passed through 0.45-μm syringe filters. His6-tagged peptides were purified by immobilized metal affinity chromatography (IMAC) loaded with nickel as described in B. Li et al. Methods Enzymol. 2009, 458, 533.
The eluted fractions were desalted using reverse phase HPLC equipped with a Waters Delta-pak C4 column (15 μm; 300 Å; 25×100 mm) or a Strata XL polymeric reverse phase SPE column. The desalted peptides were lyophilized and stored at −20° C. for future use.
Targeted peptides were dissolved in H2O to a final concentration of 3 mg/mL. To a 85 μL solution of peptides, 10 μL of 500 mM HEPES buffer (pH 7.5) was added followed by 5 μL of 0.5 mg/mL AspN protease (for modified CylLL and CylLS peptides), 0.1 mg/mL CylA protease (for modified and unmodified CylLL, CylLS peptides, and modified HalA2-GDVQAE peptide) or 0.1 mg/mL LicP protease (for LicA2). For cleavage tests of the engineered GDVQAE peptides, CylA was added to a final concentration of 0.01 mg/mL, whereas the peptide was added to a concentration of 0.3 mg/mL with 50 mM HEPES buffer (pH 7.5). The protease cleavage reaction mixtures were kept at 25° C. for 1 to 48 h. Osmotic pressure was adjusted with NaCl solution with a final concentration of 150 mM. The digested peptide mixture was directly used for antimicrobial and hemolytic assay. For the kinetic analysis of CylA and its mutant proteins, His6-CylA-27-412 and His6-CylA-27-412-E95A proteins were added to a final concentration of 500 ng/mL and 2.5 μg/mL (22 nM and 110 nM), respectively, with modified CylLS supplied at a concentration of 0.3 mg/mL (36 μM). The reactions were allowed to proceed at room temperature and were stopped by 1% TFA at different time points for LC/MS analysis. Halα and Halβ were obtained by factor Xa cleavage of modified HalA1Xa and HalA2 Xa peptides using the procedure of Y. Shi, et al. J. Am. Chem. Soc. 2011, 133, 2338. CylLL″ and CylLS″ were prepared in the same way using modified CylLL-E-1K and CylLS-E-1K peptides as described in W. Tang and W. A. van der Donk, Nat. Chem. Biol. 2013, 9, 157.
As full length CylA is not available due to its self-cleavage, His6-CylA-27-412-E95A was chosen to serve as a substituent of full length CylA as the self-cleavage was abolished whereas the conserved catalytic C-terminal region of CylA remained unchanged. To obtain the mature protease CylA-96-412, His6-CylA-27-412 was aged at 4° C. for 12 hours to allow the self-cleavage to proceed until CylA-96-412 was the dominant peak monitored by MALDI-TOF MS. The aged protein mixture was directly used as a substituent of CylA-96-412. To test the proteases' activities, CylA-96-412 and His6-CylA-27-412-E95A were supplied with a final concentration of 22 nM and 110 nM, respectively, with modified CylLS served at a concentration of 36 μM. The reactions were stopped at 3 minute, 6 minute, 12 minute and 24 minute by 1% TFA and the formation of mature CylLS″ was monitored by liquid chromatography MS (LC/MS). A 5 μL volume of sample obtained from the cleavage reaction was applied to the column that was pre-equilibrated in aqueous solvent A. The solvents used for LC were: solvent A=0.1% formic acid in 95% water/5% acetonitrile and solvent B=0.1% formic in 95% acetonitrile/5% water. A solvent gradient of 0%-80% B over 30 min was employed and the fractionated sample was directly subjected to ESI-Q/TOF MS analysis. The production of core peptide was analyzed by extracted ion chromatography monitoring the desired product mass 1017 (M+2H+).
L. lactis HP cells were grown in GM17 media under anaerobic conditions at 25° C. for 16 h. Agar plates were prepared by combining 15 mL of molten GM17 agar (cooled to 42° C.) with 150 μL of dense cell culture. The seeded agar was poured into a sterile 100 mm round dish (VWR) to solidify. Peptide samples were directly spotted on the solidified agar. Plates were incubated at 30° C. for 16 h and the antimicrobial activity was determined by the size of the zone of growth inhibition.
