Patent Application
20220290129
This invention relates to compositions of matter, methods and instruments for nucleic acid-guided nuclease editing of live microbial cells to increase biodegradation.
In the following discussion certain articles and methods will be described for background and introductory purposes. Nothing contained herein is to be construed as an “admission” of prior art. Applicant expressly reserves the right to demonstrate, where appropriate, that the methods referenced herein do not constitute prior art under the applicable statutory provisions.
There are many types of petroleum-based plastics that are manufactured and used widely in myriad ways. These synthetic plastics are difficult degrade due to high molecular weight, high hydrophobicity, and high chemical bond energy, such that they accumulate in the environment for a long time. Although many studies have reported microbial organisms with some ability to degrade petroleum-based plastics, significant degradation has not yet been achieved at commercially viable scale.
There is thus a need in the art for methods of developing and using modified microbes for the biodegradation of plastics. The present disclosure addresses this need.
This Summary is provided to introduce a selection of concepts in a simplified form that are further described below in the Detailed Description. This Summary is not intended to identify key or essential features of the claimed subject matter, nor is it intended to be used to limit the scope of the claimed subject matter. Other features, details, utilities, and advantages of the claimed subject matter will be apparent from the following written Detailed Description including those aspects illustrated in the accompanying drawings and defined in the appended claims.
The present disclosure relates to methods, compositions, and automated multi-module cell processing instruments for the production of synthetic microbes for biodegradation. The disclosure discloses methods of using nucleic-acid guided editing in microbial cell populations, e.g., bacterial and fungal cells, for production of synthetic microbes for biodegradation.
In specific aspects, the disclosure provides microbes produced by the disclosed methods, where the microbes are designed to efficiently biodegrade petroleum-based plastics. Examples of such petroleum-based substrates that can be degraded using the edited microbes described herein include, but are not limited to, polystyrene, polyethylene, polypropylene, polyurethane, polyethylene terephthalate, polyvinyl chloride and modifications or derivatives of each.
In some aspects, the microbes edited using the methods of the disclosure include bacterial cells. Known organisms with such activity include, but are not limited to, bacteria from the genera Pseudomonas, Bacillus, Brevibacillus, Cellulosimicrobium, Citrobacter, Kosakonia, Lysinibacillus, and Enterobacter. Specific bacterial strains include but are not limited to Enterobacter absuriae YT1, (Yang, et al., Environ. Sci. Technol., 48(23):13776-13784 (2014)); Bacillus sp. YP1 (Id.); Citrobacter sp. (Brandon, et al., Environ. Sci. Technol., 52(11):6526-6533 (2018)); Kosakonia sp. (Id.); Bacillus cereus strain A5 (Muhonja, et al., PLoS One, 13(7): e0198446 (2018)), Thermobifida fusca (Wei, et al., Advanced Science, 6(1900491) (2019)); Brevibacillus borstelensis strain B2,2 (Id.) and Ideonella sakaiensis (Taniguchi, et al., ACS Catal. 9(5):4089-4105 (2019)).
In some aspects, microbial organisms with known ability to biodegrade one or more plastics, such as those referenced above, can be used as the “starting” organisms that are further edited for improved biodegradation. In some aspects, the genes of organisms with ability to biodegrade plastic (or mutations found in homologous genes from these organisms) can be introduced as heterologous sequences into bacterial cells of another genera, species or strain.
In some aspects, the microbes edited using the methods of the disclosure include fungal cells, e.g., fungi of the genus Aspergillus.
In specific aspects, heterologous enzymes with known biodegradation capabilities are engineered into common research microbial species such as E. coli or S. cerevisiae.
In specific aspects, the microbes edited are found in the gut of organisms with known capabilities of digesting plastics, e.g. waxworms such as Zophobas atratus, or larvae of Tenebrio molitor (Brandon et al., supra), and Plodia interpunctella (Yang, et al., supra).
In some aspects, genes from one organism are heterologously introduced into another organism for genetic manipulation and optimization. The genes could either be integrated into the new host genome or carried on a plasmid. The optimized sequences are reintroduced to the original organism or another organism for further characterization or industrial application.
In certain embodiments, automated methods are used for nuclease-directed genome editing of one or more target genomic regions in multiple cells to enhance the ability of these cells to biodegrade petroleum-based plastics, the methods being performed in automated multi-module cell editing instruments. These methods can be used to generate libraries of living cells of interest with desired genomic changes. The automated methods carried out using the automated multi-module cell editing instruments described herein can be used with a variety of nuclease-directed genome editing techniques and can be used with or without use of one or more selectable markers.
The present disclosure thus provides, in selected embodiments, modules, instruments, and systems for automated multi-module cell editing to enhance a microbe's ability to biodegrade petroleum-based plastics, including nuclease-directed genome editing. Other specific embodiments of the automated multi-module cell editing instruments of the disclosure are designed for recursive genome editing, e.g., sequentially introducing multiple edits into genomes inside one or more cells of a cell population through two or more editing operations within the instruments.
These aspects and other features and advantages of the invention are described below in more detail.
The foregoing and other features and advantages of the present invention will be more fully understood from the following detailed description of illustrative embodiments taken in conjunction with the accompanying drawings in which:
It should be understood that the drawings are not necessarily to scale, and that like reference numbers refer to like features.
All of the functionalities described in connection with one embodiment are intended to be applicable to the additional embodiments described herein except where expressly stated or where the feature or function is incompatible with the additional embodiments. For example, where a given feature or function is expressly described in connection with one embodiment but not expressly mentioned in connection with an alternative embodiment, it should be understood that the feature or function may be deployed, utilized, or implemented in connection with the alternative embodiment unless the feature or function is incompatible with the alternative embodiment.
The practice of the techniques described herein may employ, unless otherwise indicated, conventional techniques and descriptions of organic chemistry, polymer technology, molecular biology (including recombinant techniques), cell biology, biochemistry and sequencing technology, which are within the skill of those who practice in the art. Such conventional techniques include polymer array synthesis, hybridization and ligation of polynucleotides, and detection of hybridization using a label. Specific illustrations of suitable techniques can be had by reference to the examples herein. However, other equivalent conventional procedures can, of course, also be used. Such conventional techniques and descriptions can be found in standard laboratory manuals such as Green, et al., Eds. (1999), Genome Analysis: A Laboratory Manual Series (Vols. I-IV); Weiner, Gabriel, Stephens, Eds. (2007), Genetic Variation: A Laboratory Manual; Dieffenbach, Dveksler, Eds. (2003), PCR Primer: A Laboratory Manual; Bowtell and Sambrook (2003), DNA Microarrays: A Molecular Cloning Manual; Mount (2004), Bioinformatics: Sequence and Genome Analysis; Sambrook and Russell (2006), Condensed Protocols from Molecular Cloning: A Laboratory Manual; and Sambrook and Russell (2002), Molecular Cloning: A Laboratory Manual (all from Cold Spring Harbor Laboratory Press); Stryer, L. (1995) Biochemistry (4th Ed.) W.H. Freeman, New York N.Y.; Gait, “Oligonucleotide Synthesis: A Practical Approach” 1984, IRL Press, London; Nelson and Cox (2000), Lehninger, Principles of Biochemistry 3rd Ed., W. H. Freeman Pub., New York, N.Y.; Berg et al. (2002) Biochemistry, 5th Ed., W.H. Freeman Pub., New York, N.Y.; all of which are herein incorporated in their entirety by reference for all purposes. CRISPR-specific techniques can be found in, e.g., Genome Editing and Engineering from TALENs and CRISPRs to Molecular Surgery, Appasani and Church (2018); and CRISPR: Methods and Protocols, Lindgren and Charpentier (2015); both of which are herein incorporated in their entirety by reference for all purposes.
Note that as used herein and in the appended claims, the singular forms “a,” “an,” and “the” include plural referents unless the context clearly dictates otherwise. Thus, for example, reference to “an oligonucleotide” refers to one or more oligonucleotides, and reference to “an automated system” includes reference to equivalent steps and methods for use with the system known to those skilled in the art, and so forth. Additionally, it is to be understood that terms such as “left,” “right,” “top,” “bottom,” “front,” “rear,” “side,” “height,” “length,” “width,” “upper,” “lower,” “interior,” “exterior,” “inner,” “outer” that may be used herein merely describe points of reference and do not necessarily limit embodiments of the present disclosure to any particular orientation or configuration. Furthermore, terms such as “first,” “second,” “third,” etc., merely identify one of a number of portions, components, steps, operations, functions, and/or points of reference as disclosed herein, and likewise do not necessarily limit embodiments of the present disclosure to any particular configuration or orientation.
Unless defined otherwise, all technical and scientific terms used herein have the same meaning as commonly understood by one of ordinary skill in the art to which this invention belongs. All publications mentioned herein are incorporated by reference for the purpose of describing and disclosing devices, methods and cell populations that may be used in connection with the presently described invention.
Where a range of values is provided, it is understood that each intervening value, between the upper and lower limit of that range and any other stated or intervening value in that stated range is encompassed within the invention. The upper and lower limits of these smaller ranges may independently be included in the smaller ranges, and are also encompassed within the invention, subject to any specifically excluded limit in the stated range. Where the stated range includes one or both of the limits, ranges excluding either or both of those included limits are also included in the invention.
In the following description, numerous specific details are set forth to provide a more thorough understanding of the present invention. However, it will be apparent to one of ordinary skill in the art that the present invention may be practiced without one or more of these specific details. In other instances, well-known features and procedures well known to those skilled in the art have not been described in order to avoid obscuring the invention.
The term “biodegradation” refers to the decomposition of organic material by microorganisms. Such organic materials include, but are not limited to, petroleum-based plastics and related materials.
The term “complementary” as used herein refers to Watson-Crick base pairing between nucleotides and specifically refers to nucleotides hydrogen bonded to one another with thymine or uracil residues linked to adenine residues by two hydrogen bonds and cytosine and guanine residues linked by three hydrogen bonds. In general, a nucleic acid includes a nucleotide sequence described as having a “percent complementarity” or “percent homology” to a specified second nucleotide sequence. For example, a nucleotide sequence may have 80%, 90%, or 100% complementarity to a specified second nucleotide sequence, indicating that 8 of 10, 9 of 10 or 10 of 10 nucleotides of a sequence are complementary to the specified second nucleotide sequence. For instance, the nucleotide sequence 3′-TCGA-5′ is 100% complementary to the nucleotide sequence 5′-AGCT-3′; and the nucleotide sequence 3′-TCGA-5′ is 100% complementary to a region of the nucleotide sequence 5′-TAGCTG-3′.
The term DNA “control sequences” refers collectively to promoter sequences, polyadenylation signals, transcription termination sequences, upstream regulatory domains, origins of replication, internal ribosome entry sites, nuclear localization sequences, enhancers, and the like, which collectively provide for the replication, transcription and translation of a coding sequence in a recipient cell. Not all of these types of control sequences need to be present so long as a selected coding sequence is capable of being replicated, transcribed and—for some components-translated in an appropriate host cell.
As used herein the term “donor DNA” or “donor nucleic acid” or “homology arm” refers to nucleic acid that is designed to introduce a DNA sequence modification (insertion, deletion, substitution) into a locus by homologous recombination using nucleic acid-guided nucleases. For homology-directed repair, the donor DNA must have sufficient homology to the regions flanking the “cut site” or site to be edited in the genomic target sequence. The length of the homology arm(s) will depend on, e.g., the type and size of the modification being made. In many instances and preferably, the donor DNA will have two regions of sequence homology (e.g., two homology arms) to the genomic target locus. Preferably, an “insert” region or “DNA sequence modification” region—the nucleic acid modification that one desires to be introduced into a genome target locus in a cell-will be located between two regions of homology. The DNA sequence modification may change one or more bases of the target genomic DNA sequence at one specific site or multiple specific sites. A change may include changing 1, 2, 3, 4, 5, 10, 15, 20, 25, 30, 35, 40, 50, 75, 100, 150, 200, 300, 400, or 500 or more base pairs of the target sequence. A deletion or insertion may be a deletion or insertion of 1, 2, 3, 4, 5, 10, 15, 20, 25, 30, 40, 50, 75, 100, 150, 200, 300, 400, or 500 or more base pairs of the target sequence.
The term “editing cassette” refers to a nucleic acid molecule comprising a coding sequence for transcription of a guide nucleic acid or gRNA covalently linked to a coding sequence for transcription of a donor DNA or homology arm.
The terms “guide nucleic acid” or “guide RNA” or “gRNA” refer to a polynucleotide comprising 1) a guide sequence capable of hybridizing to a genomic target locus, and 2) a scaffold sequence capable of interacting or complexing with a nucleic acid-guided nuclease.
“Homology” or “identity” or “similarity” refers to sequence similarity between two peptides or, more often in the context of the present disclosure, between two nucleic acid molecules. The term “homologous region” or “homology arm” refers to a region on the donor DNA with a certain degree of homology with the target genomic DNA sequence. Homology can be determined by comparing a position in each sequence which may be aligned for purposes of comparison. When a position in the compared sequence is occupied by the same base or amino acid, then the molecules are homologous at that position. A degree of homology between sequences is a function of the number of matching or homologous positions shared by the sequences.
The term “MAD7” refers to a CRISPR Type V nuclease derived from Eubacterium rectale, see U.S. Pat. Nos. 9,982,279 and 10,337,028.
“Nucleic acid-guided editing components” refers to one, some, or all of a nuclease, a guide nucleic acid, a donor nucleic acid, and recombination systems, if required.
The term “nickase” as used herein refers to a nuclease that cuts one strand of a double-stranded DNA at a specific recognition nucleotide sequence.
“Operably linked” refers to an arrangement of elements where the components so described are configured so as to perform their usual function. Thus, control sequences operably linked to a coding sequence are capable of effecting the transcription, and in some cases, the translation, of a coding sequence. The control sequences need not be contiguous with the coding sequence so long as they function to direct the expression of the coding sequence. Thus, for example, intervening untranslated yet transcribed sequences can be present between a promoter sequence and the coding sequence and the promoter sequence can still be considered “operably linked” to the coding sequence. In fact, such sequences need not reside on the same contiguous DNA molecule (i.e. chromosome) and may still have interactions resulting in altered regulation.
A “PAM mutation” refers to one or more edits to a target sequence that removes, mutates, or otherwise renders inactive a PAM or spacer region in the target sequence.
A “promoter” or “promoter sequence” is a DNA regulatory region capable of binding RNA polymerase and initiating transcription of a polynucleotide or polypeptide coding sequence such as messenger RNA, ribosomal RNA, small nuclear or nucleolar RNA, guide RNA, or any kind of RNA. Promoters may be constitutive or inducible. A “pol II promoter” is a regulatory sequence that is bound by RNA polymerase II to catalyze the transcription of DNA.
As used herein the term “selectable marker” refers to a gene introduced into a cell, which confers a trait suitable for artificial selection. General use selectable markers are well-known to those of ordinary skill in the art. Drug selectable markers such as ampicillin/carbenicillin, kanamycin, chloramphenicol, nourseothricin N-acetyl transferase, erythromycin, tetracycline, gentamicin, bleomycin, streptomycin, puromycin, hygromycin, blasticidin, and G418 may be employed. In other embodiments, selectable markers include, but are not limited to human nerve growth factor receptor (detected with a MAb, such as described in U.S. Pat. No. 6,365,373); truncated human growth factor receptor (detected with MAb); mutant human dihydrofolate reductase (DHFR; fluorescent MTX substrate available); secreted alkaline phosphatase (SEAP; fluorescent substrate available); human thymidylate synthase (TS; confers resistance to anti-cancer agent fluorodeoxyuridine); human glutathione S-transferase alpha (GSTA1; conjugates glutathione to the stem cell selective alkylator busulfan; chemoprotective selectable marker in CD34+ cells); CD24 cell surface antigen in hematopoietic stem cells; human CAD gene to confer resistance to N-phosphonacetyl-L-aspartate (PALA); human multi-drug resistance-1 (MDR-1; P-glycoprotein surface protein selectable by increased drug resistance or enriched by FACS); human CD25 (IL-2α; detectable by Mab-FITC); Methylguanine-DNA methyltransferase (MGMT; selectable by carmustine); rhamnose; and Cytidine deaminase (CD; selectable by Ara-C). “Selective medium” as used herein refers to cell growth medium to which has been added a chemical compound or biological moiety that selects for or against selectable markers.
The term “specifically binds” as used herein includes an interaction between two molecules, e.g., an engineered peptide antigen and a binding target, with a binding affinity represented by a dissociation constant of about 10−7M, about 10−8M, about 10−9 M, about 10−1° M, about 10−11M, about 10−12M, about 10−13M, about 10−14M or about 10−15M.
The terms “target genomic DNA sequence”, “target sequence”, or “genomic target locus” refer to any locus in vitro or in vivo, or in a nucleic acid (e.g., genome or episome) of a cell or population of cells, in which a change of at least one nucleotide is desired using a nucleic acid-guided nuclease editing system. The target sequence can be a genomic locus or extrachromosomal locus.
The term “variant” may refer to a polypeptide or polynucleotide that differs from a reference polypeptide or polynucleotide but retains essential properties. A typical variant of a polypeptide differs in amino acid sequence from another reference polypeptide. Generally, differences are limited so that the sequences of the reference polypeptide and the variant are closely similar overall and, in many regions, identical. A variant and reference polypeptide may differ in amino acid sequence by one or more modifications (e.g., substitutions, additions, and/or deletions). A variant of a polypeptide may be a conservatively modified variant. A substituted or inserted amino acid residue may or may not be one encoded by the genetic code (e.g., a non-natural amino acid). A variant of a polypeptide may be naturally occurring, such as an allelic variant, or it may be a variant that is not known to occur naturally.
A “vector” is any of a variety of nucleic acids that comprise a desired sequence or sequences to be delivered to and/or expressed in a cell. Vectors are typically composed of DNA, although RNA vectors are also available. Vectors include, but are not limited to, plasmids, fosmids, phagemids, virus genomes, BACs, YACs, PACs, synthetic chromosomes, and the like.
The compositions and methods described herein are employed to perform nuclease-directed genome editing to introduce desired edits to a population of microbial cells. In some embodiments, a single edit is introduced in a single round of editing. In some embodiments, multiple edits are introduced in a single round of editing using simultaneous editing, e.g., the introduction of two or more edits on a single vector. In some embodiments, recursive cell editing is performed where edits are introduced in successive rounds of editing.
A nucleic acid-guided nuclease complexed with an appropriate synthetic guide nucleic acid in a cell can cut the genome of the cell at a desired location. The guide nucleic acid helps the nucleic acid-guided nuclease recognize and cut the DNA at a specific target sequence (either a cellular target sequence or a curing target sequence). By manipulating the nucleotide sequence of the guide nucleic acid, the nucleic acid-guided nuclease may be programmed to target any DNA sequence for cleavage as long as an appropriate protospacer adjacent motif (PAM) is nearby. In certain aspects, the nucleic acid-guided nuclease editing system may use two separate guide nucleic acid molecules that combine to function as a guide nucleic acid, e.g., a CRISPR RNA (crRNA) and trans-activating CRISPR RNA (tracrRNA). In other aspects and preferably, the guide nucleic acid is a single guide nucleic acid construct that includes both 1) a guide sequence capable of hybridizing to a genomic target locus, and 2) a scaffold sequence capable of interacting or complexing with a nucleic acid-guided nuclease.
In general, a guide nucleic acid (e.g., gRNA) complexes with a compatible nucleic acid-guided nuclease and can then hybridize with a target sequence, thereby directing the nuclease to the target sequence. A guide nucleic acid can be DNA or RNA; alternatively, a guide nucleic acid may comprise both DNA and RNA. In some embodiments, a guide nucleic acid may comprise modified or non-naturally occurring nucleotides. In cases where the guide nucleic acid comprises RNA, the gRNA may be encoded by a DNA sequence on a polynucleotide molecule such as a plasmid, linear construct, or the coding sequence may and preferably does reside within an editing cassette. Methods and compositions for designing and synthesizing editing cassettes are described in U.S. Pat. Nos. 10,240,167; 10,266,849; 9,982,278; 10,351,877; 10,364,442; and 10,435,715; 10,465,207; 10, 669,559; 10,711,284; and 10,713,180 and U.S. Ser. No. 16/938,739, filed 24 Jul. 2020, all of which are incorporated by reference herein.
A guide nucleic acid comprises a guide sequence, where the guide sequence is a polynucleotide sequence having sufficient complementarity with a target sequence to hybridize with the target sequence and direct sequence-specific binding of a complexed nucleic acid-guided nuclease to the target sequence. The degree of complementarity between a guide sequence and the corresponding target sequence, when optimally aligned using a suitable alignment algorithm, is about or more than about 50%, 60%, 75%, 80%, 85%, 90%, 95%, 97.5%, 99%, or more. Optimal alignment may be determined with the use of any suitable algorithm for aligning sequences. In some embodiments, a guide sequence is about or more than about 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, 29, 30, 35, 40, 45, 50, 75, or more nucleotides in length. In some embodiments, a guide sequence is less than about 75, 50, 45, 40, 35, 30, 25, 20 nucleotides in length. Preferably the guide sequence is 10-30 or 15-20 nucleotides long, or 15, 16, 17, 18, 19, or 20 nucleotides in length.
In general, to generate an edit in the target sequence, the gRNA/nuclease complex binds to a target sequence as determined by the guide RNA, and the nuclease recognizes a protospacer adjacent motif (PAM) sequence adjacent to the target sequence. The target sequence can be any polynucleotide endogenous or exogenous to the microbial cell, or in vitro. A target sequence can be a sequence encoding a gene product (e.g., a protein) or a non-coding sequence (e.g., a regulatory polynucleotide, an intron, a PAM, a control sequence, etc.).
The guide nucleic acid may be and preferably is part of an editing cassette that encodes the donor nucleic acid that targets a cellular target sequence. Alternatively, the guide nucleic acid may not be part of the editing cassette and instead may be encoded on the editing vector backbone. For example, a sequence coding for a guide nucleic acid can be assembled or inserted into a vector backbone first, followed by insertion of the donor nucleic acid in, e.g., an editing cassette. In other cases, the donor nucleic acid in, e.g., an editing cassette can be inserted or assembled into a vector backbone first, followed by insertion of the sequence coding for the guide nucleic acid. Preferably, the sequence encoding the guide nucleic acid and the donor nucleic acid are located together in a rationally-designed editing cassette and are simultaneously inserted or assembled via gap repair into a linear plasmid or vector backbone to create an editing vector.
The target sequence is associated with a proto-spacer mutation (PAM), which is a short nucleotide sequence recognized by the gRNA/nuclease complex. The precise preferred PAM sequence and length requirements for different nucleic acid-guided nucleases vary; however, PAMs typically are 2-7 base-pair sequences adjacent or in proximity to the target sequence and, depending on the nuclease, can be 5′ or 3′ to the target sequence. Engineering of the PAM-interacting domain of a nucleic acid-guided nuclease may allow for alteration of PAM specificity, improve target site recognition fidelity, decrease target site recognition fidelity, or increase the versatility of a nucleic acid-guided nuclease.
In certain embodiments, the genome editing of a cellular target sequence both introduces a desired DNA change to a cellular target sequence, e.g., the genomic DNA of a cell, and removes, mutates, or renders inactive a proto-spacer mutation (PAM) region in the cellular target sequence. Rendering the PAM at the cellular target sequence inactive precludes additional editing of the cell genome at that cellular target sequence, e.g., upon subsequent exposure to a nucleic acid-guided nuclease complexed with a synthetic guide nucleic acid in later rounds of editing. Thus, cells having the desired cellular target sequence edit and an altered PAM can be selected for by using a nucleic acid-guided nuclease complexed with a synthetic guide nucleic acid complementary to the cellular target sequence. Cells that did not undergo the first editing event will be cut rendering a double-stranded DNA break, and thus will not continue to be viable. The cells containing the desired cellular target sequence edit and PAM alteration will not be cut, as these edited cells no longer contain the necessary PAM site and will continue to grow and propagate.
As for the nuclease component of the nucleic acid-guided nuclease editing system, a polynucleotide sequence encoding the nucleic acid-guided nuclease can be codon optimized for expression in particular microbial cell types, such as stem cells. The choice of nucleic acid-guided nuclease to be employed depends on many factors, such as what type of edit is to be made in the target sequence and whether an appropriate PAM is located close to the desired target sequence. Nucleases of use in the methods described herein include but are not limited to Cas 9, Cas 12/CpfI, MAD2, or MAD7 or other MADzymes. As with the guide nucleic acid, the nuclease is encoded by a DNA sequence on a vector and optionally is under the control of an inducible promoter. In some embodiments, the promoter may be separate from but the same as the promoter controlling transcription of the guide nucleic acid; that is, a separate promoter drives the transcription of the nuclease and guide nucleic acid sequences but the two promoters may be the same type of promoter. Alternatively, the promoter controlling expression of the nuclease may be different from the promoter controlling transcription of the guide nucleic acid; that is, e.g., the nuclease may be under the control of, e.g., the pTEF promoter, and the guide nucleic acid may be under the control of the, e.g., pCYC1 promoter.
Another component of the nucleic acid-guided nuclease system is the donor nucleic acid comprising homology to the cellular target sequence. The donor nucleic acid is on the same vector and even in the same editing cassette as the guide nucleic acid and preferably is (but not necessarily is) under the control of the same promoter as the editing gRNA (that is, a single promoter driving the transcription of both the editing gRNA and the donor nucleic acid). The donor nucleic acid is designed to serve as a template for homologous recombination with a cellular target sequence nicked or cleaved by the nucleic acid-guided nuclease as a part of the gRNA/nuclease complex. A donor nucleic acid polynucleotide may be of any suitable length, such as about or more than about 20, 25, 50, 75, 100, 150, 200, 500, or 1000 nucleotides in length, and up to 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13 and up to 20 kb in length if combined with a dual gRNA architecture as described in U.S. Ser. No. 62/869,24-, filed 1 Jul. 2019. In certain preferred aspects, the donor nucleic acid can be provided as an oligonucleotide of between 20-300 nucleotides, more preferably between 50-250 nucleotides. The donor nucleic acid comprises a region that is complementary to a portion of the cellular target sequence (e.g., a homology arm). When optimally aligned, the donor nucleic acid overlaps with (is complementary to) the cellular target sequence by, e.g., about 20, 25, 30, 35, 40, 50, 60, 70, 80, 90 or more nucleotides. In many embodiments, the donor nucleic acid comprises two homology arms (regions complementary to the cellular target sequence) flanking the mutation or difference between the donor nucleic acid and the cellular target sequence. The donor nucleic acid comprises at least one mutation or alteration compared to the cellular target sequence, such as an insertion, deletion, modification, or any combination thereof compared to the cellular target sequence.
As described in relation to the gRNA, the donor nucleic acid can be provided as part of a rationally-designed editing cassette, which is inserted into an editing plasmid backbone where the editing plasmid backbone may comprise a promoter to drive transcription of the editing gRNA and the donor DNA when the editing cassette is inserted into the editing plasmid backbone. Moreover, there may be more than one, e.g., two, three, four, or more editing gRNA/donor nucleic acid rationally-designed editing cassettes inserted into an editing vector; alternatively, a single rationally-designed editing cassette may comprise two to several editing gRNA/donor DNA pairs, where each editing gRNA is under the control of separate different promoters, separate like promoters, or where all gRNAs/donor nucleic acid pairs are under the control of a single promoter. In some embodiments the promoter driving transcription of the editing gRNA and the donor nucleic acid (or driving more than one editing gRNA/donor nucleic acid pair) is optionally an inducible promoter.
In addition to the donor nucleic acid, an editing cassette may comprise one or more primer sites. The primer sites can be used to amplify the editing cassette by using oligonucleotide primers; for example, if the primer sites flank one or more of the other components of the editing cassette. In addition, the editing cassette may comprise a barcode. A barcode is a unique DNA sequence that corresponds to the donor DNA sequence such that the barcode can identify the edit made to the corresponding cellular target sequence. The barcode typically comprises four or more nucleotides. In some embodiments, the editing cassettes comprise a collection or library editing gRNAs and of donor nucleic acids representing, e.g., gene-wide or genome-wide libraries of editing gRNAs and donor nucleic acids. The library of editing cassettes is cloned into vector backbones where, e.g., each different donor nucleic acid is associated with a different barcode. Also, in preferred embodiments, an editing vector or plasmid encoding components of the nucleic acid-guided nuclease system further encodes a nucleic acid-guided nuclease comprising one or more nuclear localization sequences (NLSs), such as about or more than about 1, 2, 3, 4, 5, 6, 7, 8, 9, 10, or more NLSs, particularly as an element of the nuclease sequence. In some embodiments, the engineered nuclease comprises NLSs at or near the amino-terminus, NLSs at or near the carboxy-terminus, or a combination.
Cells with a stably integrated genomic copy of the GFP gene can enable phenotypic detection of genomic edits of different classes (NHEJ, HDR, no edit) by flow cytometry, fluorescent cell imaging, or genotypic detection by sequencing of the genome-integrated GFP gene. Lack of editing, or perfect repair of cut events in the GFP gene result in cells that remain GFP-positive. Cut events that are repaired by the Non-Homologous End-Joining (NHEJ) pathway often result in nucleotide insertion or deletion events (Indel). These Indel edits often result in frame-shift mutations in the coding sequence that cause loss of GFP gene expression and fluorescence. Cut events that are repaired by the Homology-Directed Repair (HDR) pathway, using the GFP to BFP HDR donor as a repair template result in conversion of the cell fluorescence profile from that of GFP to that of BFP.