A sample of 1 mL of defibrinated rabbit blood was added into 20 mL of PBS in a 50 mL conical tube and mixed gently. The PBS-diluted blood sample was centrifuged at 800×g for 5 min at 4° C. and the supernatant containing lysed blood cells and released hemoglobin was discarded. The process was repeated 2 to 4 times until the supernatant was clear. The blood cells were then diluted with PBS to make a 5% solution, which was immediately used to test the hemolytic activity of the peptides. To an Eppendorf tube, 50 μL of 5% washed red blood cell sample was added followed by the addition of the desired peptide samples or controls. PBS was used to adjust the final volume to 85 μL. All tubes were kept in a 37° C. incubator to allow the lytic reaction to proceed. At each time point, 8 or 10 μL of reaction mixture was taken out, diluted with 190 μL of fresh PBS and centrifuged at 800×g for 5 min. The supernatant (170 μL) was transferred to a new well and the absorbance was measured at 415 nm. The absorbance of prepared blood sample at each time point was analyzed in triplicate and the maximum absorbance was determined by adding 35 μL of 0.1% Triton in PBS to 50 μL of 5% blood sample and using the same analysis procedure.
General methods. All polymerase chain reactions (PCRs) were carried out on a C1000 thermal cycler (Bio-Rad). DNA sequencing was performed by ACGT, Inc. Preparative HPLC was performed using a Waters Delta 600 instrument equipped with a Waters Delta-pak C4 column (15 μm 300 Å25×100 mm). Solid phase extraction was performed with a Strata-X polymeric reversed phase column (Phenomenex) or Vydac BioSelect C4 reversed phase column. FPLC was carried out using an AKTA FPLC system (Amersham Pharmacia Biosystems). MALDI-TOF MS was carried out on a Bruker Daltonics UltrafleXtreme MALDI TOF/TOF instrument (Bruker). The detection of peptides with low molecular weights (700-3,500 Da), peptides with medium molecular weights (3,500-20,000 Da) and proteins with high molecular weights (20,000-50,000 Da) was achieved by using different instrument settings optimized for these mass ranges.
Materials. All oligonucleotides were synthesized by Integrated DNA Technologies and used as received. Restriction endonucleases, DNA polymerases, and T4 DNA ligase were obtained from New England Biolabs. Media components were purchased from Difco Laboratories and Fisher Scientific. Chemicals were ordered from Sigma Aldrich or Fisher Scientific unless otherwise specified. Miniprep, gel extraction and PCR purification kits were purchased from Qiagen and 5 PRIME. An UltraClean microbial DNA isolation kit was obtained from Mo Bio Laboratories, Inc.
Strains and plasmids. The lichenicidin producing strain, Bacillus licheniformis ATCC 14580, was obtained from the American Type Culture Collection. E. coli DH5U and E. coli BL21 (DE3) cells were used as hosts for cloning and plasmid propagation, and hosts for protein expression, respectively. The expression vector pRSFDuet-1 was obtained from Novagen.
Extraction of genomic DNA from Bacillus licheniformis ATCC 14580. Bacillus licheniformis ATCC 14580 was cultured in LB medium at 37° C. aerobically for 12 h and the genomic DNA was extracted using an UltraClean microbial DNA isolation kit following the manufacturer's protocol.
Construction of pRSFDuet-1 derivatives for expression of LicP-25-433 and LicA2. licP and licA2 genes were amplified from the genomic DNA of Bacillus licheniformis ATCC 14580 using appropriate primers and cloned into the multiple cloning site 1 (MCS1) of a pRSFDuet-1 vector to generate pRSFDuet-1/LicP-25-433 and pRSFDuet-1/LicA2 plasmids, respectively. Primer sequences are listed in Table 11.
Construction of pRSFDuet-1 derivatives for expression of ProcA1.7-NDVNPE and NisA-NDVNPE. Engineered peptide genes were generated by multi-step overlap extension PCR. First, the amplification of the 5′ leader part was carried out by 30 cycles of denaturing (95° C. for 10 s), annealing (55° C. for 30 s), and extending (72° C. for 15 s) using forward primers for procA1.7 and nisA and appropriate leader peptide reverse primers containing the mutations (Table 11) to generate a forward megaprimer (FMP). In parallel, PCR reactions using forward primers and reverse primers for procA1.7 and nisA core peptides (Table 11) were performed to produce the 3′ core fragments (termed reverse megaprimer, RMP). The 5′ FMP fragment and 3′ RMP fragment were purified by 2% agarose gel, combined in equimolar amounts and amplified using the same PCR conditions as above with procA1.7 and nisA primers. The resulting PCR products were purified, digested and then cloned into the MCS1 of a pRSFDuet-1 vector to generate pRSFDuet-1/ProcA1.7-NDVNPE and pRSFDuet-1/NisA-NDVNPE plasmids.
Construction of pRSFDuet-1 derivatives for expression of LicP-25-433-S376A, LicP-25-433-H186A, LicP-25-433-E100A, LicP-25-433-E100A-E102A, G-LicA2, NisA-NDVNPE-I1G, NisA-NDVNPE-I1T, NisA-NDVNPE-I1C, NisA-NDVNPE-I1L, NisA-NDVNPE-I1F, NisA-NDVNPE-I1W, NisA-NDVNPE-I1K, NisA-NDVNPE-I1E, LicA2-E-1A, LicA2-E-1D, LicA2-E-1Q, LicA2-P-2A, LicA2-N-3A, LicA2-V-4A, LicA2-V-4L, LicA2-V-4F, LicA2-D-5A, and LicA2-D-5K. The expression plasmids pRSFDuet-1/LicP-25-433-S376A, pRSFDuet-1/LicP-25-433-H186A, pRSFDuet-1/LicP-25-433-E100A, pRSFDuet-1/LicP-25-433-E100A-E102A, pRSFDuet-1/G-LicA2, pRSFDuet-1/NisA-NDVNPE-I1G, pRSFDuet-1/NisA-NDVNPE-I1T, pRSFDuet-1/NisA-NDVNPE-I1C, pRSFDuet-1/NisA-NDVNPE-I1L, pRSFDuet-1/NisA-NDVNPE-I1F, pRSFDuet-1/NisA-NDVNPE-I1W, pRSFDuet-1/NisA-NDVNPE-I1K, pRSFDuet-1/NisA-NDVNPE-I1E, pRSFDuet-1/LicA2-E-1A, pRSFDuet-1/LicA2-E-1D, pRSFDuet-1/LicA2-E-1Q, pRSFDuet-1/LicA2-P-2A, pRSFDuet-1/LicA2-N-3A, pRSFDuet-1/LicA2-V-4A, pRSFDuet-1/LicA2-V-4L, pRSFDuet-1/LicA2-V-4F, pRSFDuet-1/LicA2-D-5A, and pRSFDuet-1/LicA2-D-5K were generated using QuikChange methodology based on pRSFDuet-1/LicP-25-433, pRSFDuet-1/LicA2 and pRSFDuet-1/NisA-NDVNPE as templates. Primer sequences are listed in Table 11.
Construction of pRSFDuet-1 derivatives for co-expression of LicM2 with LicA2. LicM2 was amplified from the genomic DNA of Bacillus licheniformis ATCC 14580 using appropriate primers and cloned into the MCS2 of a pRSFDuet-1 vector to generate pRSFDuet-1/LicM2-2. The expression plasmid pRSFDuet-1/LicA2/LicM2-2 was constructed by inserting the licA2 gene into the MCS1 of the pRSFDuet-1/LicM2-2 plasmid. Primer sequences are listed in Table 11.
Construction of pET28b-MBP-BamL plasmid with LicP recognition sequence. Oligonucleotides corresponding to the LicP recognition sequence NDVNPE/SGS (SEQ ID NO.: 37) were inserted into the pET28b-MBP-BamL plasmid (1) in front of the DNA sequences corresponding to the TEV cleavage site using QuikChange methodology. Primer sequences are listed in Table 11. The resultant plasmid expressed the recombinant fusion protein MBP-BamL with the LicP recognition sequence installed between the two polypeptide portions for MBP and BamL (SEQ ID NO.: 221).