Many types of petroleum-based polymers are commonly used to make single-use plastic materials, namely polyethylene (PE, including low-density polyethylene (LDPE) and high-density polyethylene (HDPE)), polystyrene (PS), polypropylene (PP), polyvinyl chloride (PVC), and polyethylene terephthalate (PET). Biodegradation can be characterized using various parameters, including but not limited to gravimetric weight loss of the plastic, molecular weight shifts of the plastic polymers, presence of water-soluble daughter products of the plastic polymers, changes in weight-averaged Molar mass (Mw) and number-average Molar mass (Mn), ATP and ADP assays, scanning electron microscopy detection of surface degradation, FT-IR measurement to detect oxidative products formed during biodegradation, gas chromatography to measure biofragmentation and existence of saturated linear alkanes, size exclusion chromatography, and NMR spectroscopy.
The biodegradation process usually includes biofragmentation of the petroleum-based polymers by secreted enzymes, followed by bioassimilation of small cleavage fragments (molar mass must be less than 500 g/mol) by the microorganisms (Montazer, et al., Can. J. Microbiol., 65:1-11 (2019)). In some aspects, the polymers are degraded back to a single molecule version of the polymer.
The initial step involves hydroxylation of C—C bonds to generate primary or secondary alcohols, which are further oxidized to aldehydes or ketones, and then to carboxylic acids. Thus, microbial oxidation decreases the number of carbonyl-groups due to the formation of carboxylic acids. Carboxylated n-alkanes are analogous to fatty acids, which can be catabolized by bacteria via the β-oxidation system pathway.
Examples of specific forms of biodegradation of exemplary petroleum-based plastics, polyethylenes and polystyrenes, as well as organisms with biodegradation activities that could be further enhanced using the methods and instruments of the invention, are as follows.
There are a number of factors in microbial degradation of PE polymers that have a significant effect on the outcome and results of PE biodegradation experiments. Accessibility of enzymes secreted by the microorganisms to the PE carbon chain is an important factor in microbial biodegradation. The microstructure of all PE material consists simply of linear carbon chains held together by hydrogen bonds. However, according to different manufacturing processes and subsequently different physical arrangements of the linear chains, polyethylene polymers can have different densities and 3-dimensional structures: low molecular weight polyethylene (LMWPE); linear low-density polyethylene (LLDPE); low-density polyethylene (LDPE); high-density polyethylene (HDPE). The proposed mechanism for biodegradation of PE is shown on
It has been reported that Alkane hydroxylases (AlkBs), enzymes involved in the alkane hydroxylase system pathway, are known to degrade linear alkanes and are the best known enzymes involved in PE degradation in β-oxidation pathway (icon, H. J. and Kim M. N., Into Biodeteior. Biodegrad., 103, 141-146 (2015). The key enzymes of interest in the alkane hydroxylase system are monoxygenases. The number and types of Alkane hydroxylases vary greatly in different bacteria, in which the induction condition and amount of goal carbon in the alkane chain are completely different. (icon and Kim, Int. Biodeteior. Biodegrad., 114:202-208 (2016)).
The P. aeruginosa genome encodes two Alkane hydroxylases, alkB1 and alkB2, while the Rhodococcus sp. TMP2 genome encodes 5 Alkane hydroxylases (alkB1, alkB2, alkB3, alkB4, and alkB5) (Takei, et al., Biotechnol. Lett., 30:1447-1452 (2008). The Alkane hydroxylase system has been investigated studied best in P. putida GPol, which expressed an Alkane hydroxylase that participates in the first step of the n-alkane oxidation pathway by hydroxylating of the terminal carbon (Rojo, Environ. Microbiol., 11:2477-2490 (2009)). AlkB of Pseudomonas aeruginosa strain E7 actively degraded low molar mass PE and played a central role in the mineralization of LAMPE into CO2 (Yoon, et al., J. Bioremed. Biodegrad., 3:145 (2012)). Also, AlkB cloned and expressed in Pseudomonas sp. E4 was active in the early stage of in LMWPE biodegradation, even in the absence of the other specific enzymes like rubredoxin and rubredoxin reductase. Laccase enzymes (phenol oxidases) expressed by Rodococcus rubber are multi-copper enzymes that have also been shown to play a major role in PE biodegradation (Santo, et al., Int. Biodeter. Biodegrad., 84:204-210 (2013)).
LDPE, mainly used to make plastic carry bags and food packaging materials, is the most abundant petroleum-polymer on earth, and represents up to 64% of single-use plastics that are discarded within a short period after use, resulting in massive and rapid accumulation in the environment (Ragaert, et al., Waste Manag., 69:24-58 (2017); Montazer, et al., Polymers, 12:123-147 (2020)). Despite recycling and energy recovery efforts, the harmful impacts of virtually “non-biodegradable” LDPE waste accumulation in landfill and in the oceans are increasing. Unlike organic wastes discarded by humans, polyethylene (PE), and other petroleum-based plastics, are extremely recalcitrant to natural biodegradation processes.
In addition to LDPE, high density polyethylene (HDPE) is also a very common non-degradable petroleum-based plastic (Devi, et al., Mar. Pollut. Bull., 96:32-40 (2015)). LDPE degrades faster than HDPE, possibly due to the fact that the polymer chains of LDPE are more closely packed than those of HDPE and that LDPE has a lower content of vinylidene defects, which have been shown to be directly correlated with oxidization of the polymer. Further, there are fewer tertiary carbons in HPDE and its molar mass is much higher, possibly making it more difficult for microorganisms and/or their oxidizing enzymes to access the polymer chains (Sudhakar, et al., Int. Biodeter. Biodegrad., 61:203-213 (2008)). The biodegradability of various pre-treated polyethylene materials, including HDPE films, LDPE films, and linear low-density polyethylene (LLDPE) films of different thicknesses, and containing pro-oxidant additives, by Rhodococcus rhodochrous (one of the most efficient bacteria for PE biodegradation) was also examined (Fontanella, et al., Polym. Degrad. Stab., 95; 1011-1021 (2010)). Biodegradation and microbial growth were measured by ATP and ADP assays, scanning electron microscopy, FTIR measurement, size exclusion chromatography, and NMR spectroscopy (Id.). Although all samples, except cobalt containing samples, were degraded by R. rhodochrous, HDPE was the least degraded and mineralization reached less than 6% after 317 days of incubation (Id.).
Biodegradation of HDPE by Aspergillus spp., was examined based on weight loss and FTIR spectrophotometric analysis (Devi, supra.) The biodegradation of unpretreated and thermally pretreated low- and high-density polyethylenes by Bacillus sp. was assessed for one year using SEM analysis (Id.). Biodegradation was also evaluated by FTIR spectroscopy analysis and weight loss percentage. Degradation of un-treated pure samples according to weight loss percentage were 10% and 3.5% in cases of LDPE and HDPE, respectively. The ability of fungal isolates was proved to utilize virgin polyethylene as the carbon source without any pre-treatment and pro-oxidant additives (Id.)
A number of studies have identified organisms that show promise in the biodegradation of synthetic plastics, and on PE in particular, including those listed in Table 1:
Acinetobacter
bumannii
Arthobacter defluvii
Bacillus
amyloliquefaciens
Bacillussubtilis
Bacillus pumilus
Bacillus subtillis
Mar. Pollut. Bull. 77, 100-106
Bacillus ssp.
Biodetior. Biodegrad., 65,
Bacillus sphericus
Biodeterior. Biodegrad., 103,
Bacillus megaterium
Bacillus subtilis
Biodeterior. Biodegrad., 65,
Bacillus cereus (MIX
Bacillus
amyloliquefaciens
J. Pharm. Pharm. Sci., 4, 690-
Bacillus subtilis
Procedia Technol., 24, 232-
Bacillus pumilus
Bacillus subtilis
Brevibacillus
borstelensis
Microbiol. 98, 1093-1100
Brevibacillus
Sci. J., 3, 95-98 (2010)
Chryseobacterium
gleum
Eur. Polym. J. 52, 146-153
Comamonas sp.
Mater. 324, 634-644(2017)
Delftia sp.
Kocuria palustris
Microbacterium
paraoxydans
Test. 3, 1094-1099 (2012)
Pseudomonas sp.
Trop. 51, 629-633 (2003)
Pseudomonas
aeruginosa
Biodeter. Biodegrad. 103,
Pseudomonas sp.
Bioremed. Biodegrad., 3, 145
Pseudomonas sp.
Res. Cent. J. Intern. 2, 200-
Pseudomonas
citronellolis
Pseudomonas sp.
Pseudomonas
aeroginosa
Microbiol. 52, 411-419 (2012)
Pseudomonas putida
Pseudomonas
siringae
Pseudomonas sp.
Rhodococcus ruber
Microbiol. Biotechnol. 65, 97-
Rhodococcus ruber
Microbiol. Biotechnol. 72,
Rhodococcus ruber
Biodegrad. 84, 204-210
Rhodococcus
rhorocuros
Degrad. Stab. 81, 441-452
Rhodococcus
rhorocuros
Degrad. Stab. 95, 1011-1021
Rhodococcus sp.
Rhodococcus sp.
Biodegrad. 63, 354-357
Staphylococcus
arlettae
Environ. Sci. Toxicol. Food
Technol. 10, 1-7 (2016)
Stentrophomonas sp.
Stentrophomonas
pavanii
Biodeterior. Biodegrad. 113,
Streptomyces spp.
Degrad. Stab. 62, 361-365
Polystyrene (PS) is the third most important polymer in use, with Styrofoam—an expanded polystyrene foam—a major product of environmental concern. Previous studies have been conducted on the biodegradation of plastic, in which several bacteria and fungi were found to be capable of degrading plastic materials, including polystyrene, but at low degradation rates (Shah, et al., Biotechnol. Adv., 26(3):246-265 (2008); Tokiwa, Int. J. Mol., 10(9):3722-3742 (2009)). Specific microorganisms, such as Rhodococcus ruber, have been shown to colonize PS film surfaces (e.g., see Riudavets, et al., J. Stored Prod. Res., 43(4):564-570 (2007)), although they have not been shown to directly change the physical and chemical properties of PS. Other organisms, such as Exiguobacterium sp. strain YT2, were found to form a biofilm on a PS film and made obvious pits and cavities, and release water-soluble daughter products, but were only able to degrade around 7.4% of the PS pieces over a 60-day incubation period. Yang, et al., Environ. Sci. Technol., 49, 12087-12093 (2015).
In specific aspects, the disclosure provides methods for improving nuclease-directed editing of cells using enrichment means to identify cells that have received the editing components needed to perform the intended editing operation.
In some aspects, the enrichment handle and method can be based on a positive versus negative signal of the surrogate. In other aspects, the enrichment method can be based on a threshold level of a surrogate, e.g., a high level of an enrichment handle versus a low or absent level of an enrichment handle.
In some aspects, the disclosure provides methods for improving nuclease-directed editing rates by enriching for microbial cells that have received an HDR donor, e.g., identifying cells that are more likely to have received the editing.
In specific aspects of the invention, the HDR is improved using fusion proteins that retain certain characteristics of RNA-directed nucleases (e.g., CRISPR nucleases) and also utilize other enzymatic activities, e.g., replication inhibition, reverse transcriptase activity, transcription enhancement activity, and the like. These nuclease fusion proteins can be used in nuclease-directed editing using the disclosed methods, with or without the enrichment methods as disclosed herein.
In certain aspects, the protocols utilize both the RNA-directed nuclease fusion proteins and a selection. Use of fusion proteins and enrichment for these editing methods may involve a single enrichment method for HDR donor, the guide nucleic acid, and the nuclease, or two or more separate enrichment events for one or more of these elements. Methods and compositions for designing and synthesizing editing cassettes where the gRNA and homology region or arm (i.e., donor DNA) are described in U.S. Pat. Nos. 10,240,167; 10,266,849; 9,982,278; 10,351,877; 10,364,442; and 10,435,715; 10,465,207; 10, 669,559; 10,711,284; and 10,713,180 and U.S. Ser. No. 16/938,739, filed 24 Jul. 2020, all of which are incorporated by reference herein.
In specific aspects, the cells receiving the HDR donor can be enriched using an initial enrichment step, e.g., using an antibiotic selection or fluorescent detection, following by an enrichment step using an enrichment of the cells receiving and expressing the co-introduced cell surface antigen.
In some implementations, the reagent cartridges 210 are disposable kits comprising reagents and cells for use in the automated multi-module cell processing/editing instrument 200. For example, a user may open and position each of the reagent cartridges 210 comprising various desired inserts and reagents within the chassis of the automated multi-module cell editing instrument 200 prior to activating cell processing. Further, each of the reagent cartridges 210 may be inserted into receptacles in the chassis having different temperature zones appropriate for the reagents contained therein.
Also illustrated in
Inserts or components of the reagent cartridges 210, in some implementations, are marked with machine-readable indicia (not shown), such as bar codes, for recognition by the robotic handling system 258. For example, the robotic liquid handling system 258 may scan one or more inserts within each of the reagent cartridges 210 to confirm contents. In other implementations, machine-readable indicia may be marked upon each reagent cartridge 210, and a processing system (not shown, but see element 237 of
Inside the chassis 290, in some implementations, will be most or all of the components described in relation to
The drive engagement mechanism 312 engages with a motor (not shown) to rotate the vial. In some embodiments, the motor drives the drive engagement mechanism 312 such that the rotating growth vial 300 is rotated in one direction only, and in other embodiments, the rotating growth vial 300 is rotated in a first direction for a first amount of time or periodicity, rotated in a second direction (i.e., the opposite direction) for a second amount of time or periodicity, and this process may be repeated so that the rotating growth vial 300 (and the cell culture contents) are subjected to an oscillating motion. Further, the choice of whether the culture is subjected to oscillation and the periodicity therefor may be selected by the user. The first amount of time and the second amount of time may be the same or may be different. The amount of time may be 1, 2, 3, 4, 5, or more seconds, or may be 1, 2, 3, 4 or more minutes. In another embodiment, in an early stage of cell growth the rotating growth vial 400 may be oscillated at a first periodicity (e.g., every 60 seconds), and then a later stage of cell growth the rotating growth vial 300 may be oscillated at a second periodicity (e.g., every one second) different from the first periodicity.
The rotating growth vial 300 may be reusable or, preferably, the rotating growth vial is consumable. In some embodiments, the rotating growth vial is consumable and is presented to the user pre-filled with growth medium, where the vial is hermetically sealed at the open end 304 with a foil seal. A medium-filled rotating growth vial packaged in such a manner may be part of a kit for use with a stand-alone cell growth device or with a cell growth module that is part of an automated multi-module cell processing system. To introduce cells into the vial, a user need only pipette up a desired volume of cells and use the pipette tip to punch through the foil seal of the vial. Open end 304 may optionally include an extended lip 302 to overlap and engage with the cell growth device. In automated systems, the rotating growth vial 300 may be tagged with a barcode or other identifying means that can be read by a scanner or camera (not shown) that is part of the automated system.