Expression and purification of LicP and LicP mutant proteins. E. coli BL21 (DE3) cells were transformed with one of the following plasmids: pRSFDuet-1/LicP-25-433, pRSFDuet-1/LicP-25-433-S376A, pRSFDuet-1/LicP-25-433-H186A, pRSFDuet-1/LicP-25-433-E100A or pRSFDuet-1/LicP-25-433-E100A-E102A, and plated on an LB plate containing 50 mg/L kanamycin. A single colony was picked and grown in 20 mL of LB containing 50 mg/L kanamycin at 37° C. for 12 h and the resulting culture was inoculated into 2 L of LB containing 50 mg/L kanamycin. Cells were cultured at 37° C. until the OD at 600 nm reached 0.5, cooled and IPTG was added to a final concentration of 0.1 mM. The cells were cultured at 18° C. for another 10 h before harvesting. The cell pellet was resuspended on ice in LanP buffer (20 mM HEPES, 1 M NaCl, pH 7.5 at 25° C.) and lysed by homogenization. The lysed sample was centrifuged at 23,700×g for 30 min and the pellet was discarded. The supernatant was passed through 0.45-μm syringe filters and the protein was purified by immobilized metal affinity chromatography (IMAC) loaded with nickel. The proteins were generally eluted from the column at an imidazole concentration between 150 mM and 300 mM and the buffer was exchanged using a GE PD-10 desalting column or a gel-filtration column pre-equilibrated with LanP buffer. Protein concentration was quantified by the absorbance at 280 nm. The extinction coefficient for His6-LicP-25-433 was calculated as 46,300 M−1 cm−1. His6-LicP-25-433-S376A was predominantly expressed in inclusion bodies.
Soluble protein was obtained by combining fractions eluted from the nickel column containing the desired protein and concentrating to a small volume. No gel filtration was performed for the mutant protein. The yield was determined to be about 50 μg for 1 L of culture. Aliquoted protein solutions were flash-frozen and kept at −80° C. until further usage.
Expression and purification of modified His6-LicA2. Modified LicA2 was obtained using a procedure similar to that reported previously using the corresponding co-expression plasmid pRSFDuet-1/LicA2/LicM2-2.
Expression and purification of unmodified His6-LicA2, His6-G-LicA2, His6-ProcA1.7-NDVNPE, His6-NisA-NDVNPE, His6-NisA-NDVNPE-I1G, His6-NisA-NDVNPE-I1T, His6-NisA-NDVNPE-I1C, His6-NisA-NDVNPE-I1L, His6-NisA-NDVNPE-I1F, His6-NisA-NDVNPE-I1W, His6-NisA-NDVNPE-I1K and His6-NisA-NDVNPE-I1E. E. coli BL21 (DE3) cells were transformed with one of the following plasmids: pRSFDuet-1/LicA2, pRSFDuet-1/G-LicA2, pRSFDuet-1/ProcA1.7-NDVNPE, pRSFDuet-1/NisA-NDVNPE, pRSFDuet-1/NisA-NDVNPE-I1G, pRSFDuet-1/NisA-NDVNPE-I1T, pRSFDuet-1/NisA-NDVNPE-I1C, pRSFDuet-1/NisA-NDVNPE-I1L, pRSFDuet-1/NisA-NDVNPE-I1F, pRSFDuet-1/NisA-NDVNPE-I1W, pRSFDuet-1/NisA-NDVNPE-I1K or pRSFDuet-1/NisA-NDVNPE-I1E. Then the cells were plated on an LB plate containing 50 mg/L kanamycin. A single colony was picked and grown in 10 mL of LB containing 50 mg/L kanamycin at 37° C. for 12 h and the resulting culture was inoculated into 1 L of LB containing 50 mg/L kanamycin. Cells were cultured at 37° C. until the OD at 600 nm reached 0.5 and IPTG was added to a final concentration of 0.2 mM. The cells continued to be cultured at 37° C. for another 3 h before harvesting. The cell pellet was resuspended at room temperature in LanA start buffer (20 mM NaH2PO4, pH 7.5 at 25° C., 500 mM NaCl, 0.5 mM imidazole, 20% glycerol) and lysed by sonication. The sample was centrifuged at 23,700×g for 30 min and the supernatant was discarded. The pellet was then resuspended in LanA buffer 1 (6 M guanidine hydrochloride, 20 mM NaH2PO4, pH 7.5 at 25° C., 500 mM NaCl, 0.5 mM imidazole) and sonicated again. The insoluble portion was removed by centrifugation at 23,700×g for 30 min and the soluble portion was passed through 0.45-μm syringe filters. His6-tagged peptides were purified by IMAC as previously described. The eluted fractions were desalted using reversed phase HPLC or a Strata X polymeric reversed phase SPE column. The desalted peptides were lyophilized and stored at −20° C. for future use.
Expression and purification of unmodified His6-LicA2-E-1A, His6-LicA2-E-1D, His6-LicA2-E-1Q, His6-LicA2-P-2A, His6-LicA2-N-3A, His6-LicA2-V-4A, His6-LicA2-V-4L, His6-LicA2-V-4F, His6-LicA2-D-5A, and His6-LicA2-D-5K. E. coli BL21 (DE3) cells were transformed with one of the following plasmids: pRSFDuet-1/LicA2-E-1A, pRSFDuet-1/LicA2-E-1D, pRSFDuet-1/LicA2-E-1Q, pRSFDuet-1/LicA2-P-2A, pRSFDuet-1/LicA2-N-3A, pRSFDuet-1/LicA2-V-4A, pRSFDuet-1/LicA2-V-4L, pRSFDuet-1/LicA2-V-4F, pRSFDuet-1/LicA2-D-5A, or pRSFDuet-1/LicA2-D-5K. Then the cells were plated on an LB plate containing 50 mg/L kanamycin. A single colony was picked and grown in 7 or 20 mL of LB containing 50 mg/L kanamycin at 37° C. for 14.5-16.5 h and the resulting culture was used to inoculate 750 mL of LB containing 50 mg/L kanamycin. Cells were cultured at 37° C. until the OD at 600 nm reached 0.5-0.6 and IPTG was added to a final concentration of 0.2 mM. The cells continued to be cultured at 37° C. for another 3 h before harvesting. The cell pellet was resuspended in LanA start buffer and lysed by sonication. The sample was centrifuged at 15,377×g for 30 min and the supernatant was discarded. The pellet was then resuspended in LanA buffer 1 and sonicated again. The insoluble portion was removed by centrifugation at 15,377×g for 30 min and the soluble portion was passed through 0.45-μm syringe filters. His6-tagged peptides were purified by IMAC as previously described (2). Eluted fractions were desalted using a Vydac Bioselect C4 reversed phase SPE column. The desalted peptides were lyophilized, dissolved in water to a final concentration of 3 mg/mL and stored at −20° C. for future use.
Intermolecular cleavage of His6-LicP-25-433-S376A by His6-LicP-25-433. His6-LicP-25-433-S376A and His6-LicP-25-433 proteins were both diluted with LanP buffer to a final concentration of 0.2 mg/mL. Parallel reactions were set up for His6-LicP-25-433 with a final protein concentration of 0.1 mg/mL in LanP buffer, His6-LicP-25-433-S376A with a final protein concentration of 0.1 mg/mL in LanP buffer, and His6-LicP-25-433-S376A and His6-LicP-25-433 combined with a final protein concentration of 0.1 mg/mL each. The three reactions were allowed to proceed at room temperature for 0, 2, 4, 7 and 19 h before being stopped by addition of SDS loading buffer and boiling at 95° C. for 10 min and analyzed by SDS-PAGE.
Sequential proteolytic cleavage of modified LicA2. HPLC-purified LicM2-modified LicA2 was dissolved in H2O to a final concentration of 3 mg/mL (340 μM). To a 17 μL solution of peptide (final peptide concentration 290 μM), 2 μL of 500 mM HEPES buffer (pH 7.5) was added followed by 1 μL of 0.5 mg/mL AspN. The reaction mixture was kept at room temperature for 12 h, and then 0.5 μL of 0.1 mg/mL LicP (final protein concentration 50 nM) was added. The reaction was then incubated at room temperature for one more hour. MALDI-TOF MS analysis was performed after each step.