The volume of the rotating growth vial 300 and the volume of the cell culture (including growth medium) may vary greatly, but the volume of the rotating growth vial 300 must be large enough to generate a specified total number of cells. In practice, the volume of the rotating growth vial 300 may range from 1-250 mL, 2-100 mL, from 5-80 mL, 10-50 mL, or from 12-35 mL. Likewise, the volume of the cell culture (cells+growth media) should be appropriate to allow proper aeration and mixing in the rotating growth vial 400. Proper aeration promotes uniform cellular respiration within the growth media. Thus, the volume of the cell culture should be approximately 5-85% of the volume of the growth vial or from 20-60% of the volume of the growth vial. For example, for a 30 mL growth vial, the volume of the cell culture would be from about 1.5 mL to about 26 mL, or from 6 mL to about 18 mL.
The rotating growth vial 300 preferably is fabricated from a bio-compatible optically transparent material—or at least the portion of the vial comprising the light path(s) is transparent. Additionally, material from which the rotating growth vial is fabricated should be able to be cooled to about 4° C. or lower and heated to about 55° C. or higher to accommodate both temperature-based cell assays and long-term storage at low temperatures. Further, the material that is used to fabricate the vial must be able to withstand temperatures up to 55° C. without deformation while spinning. Suitable materials include cyclic olefin copolymer (COC), glass, polyvinyl chloride, polyethylene, polyamide, polypropylene, polycarbonate, poly(methyl methacrylate (PMMA), polysulfone, polyurethane, and co-polymers of these and other polymers. Preferred materials include polypropylene, polycarbonate, or polystyrene. In some embodiments, the rotating growth vial is inexpensively fabricated by, e.g., injection molding or extrusion.
The motor 338 engages with drive mechanism 312 and is used to rotate the rotating growth vial 300. In some embodiments, motor 338 is a brushless DC type drive motor with built-in drive controls that can be set to hold a constant revolution per minute (RPM) between 0 and about 3000 RPM. Alternatively, other motor types such as a stepper, servo, brushed DC, and the like can be used. Optionally, the motor 338 may also have direction control to allow reversing of the rotational direction, and a tachometer to sense and report actual RPM. The motor is controlled by a processor (not shown) according to, e.g., standard protocols programmed into the processor and/or user input, and the motor may be configured to vary RPM to cause axial precession of the cell culture thereby enhancing mixing, e.g., to prevent cell aggregation, increase aeration, and optimize cellular respiration.
Main housing 336, end housings 352 and lower housing 332 of the cell growth device 330 may be fabricated from any suitable, robust material including aluminum, stainless steel, and other thermally conductive materials, including plastics. These structures or portions thereof can be created through various techniques, e.g., metal fabrication, injection molding, creation of structural layers that are fused, etc. Whereas the rotating growth vial 300 is envisioned in some embodiments to be reusable, but preferably is consumable, the other components of the cell growth device 330 are preferably reusable and function as a stand-alone benchtop device or as a module in a multi-module cell processing system.
The processor (not shown) of the cell growth device 330 may be programmed with information to be used as a “blank” or control for the growing cell culture. A “blank” or control is a vessel containing cell growth medium only, which yields 100% transmittance and 0 OD, while the cell sample will deflect light rays and will have a lower percent transmittance and higher OD. As the cells grow in the media and become denser, transmittance will decrease and OD will increase. The processor (not shown) of the cell growth device 330—may be programmed to use wavelength values for blanks commensurate with the growth media typically used in cell culture (whether, e.g., mammalian cells, bacterial cells, animal cells, yeast cells, etc.). Alternatively, a second spectrophotometer and vessel may be included in the cell growth device 330, where the second spectrophotometer is used to read a blank at designated intervals.
In use, cells are inoculated (cells can be pipetted, e.g., from an automated liquid handling system or by a user) into pre-filled growth media of a rotating growth vial 300 by piercing though the foil seal or film. The programmed software of the cell growth device 330 sets the control temperature for growth, typically 30° C., then slowly starts the rotation of the rotating growth vial 300. The cell/growth media mixture slowly moves vertically up the wall due to centrifugal force allowing the rotating growth vial 300 to expose a large surface area of the mixture to a normal oxygen environment. The growth monitoring system takes either continuous readings of the OD or OD measurements at pre-set or pre-programmed time intervals. These measurements are stored in internal memory and if requested the software plots the measurements versus time to display a growth curve. If enhanced mixing is required, e.g., to optimize growth conditions, the speed of the vial rotation can be varied to cause an axial precession of the liquid, and/or a complete directional change can be performed at programmed intervals. The growth monitoring can be programmed to automatically terminate the growth stage at a pre-determined OD, and then quickly cool the mixture to a lower temperature to inhibit further growth.
One application for the cell growth device 330 is to constantly measure the optical density of a growing cell culture. One advantage of the described cell growth device is that optical density can be measured continuously (kinetic monitoring) or at specific time intervals; e.g., every 5, 10, 15, 20, 30 45, or 60 seconds, or every 1, 2, 3, 4, 5, 6, 7, 8, 9, or 10 minutes. While the cell growth device 330 has been described in the context of measuring the optical density (OD) of a growing cell culture, it should, however, be understood by a skilled artisan given the teachings of the present specification that other cell growth parameters can be measured in addition to or instead of cell culture OD. As with optional measure of cell growth in relation to the solid wall device or module described supra, spectroscopy using visible, UV, or near infrared (NIR) light allows monitoring the concentration of nutrients and/or wastes in the cell culture and other spectroscopic measurements may be made; that is, other spectral properties can be measured via, e.g., dielectric impedance spectroscopy, visible fluorescence, fluorescence polarization, or luminescence. Additionally, the cell growth device 330 may include additional sensors for measuring, e.g., dissolved oxygen, carbon dioxide, pH, conductivity, and the like. For additional details regarding rotating growth vials and cell growth devices see U.S. Pat. No. 10,435,662, issued 8 Oct. 2019; U.S. Pat. No. 10,443,031, issued 15 Oct. 2019; and U.S. Ser. No. 16/552,981, filed 27 Aug. 2019 and Ser. No. 16/780,640, filed 3 Feb. 2020.
As described above in relation to the rotating growth vial and cell growth module, in order to obtain an adequate number of cells for transformation or transfection, cells typically are grown to a specific optical density in medium appropriate for the growth of the cells of interest; however, for effective transformation or transfection, it is desirable to decrease the volume of the cells as well as render the cells competent via buffer or medium exchange. Thus, one sub-component or module that is desired in cell processing systems to perform the methods described herein is a module or component that can grow, perform buffer exchange, and/or concentrate cells and render them competent so that they may be transformed or transfected with the nucleic acids needed for engineering or editing the cell's genome.
Permeate/filtrate member 420 is seen in the middle of
On the left of
A membrane or filter is disposed between the retentate and permeate members, where fluids can flow through the membrane but cells cannot and are thus retained in the flow channel disposed in the retentate member. Filters or membranes appropriate for use in the TFF device/module are those that are solvent resistant, are contamination free during filtration, and are able to retain the types and sizes of cells of interest. For example, in order to retain small cell types such as bacterial cells, pore sizes can be as low as 0.2 μm, however for other cell types, the pore sizes can be as high as 20 μm. Indeed, the pore sizes useful in the TFF device/module include filters with sizes from 0.20 μm, 0.21 μm, 0.22 μm, 0.23 μm, 0.24 μm, 0.25 μm, 0.26 μm, 0.27 μm, 0.28 μm, 0.29 μm, 0.30 μm, 0.31 μm, 0.32 μm, 0.33 μm, 0.34 μm, 0.35 μm, 0.36 μm, 0.37 μm, 0.38 μm, 0.39 μm, 0.40 μm, 0.41 μm, 0.42 μm, 0.43 μm, 0.44 μm, 0.45 μm, 0.46 μm, 0.47 μm, 0.48 μm, 0.49 μm, 0.50 μm and larger. The filters may be fabricated from any suitable non-reactive material including cellulose mixed ester (cellulose nitrate and acetate) (CME), polycarbonate (PC), polyvinylidene fluoride (PVDF), polyethersulfone (PES), polytetrafluoroethylene (PTFE), nylon, glass fiber, or metal substrates as in the case of laser or electrochemical etching.
The length of the channel structure 402 may vary depending on the volume of the cell culture to be grown and the optical density of the cell culture to be concentrated. The length of the channel structure typically is from 60 mm to 300 mm, or from 70 mm to 200 mm, or from 80 mm to 100 mm. The cross-section configuration of the flow channel 402 may be round, elliptical, oval, square, rectangular, trapezoidal, or irregular. If square, rectangular, or another shape with generally straight sides, the cross section may be from about 10 μm to 1000 μm wide, or from 200 μm to 800 μm wide, or from 300 μm to 700 μm wide, or from 400 μm to 600 μm wide; and from about 10 μm to 1000 μm high, or from 200 μm to 800 μm high, or from 300 μm to 700 μm high, or from 400 μm to 600 μm high. If the cross section of the flow channel 102 is generally round, oval or elliptical, the radius of the channel may be from about 50 μm to 1000 μm in hydraulic radius, or from 5 μm to 800 μm in hydraulic radius, or from 200 μm to 700 μm in hydraulic radius, or from 300 μm to 600 μm wide in hydraulic radius, or from about 200 to 500 μm in hydraulic radius. Moreover, the volume of the channel in the retentate 422 and permeate 420 members may be different depending on the depth of the channel in each member.
The TFF device may be fabricated from any robust material in which channels (and channel branches) may be milled including stainless steel, silicon, glass, aluminum, or plastics including cyclic-olefin copolymer (COC), cyclo-olefin polymer (COP), polystyrene, polyvinyl chloride, polyethylene, polyamide, polyethylene, polypropylene, acrylonitrile butadiene, polycarbonate, polyetheretheketone (PEEK), poly(methyl methylacrylate) (PMMA), polysulfone, and polyurethane, and co-polymers of these and other polymers. If the TFF device/module is disposable, preferably it is made of plastic. In some embodiments, the material used to fabricate the TFF device/module is thermally-conductive so that the cell culture may be heated or cooled to a desired temperature. In certain embodiments, the TFF device is formed by precision mechanical machining, laser machining, electro discharge machining (for metal devices); wet or dry etching (for silicon devices); dry or wet etching, powder or sandblasting, photostructuring (for glass devices); or thermoforming, injection molding, hot embossing, or laser machining (for plastic devices) using the materials mentioned above that are amenable to this mass production techniques.
The overall work flow for cell growth comprises loading a cell culture to be grown into a first retentate reservoir, optionally bubbling air or an appropriate gas through the cell culture, passing or flowing the cell culture through the first retentate port then tangentially through the TFF channel structure while collecting medium or buffer through one or both of the permeate ports 406, collecting the cell culture through a second retentate port 404 into a second retentate reservoir, optionally adding additional or different medium to the cell culture and optionally bubbling air or gas through the cell culture, then repeating the process, all while measuring, e.g., the optical density of the cell culture in the retentate reservoirs continuously or at desired intervals. Measurements of optical densities (OD) at programmed time intervals are accomplished using a 600 nm Light Emitting Diode (LED) that has been columnated through an optic into the retentate reservoir(s) containing the growing cells. The light continues through a collection optic to the detection system which consists of a (digital) gain-controlled silicone photodiode. Generally, optical density is shown as the absolute value of the logarithm with base 10 of the power transmission factors of an optical attenuator: OD=−log 10 (Power out/Power in). Since OD is the measure of optical attenuation—that is, the sum of absorption, scattering, and reflection—the TFF device OD measurement records the overall power transmission, so as the cells grow and become denser in population, the OD (the loss of signal) increases. The OD system is pre-calibrated against OD standards with these values stored in an on-board memory accessible by the measurement program.
In the channel structure, the membrane bifurcating the flow channels retains the cells on one side of the membrane (the retentate side 422) and allows unwanted medium or buffer to flow across the membrane into a filtrate or permeate side (e.g., permeate member 420) of the device. Bubbling air or other appropriate gas through the cell culture both aerates and mixes the culture to enhance cell growth. During the process, medium that is removed during the flow through the channel structure is removed through the permeate/filtrate ports 406. Alternatively, cells can be grown in one reservoir with bubbling or agitation without passing the cells through the TFF channel from one reservoir to the other.
The overall work flow for cell concentration using the TFF device/module involves flowing a cell culture or cell sample tangentially through the channel structure. As with the cell growth process, the membrane bifurcating the flow channels retains the cells on one side of the membrane and allows unwanted medium or buffer to flow across the membrane into a permeate/filtrate side (e.g., permeate member 420) of the device. In this process, a fixed volume of cells in medium or buffer is driven through the device until the cell sample is collected into one of the retentate ports 404, and the medium/buffer that has passed through the membrane is collected through one or both of the permeate/filtrate ports 406. All types of prokaryotic and eukaryotic cells—both adherent and non-adherent cells—can be grown in the TFF device. Adherent cells may be grown on beads or other cell scaffolds suspended in medium that flow through the TFF device.
The medium or buffer used to suspend the cells in the cell concentration device/module may be any suitable medium or buffer for the type of cells being transformed or transfected, such as LB, SOC, TPD, YPG, YPAD, MEM, DMEM, IMDM, RPMI, Hanks', PBS and Ringer's solution, where the media may be provided in a reagent cartridge as part of a kit. For culture of adherent cells, cells may be disposed on beads, microcarriers, or other type of scaffold suspended in medium. Most normal mammalian tissue-derived cells— except those derived from the hematopoietic system—are anchorage dependent and need a surface or cell culture support for normal proliferation. In the rotating growth vial described herein, microcarrier technology is leveraged. Microcarriers of particular use typically have a diameter of 100-300 μm and have a density slightly greater than that of the culture medium (thus facilitating an easy separation of cells and medium for, e.g., medium exchange) yet the density must also be sufficiently low to allow complete suspension of the carriers at a minimum stirring rate in order to avoid hydrodynamic damage to the cells. Many different types of microcarriers are available, and different microcarriers are optimized for different types of cells. There are positively charged carriers, such as Cytodex 1 (dextran-based, GE Healthcare), DE-52 (cellulose-based, Sigma-Aldrich Labware), DE-53 (cellulose-based, Sigma-Aldrich Labware), and HLX 11-170 (polystyrene-based); collagen- or ECM- (extracellular matrix) coated carriers, such as Cytodex 3 (dextran-based, GE Healthcare) or HyQ-sphere Pro-F 102-4 (polystyrene-based, Thermo Scientific); non-charged carriers, like HyQ-sphere P 102-4 (Thermo Scientific); or macroporous carriers based on gelatin (Cultisphere, Percell Biolytica) or cellulose (Cytopore, GE Healthcare).
In both the cell growth and concentration processes, passing the cell sample through the TFF device and collecting the cells in one of the retentate ports 404 while collecting the medium in one of the permeate/filtrate ports 406 is considered “one pass” of the cell sample. The transfer between retentate reservoirs “flips” the culture. The retentate and permeatee ports collecting the cells and medium, respectively, for a given pass reside on the same end of TFF device/module with fluidic connections arranged so that there are two distinct flow layers for the retentate and permeate/filtrate sides, but if the retentate port 404 resides on the retentate member of device/module (that is, the cells are driven through the channel above the membrane and the filtrate (medium) passes to the portion of the channel below the membrane), the permeate/filtrate port 406 will reside on the permeate member of device/module and vice versa (that is, if the cell sample is driven through the channel below the membrane, the filtrate (medium) passes to the portion of the channel above the membrane). Due to the high pressures used to transfer the cell culture and fluids through the flow channel of the TFF device, the effect of gravity is negligible.
At the conclusion of a “pass” in either of the growth and concentration processes, the cell sample is collected by passing through the retentate port 404 and into the retentate reservoir (not shown). To initiate another “pass”, the cell sample is passed again through the TFF device, this time in a flow direction that is reversed from the first pass. The cell sample is collected by passing through the retentate port 404 and into retentate reservoir (not shown) on the opposite end of the device/module from the retentate port 404 that was used to collect cells during the first pass. Likewise, the medium/buffer that passes through the membrane on the second pass is collected through the permeate port 406 on the opposite end of the device/module from the permeate port 406 that was used to collect the filtrate during the first pass, or through both ports. This alternating process of passing the retentate (the concentrated cell sample) through the device/module is repeated until the cells have been grown to a desired optical density, and/or concentrated to a desired volume, and both permeate ports (i.e., if there are more than one) can be open during the passes to reduce operating time. In addition, buffer exchange may be effected by adding a desired buffer (or fresh medium) to the cell sample in the retentate reservoir, before initiating another “pass”, and repeating this process until the old medium or buffer is diluted and filtered out and the cells reside in fresh medium or buffer. Note that buffer exchange and cell growth may (and typically do) take place simultaneously, and buffer exchange and cell concentration may (and typically do) take place simultaneously. For further information and alternative embodiments on TFFs see, e.g., U.S. Ser. No. 62/728,365, filed 7 Sep. 2018; 62/857,599, filed 5 Jun. 2019; and 62/867,415, filed 27 Jun. 2019.
In one embodiment, the reagent reservoirs or reservoirs 504 of reagent cartridge 500 are configured to hold various size tubes, including, e.g., 250 ml tubes, 25 ml tubes, 10 ml tubes, 5 ml tubes, and Eppendorf or microcentrifuge tubes. In yet another embodiment, all reservoirs may be configured to hold the same size tube, e.g., 5 ml tubes, and reservoir inserts may be used to accommodate smaller tubes in the reagent reservoir. In yet another embodiment—particularly in an embodiment where the reagent cartridge is disposable—the reagent reservoirs hold reagents without inserted tubes. In this disposable embodiment, the reagent cartridge may be part of a kit, where the reagent cartridge is pre-filled with reagents and the receptacles or reservoirs sealed with, e.g., foil, heat seal acrylic or the like and presented to a consumer where the reagent cartridge can then be used in an automated multi-module cell processing instrument. As one of ordinary skill in the art will appreciate given the present disclosure, the reagents contained in the reagent cartridge will vary depending on work flow; that is, the reagents will vary depending on the processes to which the cells are subjected in the automated multi-module cell processing instrument, e.g., protein production, cell transformation and culture, cell editing, etc.
Reagents such as cell samples, enzymes, buffers, nucleic acid vectors, expression cassettes, proteins or peptides, reaction components (such as, e.g., MgCl2, dNTPs, nucleic acid assembly reagents, gap repair reagents, and the like), wash solutions, ethanol, and magnetic beads for nucleic acid purification and isolation, etc. may be positioned in the reagent cartridge at a known position. In some embodiments of cartridge 500, the cartridge comprises a script (not shown) readable by a processor (not shown) for dispensing the reagents. Also, the cartridge 500 as one component in an automated multi-module cell processing instrument may comprise a script specifying two, three, four, five, ten or more processes to be performed by the automated multi-module cell processing instrument. In certain embodiments, the reagent cartridge is disposable and is pre-packaged with reagents tailored to performing specific cell processing protocols, e.g., genome editing or protein production. Because the reagent cartridge contents vary while components/modules of the automated multi-module cell processing instrument or system may not, the script associated with a particular reagent cartridge matches the reagents used and cell processes performed. Thus, e.g., reagent cartridges may be pre-packaged with reagents for genome editing and a script that specifies the process steps for performing genome editing in an automated multi-module cell processing instrument, or, e.g., reagents for protein expression and a script that specifies the process steps for performing protein expression in an automated multi-module cell processing instrument.
For example, the reagent cartridge may comprise a script to pipette competent cells from a reservoir, transfer the cells to a transformation module, pipette a nucleic acid solution comprising a vector with expression cassette from another reservoir in the reagent cartridge, transfer the nucleic acid solution to the transformation module, initiate the transformation process for a specified time, then move the transformed cells to yet another reservoir in the reagent cassette or to another module such as a cell growth module in the automated multi-module cell processing instrument. In another example, the reagent cartridge may comprise a script to transfer a nucleic acid solution comprising a vector from a reservoir in the reagent cassette, nucleic acid solution comprising editing oligonucleotide cassettes in a reservoir in the reagent cassette, and a nucleic acid assembly mix from another reservoir to the nucleic acid assembly/desalting module, if present. The script may also specify process steps performed by other modules in the automated multi-module cell processing instrument. For example, the script may specify that the nucleic acid assembly/desalting reservoir be heated to 50° C. for 30 min to generate an assembled product; and desalting and resuspension of the assembled product via magnetic bead-based nucleic acid purification involving a series of pipette transfers and mixing of magnetic beads, ethanol wash, and buffer.
As described in relation to
Additional details of the FTEP devices are illustrated in
In the FTEP devices of the disclosure, the toxicity level of the transformation results in greater than 30% viable cells after electroporation, preferably greater than 35%, 40%, 45%, 50%, 55%, 60%, 70%, 75%, 80%, 85%, 90%, 95% or even 99% viable cells following transformation, depending on the cell type and the nucleic acids being introduced into the cells.
The housing of the FTEP device can be made from many materials depending on whether the FTEP device is to be reused, autoclaved, or is disposable, including stainless steel, silicon, glass, resin, polyvinyl chloride, polyethylene, polyamide, polystyrene, polyethylene, polypropylene, acrylonitrile butadiene, polycarbonate, polyetheretheketone (PEEK), polysulfone and polyurethane, co-polymers of these and other polymers. Similarly, the walls of the channels in the device can be made of any suitable material including silicone, resin, glass, glass fiber, polyvinyl chloride, polyethylene, polyamide, polyethylene, polypropylene, acrylonitrile butadiene, polycarbonate, polyetheretheketone (PEEK), polysulfone and polyurethane, co-polymers of these and other polymers. Preferred materials include crystal styrene, cyclo-olefin polymer (COP) and cyclic olephin co-polymers (COC), which allow the device to be formed entirely by injection molding in one piece with the exception of the electrodes and, e.g., a bottom sealing film if present.
The FTEP devices described herein (or portions of the FTEP devices) can be created or fabricated via various techniques, e.g., as entire devices or by creation of structural layers that are fused or otherwise coupled. For example, for metal FTEP devices, fabrication may include precision mechanical machining or laser machining; for silicon FTEP devices, fabrication may include dry or wet etching; for glass FTEP devices, fabrication may include dry or wet etching, powderblasting, sandblasting, or photostructuring; and for plastic FTEP devices fabrication may include thermoforming, injection molding, hot embossing, or laser machining. The components of the FTEP devices may be manufactured separately and then assembled, or certain components of the FTEP devices (or even the entire FTEP device except for the electrodes) may be manufactured (e.g., using 3D printing) or molded (e.g., using injection molding) as a single entity, with other components added after molding. For example, housing and channels may be manufactured or molded as a single entity, with the electrodes later added to form the FTEP unit. Alternatively, the FTEP device may also be formed in two or more parallel layers, e.g., a layer with the horizontal channel and filter, a layer with the vertical channels, and a layer with the inlet and outlet ports, which are manufactured and/or molded individually and assembled following manufacture.
In specific aspects, the FTEP device can be manufactured using a circuit board as a base, with the electrodes, filter and/or the flow channel formed in the desired configuration on the circuit board, and the remaining housing of the device containing, e.g., the one or more inlet and outlet channels and/or the flow channel formed as a separate layer that is then sealed onto the circuit board. The sealing of the top of the housing onto the circuit board provides the desired configuration of the different elements of the FTEP devices of the disclosure. Also, two to many FTEP devices may be manufactured on a single substrate, then separated from one another thereafter or used in parallel. In certain embodiments, the FTEP devices are reusable and, in some embodiments, the FTEP devices are disposable. In additional embodiments, the FTEP devices may be autoclavable.
The electrodes 508 can be formed from any suitable metal, such as copper, stainless steel, titanium, aluminum, brass, silver, rhodium, gold or platinum, or graphite. One preferred electrode material is alloy 303 (UNS330300) austenitic stainless steel. An applied electric field can destroy electrodes made from of metals like aluminum. If a multiple-use (i.e., non-disposable) flow-through FTEP device is desired—as opposed to a disposable, one-use flow-through FTEP device—the electrode plates can be coated with metals resistant to electrochemical corrosion. Conductive coatings like noble metals, e.g., gold, can be used to protect the electrode plates.
As mentioned, the FTEP devices may comprise push-pull pneumatic means to allow multi-pass electroporation procedures; that is, cells to electroporated may be “pulled” from the inlet toward the outlet for one pass of electroporation, then be “pushed” from the outlet end of the flow-through FTEP device toward the inlet end to pass between the electrodes again for another pass of electroporation. This process may be repeated one to many times.
Depending on the type of cells to be electroporated (e.g., bacterial, yeast, mammalian) and the configuration of the electrodes, the distance between the electrodes in the flow channel can vary widely. For example, where the flow channel decreases in width, the flow channel may narrow to between 10 μm and 5 mm, or between 25 μm and 3 mm, or between 50 μm and 2 mm, or between 75 μm and 1 mm. The distance between the electrodes in the flow channel may be between 1 mm and 10 mm, or between 2 mm and 8 mm, or between 3 mm and 7 mm, or between 4 mm and 6 mm. The overall size of the FTEP device may be from 3 cm to 15 cm in length, or 4 cm to 12 cm in length, or 4.5 cm to 10 cm in length. The overall width of the FTEP device may be from 0.5 cm to 5 cm, or from 0.75 cm to 3 cm, or from 1 cm to 2.5 cm, or from 1 cm to 1.5 cm.
The region of the flow channel that is narrowed is wide enough so that at least two cells can fit in the narrowed portion side-by-side. For example, a typical bacterial cell is 1 μm in diameter; thus, the narrowed portion of the flow channel of the FTEP device used to transform such bacterial cells will be at least 2 μm wide. In another example, if a mammalian cell is approximately 50 μm in diameter, the narrowed portion of the flow channel of the FTEP device used to transform such mammalian cells will be at least 100 μm wide. That is, the narrowed portion of the FTEP device will not physically contort or “squeeze” the cells being transformed.
In embodiments of the FTEP device where reservoirs are used to introduce cells and exogenous material into the FTEP device, the reservoirs range in volume from 100 μL to 10 mL, or from 500 μL to 75 mL, or from 1 mL to 5 mL. The flow rate in the FTEP ranges from 0.1 mL to 5 mL per minute, or from 0.5 mL to 3 mL per minute, or from 1.0 mL to 2.5 mL per minute. The pressure in the FTEP device ranges from 1-30 psi, or from 2-10 psi, or from 3-5 psi.
To avoid different field intensities between the electrodes, the electrodes should be arranged in parallel. Furthermore, the surface of the electrodes should be as smooth as possible without pin holes or peaks. Electrodes having a roughness Rz of 1 to 10 μm are preferred. In another embodiment of the invention, the flow-through electroporation device comprises at least one additional electrode which applies a ground potential to the FTEP device.
After editing 6053, many cells in the colonies of cells that have been edited die as a result of the double-strand cuts caused by active editing and there is a lag in growth for the edited cells that do survive but must repair and recover following editing (microwells 6058), where cells that do not undergo editing thrive (microwells 6059) (vi). All cells are allowed to continue grow to establish colonies and normalize, where the colonies of edited cells in microwells 6058 catch up in size and/or cell number with the cells in microwells 6059 that do not undergo editing (vii). Once the cell colonies are normalized, either pooling 6060 of all cells in the microwells can take place, in which case the cells are enriched for edited cells by eliminating the bias from non-editing cells and fitness effects from editing; alternatively, colony growth in the microwells is monitored after editing, and slow growing colonies (e.g., the cells in microwells 6058) are identified and selected 6061 (e.g., “cherry picked”) resulting in even greater enrichment of edited cells.
In growing the cells, the medium used will depend, of course, on the type of cells being edited—e.g., bacterial, yeast or mammalian. For example, medium for yeast cell growth includes LB, SOC, TPD, YPG, YPAD, MEM and DMEM.
A module useful for performing the method depicted in
The SWIIN module 650 in
In this
In this embodiment of a SWIIN module, the perforated member includes through-holes to accommodate ultrasonic tabs disposed on the permeate member. Thus, in this embodiment the perforated member is fabricated from 316 stainless steel, and the perforations form the walls of microwells while a filter or membrane is used to form the bottom of the microwells. Typically, the perforations (microwells) are approximately 150 μm-200 μm in diameter, and the perforated member is approximately 125 μm deep, resulting in microwells having a volume of approximately 2.5 nl, with a total of approximately 200,000 microwells. The distance between the microwells is approximately 279 μm center-to-center. Though here the microwells have a volume of approximately 2.5 nl, the volume of the microwells may be from 1 to 25 nl, or preferably from 2 to 10 nl, and even more preferably from 2 to 4 nl. As for the filter or membrane, like the filter described previously, filters appropriate for use are solvent resistant, contamination free during filtration, and are able to retain the types and sizes of cells of interest. For example, in order to retain small cell types such as bacterial cells, pore sizes can be as low as 0.10 μm, however for other cell types (e.g., such as for mammalian cells), the pore sizes can be as high as 10.0 μm-20.0 μm or more. Indeed, the pore sizes useful in the cell concentration device/module include filters with sizes from 0.10 μm, 0.11 μm, 0.12 μm, 0.13 μm, 0.14 μm, 0.15 μm, 0.16 μm, 0.17 μm, 0.18 μm, 0.19 μm, 0.20 μm, 0.21 μm, 0.22 μm, 0.23 μm, 0.24 μm, 0.25 μm, 0.26 μm, 0.27 μm, 0.28 μm, 0.29 μm, 0.30 μm, 0.31 μm, 0.32 μm, 0.33 μm, 0.34 μm, 0.35 μm, 0.36 μm, 0.37 μm, 0.38 μm, 0.39 μm, 0.40 μm, 0.41 μm, 0.42 μm, 0.43 μm, 0.44 μm, 0.45 μm, 0.46 μm, 0.47 μm, 0.48 μm, 0.49 μm, 0.50 μm and larger. The filters may be fabricated from any suitable material including cellulose mixed ester (cellulose nitrate and acetate) (CME), polycarbonate (PC), polyvinylidene fluoride (PVDF), polyethersulfone (PES), polytetrafluoroethylene (PTFE), nylon, or glass fiber.