Competition assay of LicP activity with modified and linear LicA2. To a reaction vessel containing 70 μL deionized H2O, 5 μL each of 3 mg/mL modified LicA2 and linear G-LicA2 peptides were added (final peptide concentration 17 μM each) followed by 10 μL of 500 mM HEPES buffer (pH 7.5). Then, 10 μL of 0.01 mg/mL LicP was supplied (final protein concentration 21 nM) and the reaction was incubated at room temperature before being quenched by addition of formic acid to a final concentration of 1% at different time points. To observe the complete consumption of both peptides, substrates were incubated as above except that 10 μL of 1 mg/mL LicP was added (final protein concentration 2.1 μM). The reaction mixture was kept at room temperature for 12 h before being quenched with 1% formic acid for MS analysis.
Comparison of the proteolytic activity of LicP and TEV on MBP-BamL. A sample of 1 mL of MBP-BamL ((SEQ ID NO.: 221); 50 μM) was incubated with the same molar amount of LicP or TEV (final concentration 0.54 μM) at 4° C. At different time points, the reaction was quenched by adding an equal volume of loading dye and heating for 10 min at 90° C. The results were analyzed by Coomassie-stained SDS-PAGE.
The size difference between MBP and BamL bands is due to different recognition site locations of LicP and TEV in the construct.
LicP assay and Gel Analysis for wild type His6-LicA2 and His6-LicA2 mutants. A sample containing 100 μM peptide was incubated with 0.4 μM His6-LicP(25-433) and 2 mM DTT in 50 mM HEPES (pH 7.5) buffer with a total reaction volume of 300 μL. After 15 min, 30 min, 1 h, 2 h, 4 h, and 7.5 h, the reactions were centrifuged for 30 s at 2000×g (because of observed precipitation) then 40 μL aliquots were removed and quenched by addition of 10.4 μL 5% aqueous formic acid to a final concentration of 1%.
Formic-acid quenched samples were diluted 25% with 95:5 NuPAGE LDS sample buffer (4×):β-mercaptoethanol to a final concentration of 60 μM peptide and 0.24 μM LicP. Solutions of 60 μM LicA2 substrates, 0.24 μM LicP, and 40-fold diluted Polypeptide Standards (#161-0326) were prepared, each containing 25% 95:5 NuPAGE LDS sample buffer (4×):β-mercaptoethanol. All SDS-PAGE samples were heated for 10 min at 70° C. then a 10-20% Mini-Protean Tris-Tricine gel (#456-3116) loaded with 5 μL per lane was run at 100 V for 125 min in 100 mM Tris, 100 mM Tricine, 0.1% SDS buffer while cooling the entire apparatus in ice.
The gels were subjected to consecutive coomassie and silver staining as described herein. The gels were rocked for 1 h in 50% MeOH/7% AcOH followed by rocking for 45 min in coomassie stain (0.25% coomassie/50% MeOH/10% AcOH), rinsing with H2O, rocking overnight in 20% MeOH/10% AcOH, and then rinsing with H2O again. All of the following steps were conducted with agitation on a Barnstead LabLine multipurpose rotator. The gels were washed with 50% MeOH/7% AcOH for 1.5 h, followed by washing with H2O (3×10 min). The gels were then sensitized 1 min in 0.02% Na2S2O3, then washed with H2O (2×1 min), followed by staining for 30 min in a solution containing 44 mL of H2O, 5.9 mL of 0.1 M AgNO3, and 37.5 μL of 37% formaldehyde. The gels were then washed with H2O (<1 min) then developed in a solution containing 100 mL of 3% Na2CO3, 2 mL of 0.02% Na2S2O3, and 50 μL of 37% formaldehyde for <5 min. Developing was quenched by washing with 12% AcOH for 30 min, followed by washing with H2O (2×30 min). Images of gels were acquired using an HP scanjet 8250.
Identification of the cleavage sites of the LanP proteases encoded in the genome of B. licheniformis 9945A and Bacillus cereus VD045. LanP proteases were purified using a similar procedure as described for LicP. Dehydrated and cyclized LanA2 encoded in the genome of B. licheniformis 9945A was obtained by coexpressing the precursor peptide with its corresponding LanM synthetase in E. coli, whereas linear LanA3 encoded in the genome of Bacillus cereus VD045 was obtained by expression in E. coli. The precursor peptides were incubated with their corresponding proteases and the results were analyzed using MALDI-TOF MS.
Removal of leader peptides of modified or linear LicA2. Modified or linear LicA2 peptides were dissolved in H2O to make a 3 mg/mL solution (340 μM). To a 17 μL solution of peptide (final peptide concentration 290 μM), 2 μL of 500 mM HEPES buffer (pH 7.5) was added followed by 1 μL of 1 mg/mL LicP (final protein concentration 1.1 μM). The reaction was incubated at room temperature for 6 h followed by MS analysis.
Proteolytic cleavage of the leader peptides of engineered peptides. ProcA1.7-NDVNPE was dissolved in H2O to a final concentration of 3 mg/mL (250 μM), whereas for NisA-NDVNPE and its mutant peptides, a 10 mg/mL peptide solution was made (1.3 mM). For ProcA1.7-NDVNPE, 15 μL of peptide solution (final peptide concentration 190 μM) was pre-mixed with 1 μL of 50 mM DTT and 2 μL of 500 mM HEPES buffer (pH 7.5), to which 2 μL of 0.1 mg/mL LicP (final protein concentration 210 nM) was added. The reaction was incubated at room temperature for 4 h before analysis. For NisA-NDVNPE-I1T and NisA-NDVNPE-I1C, 1 μL of peptide (final peptide concentration 65 μM) was pre-mixed with 1 μL of 50 mM DTT and 2 μL of 500 mM HEPES buffer (pH 7.5) in 14 μL H2O, then 2 μL of 0.1 mg/mL LicP (final protein concentration 210 nM) was added. The reaction was incubated at room temperature for 20 h before analysis. For NisA-NDVNPE and other NisA mutant peptides, 1 mg/mL LicP (final protein concentration 2.1 μM) was employed instead of 0.1 mg/mL LicP and the reaction was kept at room temperature for 30 h before analysis.
Materials. All oligonucleotides were synthesized by Integrated DNA Technologies and used as received. Restriction endonucleases, DNA polymerases, and T4 DNA ligase were obtained from New England Biolabs. Media components were purchased from Difco Laboratories and Fisher Scientific. Chemicals were ordered from Sigma Aldrich or Fisher Scientific unless otherwise specified. Miniprep, gel extraction and PCR purification kits were purchased from Qiagen and 5 PRIME. Synthetic genes were obtained from IDT, Inc. For LanP from Bacillus cereus VD156, the DNA was ordered in two gBlocks, whereas for the substrate it was ordered as one oligonucleotide. An UltraClean microbial DNA isolation kit was obtained from Mo Bio Laboratories, Inc.
Strains and plasmids. Bacillus licheniformis ATCC 14580 and Bacillus licheniformis ATCC 9945A were obtained from American Type Culture Collection. E. coli DH5a and E. coli BL21 (DE3) cells were used as hosts for cloning and plasmid propagation, and hosts for protein expression, respectively. The expression vector pRSFDuet-1 was obtained from Novagen.
Extraction of genomic DNA from B. licheniformis ATCC 14580 and B. licheniformis ATCC 9945A. Bacteria were cultured in LB medium at 37° C. aerobically for 12 h and the genomic DNA was extracted using an UltraClean microbial DNA isolation kit following the manufacturer's protocol.
Construction of pRSFDuet-1 derivatives for co-expression of LanM2-9945A with LanA2-9945A, and for expression of LanP-42-476-9945A. The genes for the LanM2 and LanA2 encoded in the genome of B. licheniformis ATCC 9945A (hereafter LanM2-9945A and LanA2-9945A, respectively) were amplified from the genomic DNA using appropriate primers and cloned into a pRSFDuet-1 vector to generate pRSFDuet-1/LanA2-9945A/LanM2-9945A-2 using Gibson assembly (LanA2 in MCS1 and LanM2 in MCS2). The gene encoding residues 42-476 of the class II LanP (designated LanP-42-476-9945A) was amplified from the genomic DNA of B. licheniformis ATCC 9945A using appropriate primers and cloned into the MCS1 of a pRSFDuet-1 vector to generate pRSFDuet-1/LanP-42-476-9945A using Gibson assembly. Primer sequences are listed in Table 11.