The cross-section configuration of the mated serpentine channel may be round, elliptical, oval, square, rectangular, trapezoidal, or irregular. If square, rectangular, or another shape with generally straight sides, the cross section may be from about 2 mm to 15 mm wide, or from 3 mm to 12 mm wide, or from 5 mm to 10 mm wide. If the cross section of the mated serpentine channel is generally round, oval or elliptical, the radius of the channel may be from about 3 mm to 20 mm in hydraulic radius, or from 5 mm to 15 mm in hydraulic radius, or from 8 mm to 12 mm in hydraulic radius.
Serpentine channels 660a and 660b can have approximately the same volume or a different volume. For example, each “side” or portion 660a, 660b of the serpentine channel may have a volume of, e.g., 2 mL, or serpentine channel 660a of permeate member 608 may have a volume of 2 mL, and the serpentine channel 660b of retentate member 604 may have a volume of, e.g., 3 mL. The volume of fluid in the serpentine channel may range from about 2 mL to about 80 mL, or about 4 mL to 60 mL, or from 5 mL to 40 mL, or from 6 mL to 20 mL (note these volumes apply to a SWIIN module comprising a, e.g., 50-500K perforation member). The volume of the reservoirs may range from 5 mL to 50 mL, or from 7 mL to 40 mL, or from 8 mL to 30 mL or from 10 mL to 20 mL, and the volumes of all reservoirs may be the same or the volumes of the reservoirs may differ (e.g., the volume of the permeate reservoirs is greater than that of the retentate reservoirs).
The serpentine channel portions 660a and 660b of the permeate member 608 and retentate member 604, respectively, are approximately 200 mm long, 130 mm wide, and 4 mm thick, though in other embodiments, the retentate and permeate members can be from 75 mm to 400 mm in length, or from 100 mm to 300 mm in length, or from 150 mm to 250 mm in length; from 50 mm to 250 mm in width, or from 75 mm to 200 mm in width, or from 100 mm to 150 mm in width; and from 2 mm to 15 mm in thickness, or from 4 mm to 10 mm in thickness, or from 5 mm to 8 mm in thickness. Embodiments the retentate (and permeate) members may be fabricated from PMMA (poly(methyl methacrylate) or other materials may be used, including polycarbonate, cyclic olefin co-polymer (COC), glass, polyvinyl chloride, polyethylene, polyamide, polypropylene, polysulfone, polyurethane, and co-polymers of these and other polymers. Preferably at least the retentate member is fabricated from a transparent material so that the cells can be visualized (see, e.g.,
Because the retentate member preferably is transparent, colony growth in the SWIIN module can be monitored by automated devices such as those sold by JoVE (ScanLag™ system, Cambridge, Mass.) (also see Levin-Reisman, et al., Nature Methods, 7:737-39 (2010)). Cell growth for, e.g., mammalian cells may be monitored by, e.g., the growth monitor sold by IncuCyte (Ann Arbor, Mich.) (see also, Choudhry, PLos One, 11(2):e0148469 (2016)). Further, automated colony pickers may be employed, such as those sold by, e.g., TECAN (Pickolo™ system, Mannedorf, Switzerland); Hudson Inc. (RapidPick™, Springfield, N.J.); Molecular Devices (QPix 400™ system, San Jose, Calif.); and Singer Instruments (PIXL™ system, Somerset, UK).
Due to the heating and cooling of the SWIIN module, condensation may accumulate on the retentate member which may interfere with accurate visualization of the growing cell colonies. Condensation of the SWIIN module 650 may be controlled by, e.g., moving heated air over the top of (e.g., retentate member) of the SWIIN module 650, or by applying a transparent heated lid over at least the serpentine channel portion 660b of the retentate member 604. See, e.g.,
In SWIIN module 650 cells and medium—at a dilution appropriate for Poisson or substantial Poisson distribution of the cells in the microwells of the perforated member—are flowed into serpentine channel 660b from ports in retentate member 604, and the cells settle in the microwells while the medium passes through the filter into serpentine channel 660a in permeate member 608. The cells are retained in the microwells of perforated member 601 as the cells cannot travel through filter 603. Appropriate medium may be introduced into permeate member 608 through permeate ports 611. The medium flows upward through filter 603 to nourish the cells in the microwells (perforations) of perforated member 601. Additionally, buffer exchange can be effected by cycling medium through the retentate and permeate members. In operation, the cells are deposited into the microwells, are grown for an initial, e.g., 2-100 doublings, editing is induced by, e.g., raising the temperature of the SWIIN to 42° C. to induce a temperature inducible promoter or by removing growth medium from the permeate member and replacing the growth medium with a medium comprising a chemical component that induces an inducible promoter.
Once editing has taken place, the temperature of the SWIIN may be decreased, or the inducing medium may be removed and replaced with fresh medium lacking the chemical component thereby de-activating the inducible promoter. The cells then continue to grow in the SWIIN module 650 until the growth of the cell colonies in the microwells is normalized. For the normalization protocol, once the colonies are normalized, the colonies are flushed from the microwells by applying fluid or air pressure (or both) to the permeate member serpentine channel 660a and thus to filter 603 and pooled. Alternatively, if cherry picking is desired, the growth of the cell colonies in the microwells is monitored, and slow-growing colonies are directly selected; or, fast-growing colonies are eliminated.
Imaging of cell colonies growing in the wells of the SWIIN is desired in most implementations for, e.g., monitoring both cell growth and device performance and imaging is necessary for cherry-picking implementations. Real-time monitoring of cell growth in the SWIIN requires backlighting, retentate plate (top plate) condensation management and a system-level approach to temperature control, air flow, and thermal management. In some implementations, imaging employs a camera or CCD device with sufficient resolution to be able to image individual wells. For example, in some configurations a camera with a 9-pixel pitch is used (that is, there are 9 pixels center-to-center for each well). Processing the images may, in some implementations, utilize reading the images in grayscale, rating each pixel from low to high, where wells with no cells will be brightest (due to full or nearly-full light transmission from the backlight) and wells with cells will be dim (due to cells blocking light transmission from the backlight). After processing the images, thresholding is performed to determine which pixels will be called “bright” or “dim”, spot finding is performed to find bright pixels and arrange them into blocks, and then the spots are arranged on a hexagonal grid of pixels that correspond to the spots. Once arranged, the measure of intensity of each well is extracted, by, e.g., looking at one or more pixels in the middle of the spot, looking at several to many pixels at random or pre-set positions, or averaging X number of pixels in the spot. In addition, background intensity may be subtracted. Thresholding is again used to call each well positive (e.g., containing cells) or negative (e.g., no cells in the well). The imaging information may be used in several ways, including taking images at time points for monitoring cell growth. Monitoring cell growth can be used to, e.g., remove the “muffin tops” of fast-growing cells followed by removal of all cells or removal of cells in “rounds” as described above, or recover cells from specific wells (e.g., slow-growing cell colonies); alternatively, wells containing fast-growing cells can be identified and areas of UV light covering the fast-growing cell colonies can be projected (or rastered with shutters) onto the SWIIN to irradiate or inhibit growth of those cells. Imaging may also be used to assure proper fluid flow in the serpentine channel 660.
In addition to the reservoir for storing the cells, the system 700 may include a reservoir for storing editing cassettes or ribozyme-containing editing cassettes 716 and a vector backbone 718. Both the editing cassettes or ribozyme-containing editing cassettes and the vector backbone are transferred from the reagent cartridge to, e.g., an electroporation device 708, which already contains the cell culture grown to a target OD and rendered electrocompetent via filtration module 730. In electroporation device 708, the assembled nucleic acids are introduced into the cells. Following electroporation, the cells are transferred into a combined recovery/selection module 710. For examples of multi-module cell editing instruments, see U.S. Pat. No. 10,253,316, issued 9 Apr. 2019; U.S. Pat. No. 10,329,559, issued 25 Jun. 2019; U.S. Pat. No. 10,323,242, issued 18 Jun. 2019; U.S. Pat. No. 10,421,959, issued 24 Sep. 2019; U.S. Pat. No. 10,465,185, issued 5 Nov. 2019; and U.S. Ser. No. 16/412,195, filed 14 May 2019; Ser. No. 16/571,091, filed 14 Sep. 2019; Ser. No. 16/666,964, filed 29 Oct. 2019; Ser. No. 16/680,643, filed 12 Nov. 2019; Ser. No. 16/750,369, filed 23 Jan. 2020, all of which are herein incorporated by reference in their entirety.
Following recovery, and, optionally, selection 710, the cells are transferred to a growth, and editing module 740. The cells are allowed to grow and editing take place. In some embodiments, editing is induced by transcription of one or both of the nuclease and the gRNA being under the control of an inducible promoter. In some embodiments, the inducible promoter is a pL promoter where the promoter is activated by a rise in temperature and “deactivated” by lowering the temperature.
The recovery, selection, growth, editing and storage modules may all be separate, may be arranged and combined as shown in
Once the cells are edited and re-grown (e.g., recovered from editing), the cells may be stored, e.g., in a storage module 712, where the cells can be kept at, e.g., 4° C. until the cells are retrieved for further study. Alternatively, the cells may be used in another round of editing. The multi-module cell processing instrument is controlled by a processor 742 configured to operate the instrument based on user input, as directed by one or more scripts, or as a combination of user input or a script. The processor 742 may control the timing, duration, temperature, and operations of the various modules of the system 700 and the dispensing of reagents. For example, the processor 742 may cool the cells post-transformation until editing is desired, upon which time the temperature may be raised to a temperature conducive of genome editing and cell growth. The processor may be programmed with standard protocol parameters from which a user may select, a user may specify one or more parameters manually or one or more scripts associated with the reagent cartridge may specify one or more operations and/or reaction parameters. In addition, the processor may notify the user (e.g., via an application to a smart phone or other device) that the cells have reached the target OD as well as update the user as to the progress of the cells in the various modules in the multi-module system.
The automated multi-module cell processing instrument 700 is a nuclease-directed genome editing system and can be used in single editing systems where, e.g., two or more edits to a cellular genome are introduced using a single editing process via multiplex editing cassettes. The system may be configured to perform sequential editing, e.g., using different nuclease-directed systems sequentially to provide two or more genome edits in a cell in each of two or more rounds of editing; and/or recursive editing, e.g. utilizing a single nuclease-directed system to introduce sequentially two or more genome edits in a cell in each of two or more round of editing.
After recovery, the cells may be transferred to a storage module 812, where the cells can be stored at, e.g., 4° C. for later processing, or the cells may be diluted and transferred to a selection/singulation/growth/induction/editing/normalization (SWIIN) module 820. In the SWIIN 820, the cells are arrayed such that there is an average of one cell per microwell. The arrayed cells may be in selection medium to select for cells that have been transformed or transfected with the editing vector(s). Once singulated, the cells grow through 2-50 doublings and establish colonies. Once colonies are established, editing is induced by providing conditions (e.g., temperature, addition of an inducing or repressing chemical) to induce editing. Editing is then initiated and allowed to proceed, the cells are allowed to grow to terminal size (e.g., normalization of the colonies) in the microwells and then are treated to conditions that cure the editing vector from this round. Once cured, the cells can be flushed out of the microwells and pooled, then transferred to the storage (or recovery) unit 812 or can be transferred back to the growth module 804 for another round of editing. In between pooling and transfer to a growth module, there typically is one or more additional steps, such as cell recovery, medium exchange (rendering the cells electrocompetent), cell concentration (typically concurrently with medium exchange by, e.g., filtration. Note that the selection/singulation/growth/induction/editing/normalization and curing modules may be the same module, where all processes are performed in, e.g., a solid wall device, or selection and/or dilution may take place in a separate vessel before the cells are transferred to the solid wall singulation/growth/induction/editing/normalization/editing module (SWIIN). Similarly, the cells may be pooled after normalization, transferred to a separate vessel, and cured in the separate vessel. As an alternative to singulation in, e.g., a solid wall device, the transformed cells may be grown in—and editing can be induced in—bulk liquid. Once the putatively-edited cells are pooled, they may be subjected to another round of editing, beginning with growth, cell concentration and treatment to render electrocompetent, and transformation by yet another donor nucleic acid in another editing cassette via the electroporation module 808.
In electroporation device 808, the yeast cells selected from the first round of editing are transformed by a second set of editing cassettes or ribozyme-containing editing cassettes and vector backbones and the cycle is repeated until the cells have been transformed and edited by a desired number of, e.g., editing cassettes. The multi-module cell processing instrument exemplified in
It should be apparent to one of ordinary skill in the art given the present disclosure that the process described may be recursive and multiplexed; that is, cells may go through the workflow described in relation to
In any recursive process, it is advantageous to “cure” the editing vectors (e.g., the gap-repaired vector backbone+the editing cassette or ribozyme-containing editing cassette). “Curing” is a process in which one or more editing vectors used in the prior round of editing is eliminated from the transformed cells. Curing can be accomplished by, e.g., cleaving the editing vector(s) using a curing plasmid thereby rendering the editing vectors nonfunctional; diluting the editing vector(s) in the cell population via cell growth (that is, the more growth cycles the cells go through, the fewer daughter cells will retain the editing vector(s)), or by, e.g., utilizing a heat-sensitive origin of replication on the editing vector. The conditions for curing will depend on the mechanism used for curing; that is, in this example, how the curing plasmid cleaves the editing vector.
The libraries are designed to increase various pathways involved in biodegradation. Amino acid production pathways are a good model for the production of such edited microbes. For example, lysine is naturally synthesized in E. coli along the diaminopimelate (DAP) biosynthetic pathway. Strain engineering strategies for increasing lysine production in E. coli and other industrially-relevant production hosts such as Corynebacterium glutamicum have historically focused on the genes in the DAP pathway as obvious targets for mutagenesis and over-expression. Beyond this short list of genes encoding the lysine biosynthetic enzymes, however, it is likely that additional loci throughout the E. coli genome may also contribute appreciably (if less directly) to improved lysine yields in an industrial production setting. For this reason, targeted mutagenesis strategies which enable a broader query of the entire genome are also of significant value to the those designing libraries for particular microbial function.