Expression plasmids for LanP and one LanA substrate from B. cereus VD156. The lanP gene and one of the genes for its putative substrates were synthesized as codon-optimized dsDNA oligos and cloned into pRSFDuet vector between BamHI and NotI restriction sites via Gibson Assembly. For expression in E. coli, the N-terminal secretion signal of the protease (the first 27 amino acid as predicted by signalP, see sequence shown in red in Table S2) was removed, and E. coli B121(DE3) cells were transformed with the resulting plasmids containing the truncated protease gene (VD156P(del)) or the four substrate genes. The cells were incubated at 37° C. until the OD600 reached 0.5-0.8, then induced with 0.20 mM IPTG, and expressed at 18° C. for 20 h. The synthetic gene sequences are listed in Table 12.
Identification of the cleavage sites of the LanP proteases encoded in the genome of B. licheniformis 9945A and B. cereus VD156. The LanP proteases from B. licheniformis 9945A and B. cereus VD156 were purified using a similar procedure as described for LicP. A His6-tagged analog of dehydrated and cyclized LanA2 encoded in the genome of B. licheniformis 9945A was obtained by coexpression of the precursor peptide with its corresponding LanM synthetase in E. coli using the same procedure as described herein for LicA/LicM. Linear LanA3 encoded in the genome of B. cereus VD156 was also obtained by expression in E. coli as an N-terminal His6-tagged peptide. The peptides were incubated with their corresponding purified proteases and the results were analyzed using MALDI-TOF MS.
aThe sequence shown in red is predicted to be a secretion signal and was removed in the expression constructs. Also shown is the sequence of the LanA3 precursor peptide. The observed cleavage occurred after the Arg shown in blue.
Additional nucleic acid and amino acid sequences described herein are listed in Table 13.
licheniformis
cereus Q1 (CerP)
cereus FRI-35
tusciae DSM 2912
Enterococcus
caccae ATCC
cereus VPC1401
bombysepticus
thuringiensis DB27
Planomicrobium
glaciei CHR43
cereus VD045
licheniformis
All references, including publications, patent applications, and patents, cited herein are hereby incorporated by reference to the same extent as if each reference were individually and specifically indicated to be incorporated by reference and were set forth in its entirety herein. Also incorporated by reference in their entirety are any polynucleotide and polypeptide sequences which reference an accession number correlating to an entry in a public database, such as those maintained by The Institute for Genomic Research (TIGR) on the world wide web at tigr.org and/or the National Center for Biotechnology Information (NCBI) on the world wide web at ncbi.nlm.nih.gov.
All methods described herein can be performed in any suitable order unless otherwise indicated herein or otherwise clearly contradicted by context. The use of any and all examples, or exemplary language (e.g., “such as”) provided herein, is intended merely to better illuminate the invention and does not pose a limitation on the scope of the invention unless otherwise claimed. No language in the specification should be construed as indicating any non-claimed element as essential to the practice of the invention.
Preferred aspects of this invention are described herein, including the best mode known to the inventors for carrying out the invention. Variations of those preferred aspects may become apparent to those of ordinary skill in the art upon reading the foregoing description. The inventors expect a person having ordinary skill in the art to employ such variations as appropriate, and the inventors intend for the invention to be practiced otherwise than as specifically described herein. Accordingly, this invention includes all modifications and equivalents of the subject matter recited in the claims appended hereto as permitted by applicable law. Moreover, any combination of the above-described elements in all possible variations thereof is encompassed by the invention unless otherwise indicated herein or otherwise clearly contradicted by context.
The present application is a continuation of U.S. patent application Ser. No. 15/311,188, filed Mar. 3, 2017, which is the U.S. National Stage Application of PCT/US15/30437, filed May 12, 2015, which claims benefit of priority under 35 U.S.C. 119 to U.S. provisional patent application Ser. No. 61/992,193, filed May 12, 2014, and entitled “HIGHER PERFORMANCE PROTEASES FOR SCARLESS TAG REMOVAL,” the contents of which are herein incorporated by reference in its entirety.
This invention was made with government support under R01GM58822 and 5T32GM070421 awarded by the National Institutes of Health. The government has certain rights in the invention.
| Number | Date | Country | |
|---|---|---|---|
| Parent | 15311188 | Mar 2017 | US |
| Child | 18070554 | US |