For example, nucleic acid-guided nuclease editing of 200,000 loci in the MG1655 strain of E. coli harboring an engine plasmid can result in genome-wide mutagenesis for optimization of biological activity. The engine plasmid comprises a coding sequence for the MAD7 nuclease under the control of the inducible pL promoter, the λ Red operon recombineering system under the control of the inducible pBAD promoter (inducible by the addition of arabinose in the cell growth medium), the c1857 gene under the control of a constitutive promoter, as well as a selection marker and an origin of replication. The λ Red recombineering system repairs the double-stranded breaks resulting from the cut by the MAD7 nuclease. The c1857 gene at 30° C. actively represses the pL promoter (which drives the expression of the MAD7 nuclease and the editing or CREATE cassette on the editing cassette such as the exemplary editing vector); however, at 42° C., the c1857 repressor gene unfolds or degrades, and in this state the c1857 repressor protein can no longer repress the pL promoter leading to active transcription of the coding sequence for the MAD7 nuclease and the editing (e.g., CREATE) cassette.
Mutagenesis libraries specifically targeting the genes in the DAP pathway—along with a number of genes whose enzymes convert products feeding into the DAP pathway— were designed for saturation mutagenesis. Additionally, to more deeply explore the rest of the genome for new targets involved in lysine biosynthesis, libraries were designed to target all annotated loci with either premature stop codons (for a knock-out phenotype) or insertion of a set of five synthetic promoter variants (for expression modulation phenotypes). See, e.g., U.S. Ser. No. 16/904,827, filed 18 Jun. 2020 for methods, materials and identified mutants enhancing lysine production.
The 200,000 nucleic acid mutations or edits described herein were generated using MAD7, along with a gRNA and donor DNA. A nucleic acid-guided nuclease such as MAD7 is complexed with an appropriate synthetic guide nucleic acid in a cell and can cut the genome of the cell at a desired location. The guide nucleic acid helps the nucleic acid-guided nuclease recognize and cut the DNA at a specific target sequence. By manipulating the nucleotide sequence of the guide nucleic acid, the nucleic acid-guided nuclease is programmed to target any DNA sequence for cleavage as long as an appropriate protospacer adjacent motif (PAM) is nearby. In certain aspects, the nucleic acid-guided nuclease editing system may use two separate guide nucleic acid molecules that combine to function as a guide nucleic acid, e.g., a CRISPR RNA (crRNA) and trans-activating CRISPR RNA (tracrRNA). In other aspects, the guide nucleic acid may be a single guide nucleic acid that includes both the crRNA and tracrRNA sequences.
In the methods to generate the 200,000 member library, the guide nucleic acids were provided as a sequence to be expressed from a plasmid or vector comprising both the guide sequence and the scaffold sequence as a single transcript under the control of an inducible promoter. The guide nucleic acids are engineered to target a desired target sequence by altering the guide sequence so that the guide sequence is complementary to a desired target sequence, thereby allowing hybridization between the guide sequence and the target sequence. In general, to generate an edit in the target sequence, the gRNA/nuclease complex binds to a target sequence as determined by the guide RNA, and the nuclease recognizes a protospacer adjacent motif (PAM) sequence adjacent to the target sequence. The target sequences for the genome-wide mutagenesis here encompassed 200,000 loci throughout the E. coli genome. Various types of edits were introduced herein, including site-directed mutagenesis, saturation mutagenesis, promoter swaps and ladders, knock-in and knock-out edits, SNP or short tandem repeat swaps, and start/stop codon exchanges.
The following examples are put forth so as to provide those of ordinary skill in the art with a complete disclosure and description of how to make and use the present invention, and are not intended to limit the scope of what the inventors regard as their invention, nor are they intended to represent or imply that the experiments below are all of or the only experiments performed. It will be appreciated by persons skilled in the art that numerous variations and/or modifications may be made to the invention as shown in the specific aspects without departing from the spirit or scope of the invention as broadly described. The present aspects are, therefore, to be considered in all respects as illustrative and not restrictive.
Singleplex automated genomic editing using MAD7 nuclease was successfully performed with an automated multi-module instrument as described in, e.g., U.S. Pat. No. 9,982,279; and U.S. Ser. No. 16/024,831 filed 30 Jun. 2018; Ser. No. 16/024,816 filed 30 Jun. 2018; Ser. No. 16/147,353 filed 28 Sep. 2018; Ser. No. 16/147,865 filed 30 Sep. 2018; and Ser. No. 16/147,871 filed 30 Jun. 2018.
An ampR plasmid backbone and a lacZ_F172* editing cassette were assembled via Gibson Assembly® into an “editing vector” in an isothermal nucleic acid assembly module included in the automated instrument. lacZ_F172 functionally knocks out the lacZ gene. “lacZ_F172*” indicates that the edit happens at the 172nd residue in the lacZ amino acid sequence. Following assembly, the product was de-salted in the isothermal nucleic acid assembly module using AMPure beads, washed with 80% ethanol, and eluted in buffer. The assembled editing vector and recombineering-ready, electrocompetent cells were transferred into a transformation module for electroporation. The cells and nucleic acids were combined and allowed to mix for 1 minute, and electroporation was performed for 30 seconds. The parameters for the poring pulse were: voltage, 2400 V; length, 5 ms; interval, 50 ms; number of pulses, 1; polarity, +. The parameters for the transfer pulses were: Voltage, 150 V; length, 50 ms; interval, 50 ms; number of pulses, 20; polarity, +/−. Following electroporation, the cells were transferred to a recovery module (another growth module), and allowed to recover in SOC medium containing chloramphenicol. Carbenicillin was added to the medium after 1 hour, and the cells were allowed to recover for another 2 hours. After recovery, the cells were held at 4° C. until recovered by the user.
After the automated process and recovery, an aliquot of cells was plated on MacConkey agar base supplemented with lactose (as the sugar substrate), chloramphenicol and carbenicillin and grown until colonies appeared. White colonies represented functionally edited cells, purple colonies represented un-edited cells. All liquid transfers were performed by the automated liquid handling device of the automated multi-module cell processing instrument.
The result of the automated processing was that approximately 1.0E−03 fraction of cells were transformed (comparable to conventional benchtop results), and the editing efficiency was 83.5%. The lacZ_172 edit in the white colonies was confirmed by sequencing of the edited region of the genome of the cells. Further, steps of the automated cell processing were observed remotely by webcam and text messages were sent to update the status of the automated processing procedure.
Recursive editing was successfully achieved using the automated multi-module cell processing system. An ampR plasmid backbone and a lacZ_V10* editing cassette were assembled via Gibson Assembly® into an “editing vector” in an isothermal nucleic acid assembly module included in the automated system. Similar to the lacZ_F172 edit, the lacZ_V10 edit functionally knocks out the lacZ gene. “lacZ_V10” indicates that the edit happens at amino acid position 10 in the lacZ amino acid sequence. Following assembly, the product was de-salted in the isothermal nucleic acid assembly module using AMPure beads, washed with 80% ethanol, and eluted in buffer. The first assembled editing vector and the recombineering-ready electrocompetent E. Coli cells were transferred into a transformation module for electroporation. The cells and nucleic acids were combined and allowed to mix for 1 minute, and electroporation was performed for 30 seconds. The parameters for the poring pulse were: voltage, 2400 V; length, 5 ms; interval, 50 ms; number of pulses, 1; polarity, +. The parameters for the transfer pulses were: Voltage, 150 V; length, 50 ms; interval, 50 ms; number of pulses, 20; polarity, +/−. Following electroporation, the cells were transferred to a recovery module (another growth module) allowed to recover in SOC medium containing chloramphenicol. Carbenicillin was added to the medium after 1 hour, and the cells were grown for another 2 hours. The cells were then transferred to a centrifuge module and a media exchange was then performed. Cells were resuspended in TB containing chloramphenicol and carbenicillin where the cells were grown to OD600 of 2.7, then concentrated and rendered electrocompetent.
During cell growth, a second editing vector was prepared in an isothermal nucleic acid assembly module. The second editing vector comprised a kanamycin resistance gene, and the editing cassette comprised a galK Y145* edit. If successful, the galK Y145* edit confers on the cells the ability to uptake and metabolize galactose. The edit generated by the galK Y154* cassette introduces a stop codon at the 154th amino acid reside, changing the tyrosine amino acid to a stop codon. This edit makes the galK gene product nonfunctional and inhibits the cells from being able to metabolize galactose. Following assembly, the second editing vector product was de-salted in the isothermal nucleic acid assembly module using AMPure beads, washed with 80% ethanol, and eluted in buffer. The assembled second editing vector and the electrocompetent cells (that were transformed with and selected for the first editing vector) were transferred into a transformation module for electroporation, using the same parameters as detailed above. Following electroporation, the cells were transferred to a recovery module (another growth module), allowed to recover in SOC medium containing carbenicillin. After recovery, the cells were held at 4° C. until retrieved, after which an aliquot of cells were plated on LB agar supplemented with chloramphenicol, and kanamycin. To quantify both lacZ and galK edits, replica patch plates were generated on two media types: 1) MacConkey agar base supplemented with lactose (as the sugar substrate), chloramphenicol, and kanamycin, and 2) MacConkey agar base supplemented with galactose (as the sugar substrate), chloramphenicol, and kanamycin. All liquid transfers were performed by the automated liquid handling device of the automated multi-module cell processing system.
In this recursive editing experiment, 41% of the colonies screened had both the lacZ and galK edits, the results of which were comparable to the double editing efficiencies obtained using a “benchtop” or manual approach.
5 nM oligonucleotides synthesized on a chip were amplified using Q5 polymerase in 50 μL volumes. The PCR conditions were 95° C. for 1 minute; 8 rounds of 95° C. for 30 seconds/60° C. for 30 seconds/72° C. for 2.5 minutes; with a final hold at 72° C. for 5 minutes. Following amplification, the PCR products were subjected to SPRI cleanup, where 30 μL SPRI mix was added to the 50 μL PCR reactions and incubated for 2 minutes. The tubes were subjected to a magnetic field for 2 minutes, the liquid was removed, and the beads were washed 2x with 80% ethanol, allowing 1 minute between washes. After the final wash, the beads were allowed to dry for 2 minutes, 50 μL 0.5x TE pH 8.0 was added to the tubes, and the beads were vortexed to mix. The slurry was incubated at room temperature for 2 minutes, then subjected to the magnetic field for 2 minutes. The eluate was removed and the DNA quantified.
Following quantification, a second amplification procedure was carried out using a dilution of the eluate from the SPRI cleanup. PCR was performed under the following conditions: 95° C. for 1 minute; 18 rounds of 95° C. for 30 seconds/72° C. for 2.5 minutes; with a final hold at 72° C. for 5 minutes. Amplicons were checked on a 2% agarose gel and pools with the cleanest output(s) were identified. Amplification products appearing to have heterodimers or chimeras were not used.
Purified backbone vector was linearized by restriction enzyme digest with StuI. Up to 20 μg of purified backbone vector was in a 100 μL total volume in StuI-supplied buffer. Digestion was carried out at 30° C. for 16 hrs. Linear backbone was dialyzed to remove salt on 0.025 μm MCE membrane for ˜60 min on nuclease-free water. Linear backbone concentration was measured using dye/fluorometer-based quantification.
The afternoon before transformation was to occur, 10 mL of YPAD was added to S. cerevisiae cells, and the culture was shaken at 250 rpm at 30° C. overnight. The next day, approximately 2 mL of the overnight culture was added to 100 mL of fresh YPAD in a 250-mL baffled flask and grown until the OD600 reading reached 0.3+/−0.05. The culture was then placed in a 30° C. incubator shaking at 250 rpm and allowed to grow for 4-5 hours, with the OD checked every hour. When the culture reached ˜1.5 OD600, two 50 mL aliquots of the culture were poured into two 50-mL conical vials and centrifuged at 4300 rpm for 2 minutes at room temperature. The supernatant was removed from the 50 mL conical tubes, avoiding disturbing the cell pellet. 25 mL of lithium acetate/DTT solution was added to each conical tube and the pellet was gently resuspended using an inoculating loop, needle, or long toothpick.
Following resuspension, both cell suspensions were transferred to a 250-mL flask and placed in the shaker to shake at 30° C. and 200 rpm for 30 minutes. After incubation was complete, the suspension was transferred to one 50-mL conical tube and centrifuged at 4300 RPM for 3 minutes. The supernatant was then discarded. From this point on, cold liquids were used and kept on ice until electroporation was complete. 50 mL of 1 M sorbitol was added to the cells and the pellet was resuspended. The cells were centrifuged at 4300 rpm for 3 minutes at 4° C., and the supernatant was discarded. The centrifugation and resuspension steps were repeated for a total of three washes. 50 μL of 1 M sorbitol was then added to one pellet, the cells were resuspended, then this aliquot of cells was transferred to the other tube and the second pellet was resuspended. The approximate volume of the cell suspension was measured, then brought to a 1 mL volume with cold 1 M sorbitol. The cell/sorbitol mixture and transferred into a 2-mm cuvette. Impedance measurement of the cells was measured in the cuvette. At this point the KW must be >20. If this is not the case the cells should be washed in cold sorbitol two to three additional times.
Transformation was then performed using 500 ng of linear backbone along with 50 ng editing cassettes with the competent S. cerevisiae cells. 2 mm electroporation cuvettes were placed on ice and the plasmid/cassette mix was added to each corresponding cuvette. 100 μL of electrocompetent cells were added to each cuvette and the linear backbone and cassettes. Each sample was electroporated using the following conditions on a NEPAGENE electroporator: Poring pulse: 1800V, 5.0 second pulse length, 50.0 msec pulse interval, 1 pulse; Transfer pulse: 100 V, 50.0 msec pulse length, 50.0 msec pulse interval, with 3 pulses. Once the transformation process is complete, 900 μL of room temperature YPAD Sorbitol media was added to each cuvette. The cells were then transferred and suspended in a 15 mL tube and incubated shaking at 250 RPM at 30° C. for 3 hours. 9 mL of YPAD and 10 μL of Hygromycin B 1000× stock was added to the 15 mL tube.
The edited libraries can be used to identify microbes that are optimized for production of tyrosine. Additional guidance can be found in U.S. Ser. No. 62/907,402 filed September 27 and 62/912,544, which are hereby incorporated by reference for all purposes.
Library stocks of edited S. cerevisiae were diluted and plated onto 245×245 mm YPD agar plates (Teknova) containing 250 μg/mL Hygromycin (Teknova) using sterile glass beads. Libraries were diluted an appropriate amount to yield ˜1500-2000 colonies on the plates. Plates were incubated 36-48 h at 30° C. and then stored at 4° C. until use. Colonies were picked using a QPix™ 420 (Molecular Devices) and deposited into sterile 1.2 mL square 96-well plates (Thomas Scientific) containing 300 μL of growth medium (Synthetic Complete (SC) medium (-Tyr, -Phe) containing 250 μg/mL Hygromycin (Sigma)). SC (-Tyr, -Phe) medium was prepared by adding 6.71 g/L YNB+nitrogen (AmSO4) (Sunrise Science Products), 0.69 g/L CSM (-Tyr, -Phe) (Sunrise Science Products) and 20 g/L glucose (Tenova) to water. Plates were sealed (AirPore sheets (Qiagen)) and incubated for 36-48 h in a shaker incubator (Climo-Shaker ISF1-X (Kuhner), 30° C., 85% humidity, 250 rpm). Plate cultures were then diluted 20-fold (15 μL culture into 285 μL medium) into new 96-well plates containing fresh SC (-Tyr, -Phe) medium. Production plates were incubated for 24 h in a shaker incubator (Climo-Shaker ISF1-X (Kuhner), 30° C., 85% humidity, 250 rpm).
Production plates were centrifuged (Centrifuge 5920R, Eppendorf) at 3,000 g for 10 min to pellet cells. The supernatants from production plates were diluted 500-fold into water (10 μL of supernatant with 490 μL) of water in 1.2 mL square 96-well plates. Samples were thoroughly mixed and then diluted a subsequent 10-fold further into acetonitrile (LC/MS grade, Fisher), (20 μL sample with 180 μL of acetonitrile) into a 96-well Plate (polypropylene, 335 μL/well, Conical Bottom (Thomas Scientific). Plates were heat sealed and thoroughly mixed.
Tyrosine concentrations were determined using a RapidFire high-throughput mass spectrometry system (Agilent) coupled to a 6470 Triple Quad mass spectrometer (Agilent). The RapidFire conditions were as follows: Pump 1: 95% acetonitrile (LC/MS grade, Fisher), 5% water (LC/MS grade, Fisher), 0.1% formic acid (Sigma) 1.5 mL/min, Pump 2: 100% water, 1.25 mL/min, Pump 3: 5% acetonitrile, 95% water, 1.25 mL/min. RapidFire method: Aspirate: 600 ms, Load/wash: 3000 ms, Extra wash: 0 ms, Elute: 3000 ms, Re-equilibrate: 500 ms 10 μL injection loop. The following mass spectrometry conditions were used for tyrosine detection.
Precursor ion: 182.1 m/z, Product ion (quantifying): 136.0 m/z, Dwell: 50, Fragmentor: 72, Collision energy: 12, Cell accelerator voltage: 4, Polarity: positive
Precursor ion: 182.1 m/z, Product ion (qualifying): 91.0 m/z, Dwell: 50, Fragmentor: 72, Collision energy: 36, Cell accelerator voltage: 4, Polarity: positive
Source conditions: Gas Temp: 300° C., Gas Flow: 10 L/min, Nebulizer: 45 psi, Sheath gas temp: 350° C., Sheath gas flow: 11 L/min, Capillary voltage: 3500V (positive), Nozzle voltage: 500V (positive)
Data was analyzed using MassHunter Quantitative Analysis software (Agilent) with a standard curve of Tyrosine used for quantitation of Tyrosine in the samples. Each 96-well plate of samples contained 4 replicates of the wildtype strain to calculate the relative Tyrosine of samples compared to the base strain used for editing. Hits from the primary screen were re-tested in quadruplicate using a similar protocol as described above. For methods and materials as well as identified S. cerevisiae mutants with enhanced tyrosine production, see U.S. Ser. No. 16/026,359, filed 21 Sep. 2020.
Library stocks of edited S. cerevisiae were diluted and plated onto 245×245 mm LB agar plates (Teknova) containing 100 μg/mL carbenicillin (Teknova) and 25 μg/mL chloramphenicol (Teknova) using sterile glass beads. Libraries were diluted an appropriate amount to yield ˜2000-3000 colonies on the plates. Plates were incubated ˜16 h at 30° C. and then stored at 4° C. until use. Colonies were picked using a QPix™ 420 (Molecular Devices) and deposited into sterile 1.2 mL square 96-well plates (Thomas Scientific) containing 300 μL of overnight growth medium (EZ Rich Defined Medium, w/o lysine (Teknova), 100 μg/mL carbenicillin and 25 μg/mL chloramphenicol). Plates were sealed (AirPore sheets (Qiagen)) and incubated for ˜19 h in a shaker incubator (Climo-Shaker ISF1-X (Kuhner), 30° C., 85% humidity, 250 rpm). Plate cultures were then diluted 20-fold (15 μL culture into 285 μL medium) into new 96-well plates containing lysine production medium (20 g/L ammonium sulfate (Teknova), 200 mM MOPS buffer (Teknova), 3 mg/L Iron(II) sulfate heptahydrate (Sigma), 3 mg/L Manganese (II) sulfate monohydrate (Sigma), 0.5 mg/L Biotin (Sigma), 1 mg/L Thiamine hydrochloride (Sigma), 0.7 g/L Potassium chloride (Teknova), 20 g/L glucose (Teknova), 5 g/L Potassium phosphate monobasic (Sigma), 1 mL/L Trace metal mixture (Teknova), 1 mM Magnesium sulfate (Teknova), 100 μg/mL carbenicillin and 25 μg/mL chloramphenicol). Production plates were incubated for 24 h in a shaker incubator (Climo-Shaker ISF1-X (Kuhner), 30° C., 85% humidity, 250 rpm).
Production plates were centrifuged (Centrifuge 5920R, Eppendorf) at 3,000 g for 10 min to pellet cells. The supernatants from production plates were diluted 100-fold into water (5 μL of supernatant with 495 μL) of water in 1.2 mL square 96-well plates. Samples were thoroughly mixed and then diluted a subsequent 10-fold further into a 50:50 mixture of acetonitrile and water (20 μL sample with 180 μL of the acetonitrile/water mixture) into a 96-well Plate (polypropylene, 335 μL/well, Conical Bottom (Thomas Scientific). Plates were heat sealed and thoroughly mixed.
Lysine concentrations were determined using a RapidFire high-throughput mass spectrometry system (Agilent) coupled to a 6470 Triple Quad mass spectrometer (Agilent). The RapidFire conditions were as follows: Pump 1: 80% acetonitrile (LC/MS grade, Fisher), 20% water (LC/MS grade, Fisher), 1.5 mL/min, Pump 2: 100% water, 1.25 mL/min, Pump 3: 5% acetonitrile, 95% water, 1.25 mL/min. RapidFire method: Aspirate: 600 ms, Load/wash: 2000 ms, Extra wash: 0 ms, Elute: 3000 ms, Re-equilibrate: 500 ms. 10 μL injection loop. The following mass spectrometry conditions were used for lysine detection.
Precursor ion: 147.1 m/z, Product ion (quantifying): 84 m/z, Dwell: 20, Fragmentor: 80, Collision energy: 20, Cell accelerator voltage: 4, Polarity: positive Precursor ion: 147.1 m/z, Product ion (qualifying): 130 m/z, Dwell: 20, Fragmentor: 80, Collision energy: 8, Cell accelerator voltage: 4, Polarity: positive Source conditions: Gas Temp: 300° C., Gas Flow: 10 L/min, Nebulizer: 45 psi, Sheath gas temp: 350° C., Sheath gas flow: 11 L/min, Capillary voltage: 3000V (positive), Nozzle voltage: 1500V (positive)
Data was analyzed using MassHunter Quantitative Analysis software (Agilent) with a standard curve of lysine used for quantitation of lysine in the samples. Each 96-well plate of samples contained 4 replicates of the wildtype strain and 4 replicates of the dapA E84T positive control strain to calculate the relative lysine yield of samples compared to the controls. Hits from the primary screen were re-tested in quadruplicate using a similar protocol as described above.
The afternoon before transformation was to occur, 10 mL of YPAD was added to Bacillus sp. YP1 cells, and the culture was shaken at 250 rpm at 30° C. overnight. The next day, approximately 2 mL of the overnight culture was added to 100 mL of fresh YPAD in a 250-mL baffled flask and grown until the OD600 reading reached 0.3+/−0.05. The culture was then placed in a 30° C. incubator shaking at 250 rpm and allowed to grow for 4-5 hours, with the OD checked every hour. When the culture reached ˜1.5 OD600, two 50 mL aliquots of the culture were poured into two 50-mL conical vials and centrifuged at 4300 rpm for 2 minutes at room temperature. The supernatant was removed from the 50 mL conical tubes, avoiding disturbing the cell pellet. 25 mL of lithium acetate/DTT solution was added to each conical tube and the pellet was gently resuspended using an inoculating loop, needle, or long toothpick.
Following resuspension, both cell suspensions were transferred to a 250-mL flask and placed in the shaker to shake at 30° C. and 200 rpm for 30 minutes. After incubation was complete, the suspension was transferred to one 50-mL conical tube and centrifuged at 4300 RPM for 3 minutes. The supernatant was then discarded. From this point on, cold liquids were used and kept on ice until electroporation was complete. 50 mL of 1 M sorbitol was added to the cells and the pellet was resuspended. The cells were centrifuged at 4300 rpm for 3 minutes at 4° C., and the supernatant was discarded. The centrifugation and resuspension steps were repeated for a total of three washes. 50 μL of 1 M sorbitol was then added to one pellet, the cells were resuspended, then this aliquot of cells was transferred to the other tube and the second pellet was resuspended. The approximate volume of the cell suspension was measured, then brought to a 1 mL volume with cold 1 M sorbitol. The cell/sorbitol mixture and transferred into a 2-mm cuvette. Impedance measurement of the cells was measured in the cuvette. At this point the KW must be >20. If this is not the case the cells should be washed in cold sorbitol two to three additional times.
Transformation was then performed using 500 ng of linear backbone along with 50 ng editing cassettes with the competent Bacillus sp. cells. 2 mm electroporation cuvettes were placed on ice and the plasmid/cassette mix was added to each corresponding cuvette. 100 μL of electrocompetent cells were added to each cuvette and the linear backbone and cassettes. Each sample was electroporated using the following conditions on a NEPAGENE electroporator: Poring pulse: 1800V, 5.0 second pulse length, 50.0 msec pulse interval, 1 pulse; Transfer pulse: 100 V, 50.0 msec pulse length, 50.0 msec pulse interval, with 3 pulses. Once the transformation process is complete, 900 μL of room temperature YPAD Sorbitol media was added to each cuvette. The cells were then transferred and suspended in a 15 mL tube and incubated shaking at 250 RPM at 30° C. for 3 hours. 9 mL of YPAD and 10 μL of Hygromycin B 1000× stock was added to the 15 mL tube.
Library stocks of edited Bacillus sp. YP1 are diluted and plated onto 245×245 mm YPD agar plates (Teknova) using sterile glass beads. Libraries are diluted an appropriate amount to yield ˜1500-2000 colonies on the plates. Plates are incubated 36-48 h at 30° C. and then stored at 4° C. until use. Colonies are picked using a QPix™ 420 (Molecular Devices) and deposited into sterile 1.2 mL square 96-well plates (Thomas Scientific) containing 300 μL of growth medium (Nonselective media for Bacillus (Millipore Sigma)). Plates are sealed (AirPore sheets (Qiagen)) and incubated for 36-48 h in a shaker incubator (Climo-Shaker ISF1-X (Kuhner), 30° C., 85% humidity, 250 rpm). Plate cultures are then diluted 20-fold (15 μL culture into 285 μL medium) into new 96-well plates containing fresh growth medium. Production plates were incubated for 24 h in a shaker incubator (Climo-Shaker ISF1-X (Kuhner), 30° C., 85% humidity, 250 rpm).
Production plates are centrifuged (Centrifuge 5920R, Eppendorf) at 3,000 g for 10 min to pellet cells. The supernatants from production plates were diluted 500-fold into water (10 μL of supernatant with 490 μL) of water in 1.2 mL square 96-well plates. Samples are thoroughly mixed and diluted a subsequent 10-fold further into acetonitrile (LC/MS grade, Fisher), (20 μL sample with 180 μL of acetonitrile) into a 96-well Plate (polypropylene, 335 μL/well, Conical Bottom (Thomas Scientific). Plates are heat sealed and thoroughly mixed.
PE biodegradation is measured using a few approaches to detect biodegradation. Numerous detection methods can be used, e.g., as set forth in Montazer Z et al. Polymers, 12, 123-147 (2020).
One detection method is measurement of gravimetric weight loss of the plastic using a sufficiently sensitive scale. See, e.g., Harshvardhan, K. and Jha, B. Mar. Pollut. Bull. 77, 100-106 (2013); Usha R. et al., Libyan Agric. Res. Cent. J. Intern. 2, 200-204 (2011); To accurately measure the dry mass of residual polymers after biodegradation, the PE material is removed from any media and PE with bound Bacillus cells filtered using a suitable filter paper. The filtered PE particles are washed with 2% (m/v) sodium dodecyl sulfate to lyse any remaining cells that attached to the PE surface. The PE particles remaining on the filter paper are further rinsed with distilled water and dried overnight at 60° C. before weighing. The rate of biodegradation is the percentage of polymer weights loss per unit time of experiment. See Kyaw, B. M. et al., Indian J. Microbiol. 52, 411-419 (2012); Montazer, Z. et al., Can. J. Microbiol., 65, 1-11 (2019);
Another such detection method is the ability of the edited Bacillus sp. colonies to colonize and form biofilms on PE and secrete biosurfactants. Biofilm formation is assessed using biofilm quantification by direct counting of the number of cells adhering to the surface of the PE film and/or protein concentration, as described in Das, M. P. and Kumar, S., 3 Biotech, 5, 81-86 (2015). The PE film is provided as the sole carbon source for the edited Bacillus sp. of the library. In addition, secretion of biosurfactant molecules by the Bacillus sp., such as the enzyme surfactin, is measured to assess the ability to degrade the PE rather than just biofilm formation (Vimala, P. P. and Mathew, L. Procedia Technol., 24, 232-239 (2016)).
PE biodegradation is also measured by Fourier transform infrared (FT-IR) spectrometry, which detects oxidative products of physical plastic degradation such as the appearance of carbonyl-groups, or scanning electron microscopy, which measures physical deterioration of the polymer surface following colonization of the PE film by the edited Bacillus. See, e.g., Harshvardhan, K. and Jha, B., supra.
While this invention is satisfied by embodiments in many different forms, as described in detail in connection with preferred embodiments of the invention, it is understood that the present disclosure is to be considered as exemplary of the principles of the invention and is not intended to limit the invention to the specific embodiments illustrated and described herein. Numerous variations may be made by persons skilled in the art without departure from the spirit of the invention. The scope of the invention will be measured by the appended claims and their equivalents. The abstract and the title are snot to be construed as limiting the scope of the present invention, as their purpose is to enable the appropriate authorities, as well as the general public, to quickly determine the general nature of the invention. In the claims that follow, unless the term “means” is used, none of the features or elements recited therein should be construed as means-plus-function limitations pursuant to 35 U.S.C. § 112, ¶6.
